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Appl Environ Microbiol. Apr 2009; 75(7): 1876–1884.
Published online Feb 5, 2009. doi:  10.1128/AEM.01042-08
PMCID: PMC2663209

Transcription Analysis of Genes Encoding Homologues of Reductive Dehalogenases in “Dehalococcoides” sp. Strain CBDB1 by Using Terminal Restriction Fragment Length Polymorphism and Quantitative PCR[down-pointing small open triangle]

Abstract

The transcription of reductive dehalogenase homologous (rdh) genes of “Dehalococcoides” sp. strain CBDB1 was investigated during the growth and reductive dechlorination of 1,2,3- and 1,2,4-trichlorobenzene (TCB). A method was developed to monitor the expression of all 32 rdhA genes present in the genome based on reverse transcription-PCR amplification with 13 degenerate primer pairs and terminal restriction fragment length polymorphism (t-RFLP) analysis. With this approach, the upregulation of the transcription of 29 rdhA genes was indicated in response to 1,2,3- and 1,2,4-TCB added after a substrate depletion period of 72 h. The transcription of the remaining three rdhA genes additionally was detected using specific primers. While most rdhA genes were upregulated similarly in cultures after induction with 1,2,3-TCB or 1,2,4-TCB, three rdhA genes responded differentially to 1,2,3- and 1,2,4-TCB, as revealed by the comparison of t-RFLP profiles. The enhanced transcription of cbdbA1453 and cbdbA187 was observed in the presence of 1,2,3-TCB, while the transcription of cbdbA1624 was strongly induced by 1,2,4-TCB. Comparison of t-RFLP profiles obtained from cDNA and genomic DNA indicated a particularly high induction of the transcription of cbrA (=cbdbA84) by both TCBs. As indicated by reverse transcription-quantitative PCR, the transcription of these plus two other rdhA genes (cbdbA1588 and cbdbA1618) increased within 48 to 72 h by one or two orders of magnitude. Subsequently, transcript levels slowly decreased and approached initial transcript levels several days after complete dehalogenation. Finally, cbrA was transcribed to a level of 22 transcripts per cbrA gene, suggesting that cbrA mRNA could be an appropriate biomarker for the investigation of the natural dechlorination potential at chlorobenzene-contaminated sites.

Highly chlorinated aromatic compounds, like polychlorinated benzenes, dibenzo-p-dioxins and -furans, and polychlorinated biphenyls (PCB) persist in the environment for long periods, tend to accumulate in food chains, and are ubiquitously distributed. Many freshwater and marine sediments worldwide have been polluted by high concentrations of these compounds (4, 7, 43). Under reducing conditions often prevailing in sediments, reductive dehalogenation is considered the main biological process for the natural attenuation of highly chlorinated compounds. Several anaerobic bacteria are known to grow by respiratory dehalogenation with chlorinated compounds (for a review, see reference 40). Physiological studies with pure cultures of “Dehalococcoides” spp. and related bacteria of subphylum II of the Chloroflexi have shown that they are highly specialized in using reductive dechlorination as a respiratory process (3, 12, 13, 30, 41). The genome sequences of “Dehalococcoides ethenogenes” strain 195 and Dehalococcoides sp. strain CBDB1 confirmed their metabolic restriction to respiratory dehalogenation (24, 38) and revealed the presence of 17 and 32 reductive dehalogenase homologous (rdh) genes, respectively, some of which are thought to be involved in the dechlorination of a broad spectrum of halogenated compounds. The rdhA genes code for the catalytic subunit of the terminal electron-transferring enzyme, an iron-sulfur corrinoid protein (14), and contain a twin arginine translocation (TAT) signal sequence that is responsible for transport to or across the cytoplasmic membrane. The rdhA genes generally are colocalized with rdhB genes that encode potential membrane-anchoring proteins.

The functions of only a few RdhA proteins of Dehalococcoides species have been elucidated by biochemical studies. The trichloroethene-reductive dehalogenase (TceA) of D. ethenogenes strain 195 catalyzes the reductive dechlorination of trichloroethene to ethene and of several other haloalkanes and haloalkenes. Membrane localization studies indicated that TceA is located on the exterior of the cytoplasmic membrane (28), as was also suggested for the chlorobenzene-reductive dehalogenase of strain CBDB1 (15). An additional enzyme is necessary in D. ethenogenes strain 195 for the complete dechlorination of tetrachloroethene (PCE) to ethene, the PCE-reductive dehalogenase (PceA) (29). Recent proteomic studies demonstrated the high abundance of both PceA and TceA during growth on PCE (11, 31). A reductive dehalogenase was partially purified from Dehalococcoides sp. strain VS and exhibited the highest activity with dichloroethenes and vinyl chloride (33). Strain BAV1 contains a different enzyme, BvcA, but it has a similar function, as indicated by the transcription analysis of cells grown with vinyl chloride (23).

The first chlorobenzene-reductive dehalogenase, CbrA, recently was identified and purified from Dehalococcoides sp. strain CBDB1 (2); it has no orthologues in other known genome sequences, including that of D. ethenogenes strain 195. CbrA catalyzes the dechlorination of 1,2,3,4-tetrachlorobenzene and 1,2,3-trichlorobenzene (1,2,3-TCB). Strain CBDB1 possesses a very high capability to reductively dechlorinate different classes of chlorinated aromatic compounds. Hexachlorobenzene, pentachlorobenzene, all three isomers of tetrachlorobenzene, 1,2,3- and 1,2,4-TCB (3, 18), five chlorinated dibenzo-p-dioxins (6), pentachlorophenol, all three tetrachlorophenols, all six trichlorophenols, and three dichlorophenols (1) were reductively dechlorinated at positions flanked by one or two chlorine atoms, leading to branched dechlorination pathways. The PCB-dechlorinating strain DF-1, which is distantly related to Dehalococcoides, also is able to dechlorinate chlorinated benzenes but is restricted to the cleavage of carbon-chlorine bonds at positions flanked by two chlorines (46). D. ethenogenes strain 195 dechlorinates a less diverse range of chlorinated benzenes and chlorinated dibenzo-p-dioxins than strain CBDB1 (10), and, interestingly, can only cometabolically catalyze the dechlorination of 1,2,3- and 1,2,4-TCB during growth on PCE.

The elucidation of the function of the multiple rdhA genes of Dehalococcoides sp. strain CBDB1 will aid our understanding of how the organism catalyzes the dehalogenation of this remarkably wide range of different chloroaromatics. Since proteomic studies still are hampered by the need for relatively large amounts of protein, which are difficult to obtain from bacteria cultivated with poorly soluble chloroaromatics, we have conducted transcription analyses targeting the whole set of 32 rdhA genes encoded in strain CBDB1. Using a terminal restriction fragment length polymorphism (t-RFLP) approach for the differentiation of individual rdhA transcripts and quantitative PCR (qPCR) for quantification, the transcriptional response of Dehalococcoides sp. strain CBDB1 to two chlorinated benzenes, 1,2,3- and 1,2,4-TCB, is presented.

MATERIALS AND METHODS

Bacterial strains and cultivation conditions.

Strain CBDB1 was grown in Ti(III)-citrate-reduced, carbonate-buffered synthetic medium with hydrogen as the electron donor and 5 mM acetate as the carbon source, as previously described (3). Following the inoculation of the anaerobic medium, the headspace was pressurized with 20% CO2 and 80% N2 (1.3 bar), hydrogen was added to a pressure of 1.5 bar, and cultivation was performed at a temperature of 30°C with shaking (120 rpm). The electron acceptors 1,2,3-TCB and 1,2,4-TCB were added from a 5.5 M stock solution in acetone at a final concentration of 60 μM. Two-liquid phase (TLP) cultures (50 ml) were supplied with 1,2,3- or 1,2,4-TCB dissolved in N2-gassed hexadecane (200 mM), resulting in a nominal concentration of 10 mM. Escherichia coli XL1-Blue MRF′ (Stratagene, Amsterdam, The Netherlands), used as the host for cloning vectors, was cultivated on a rotary shaker at 140 rpm in nutrient broth or statically on nutrient agar at 37°C with final concentrations of ampicillin, 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside, and isopropyl-β-d-thiogalactopyranoside of 125 μg ml−1, 80 μg ml−1, and 0.5 mM, respectively.

Chemicals.

Chlorobenzenes were purchased from Sigma-Aldrich (Seelze, Germany) at a purity of 99%.

Analytical techniques.

Chlorobenzenes were extracted from 0.5-ml samples with 1 ml of hexane and analyzed using a Shimadzu GC 14A equipped with a flame ionization detector and a DB608-megabore-capillary column (30 m; inside diameter, 0.331 mm; 0.5-μm film thickness; J&W Scientific, Folsom, CA). Helium 5.0 served as the carrier gas (flow rate, 0.3 m s−1; column pressure, 2.4 × 105 Pa) and nitrogen 5.0 as the make-up gas. The following temperature program was used: initial hold at 40°C for 1.1 min; increase at a rate of 40°C min−1 to 70°C (hold for 1.9 min); 20°C min−1 to 140°C (hold for 1.2 min); 40°C min−1 to 160°C (hold for 1.2 min); 25°C min−1 to 220°C (hold for 0.5 min); and 40°C min−1 to 280°C, with a final hold for 5 min. Injector and detector temperatures were 250 and 280°C, respectively. A 10-level calibration curve (0.7 to 364 μM) was generated with authentic chlorobenzene standards and 1,3,5-tribromobenzene as an internal standard.

Transcription experiment.

TLP cultures supplemented with 1,2,3- or 1,2,4-TCB served as precultures to grow strain CBDB1 to cell numbers of 108 to 109 cells ml−1 based on the qPCR measurements of 16S rRNA gene copy numbers (see below). For a single transcription experiment, five replicate 50-ml cultures were inoculated with 10% [vol/vol] cells from one preculture. To reduce the level of dehalogenase gene transcripts that originated from precultivation, inoculated cultures were incubated without a chlorinated electron acceptor for 72 h. To start the transcription analysis, either 1,2,3- or 1,2,4-TCB was added from an acetone stock solution, and samples were taken immediately and after 24, 48, 72, and 168 h. Eight transcription experiments were performed, four with 1,2,3-TCB and four with 1,2,4-TCB as the electron acceptor. Half of each type of culture was inoculated with half the cells pregrown with 1,2,3-TCB and the other half pregrown with 1,2,4-TCB, thus representing four slightly different experimental conditions. For clarity, the results of only one replicate of each of the four variants are presented, but the same general observations were made in the parallel setups. At each sampling point, the concentration of formed dichlorobenzenes (DCBs) was measured in all replicate cultures, and one culture was completely harvested for nucleic acid extraction. Cultures supplemented with the respective amount of acetone served as controls.

Extraction of nucleic acids.

DNA was extracted from 1 ml of liquid culture using bead beating and ethanol precipitation as described previously (9). For RNA extraction, 45 ml of the cultures was harvested by centrifugation at 3,500 × g for 45 min at 4°C. The supernatant was discarded, except for a 1-ml residue. The pellet was resuspended in this volume, transferred to a 1.5-ml tube, and centrifuged at 5,000 × g for 30 min at 4°C. Of the supernatant, 900 μl was discarded. Luciferase mRNA (9.9 × 108 copies; Promega, Mannheim, Germany) was added to the residual 100-μl volume and served as an internal standard for normalization due to losses during mRNA preparation and reverse transcription (RT) inefficiencies. RNA was extracted immediately using the Total RNA Mini kit (A&A Biotechnologie, Gdynia, Poland). Contaminating DNA was removed using a DNase I kit (Fermentas, St. Leon-Rot, Germany) with treatment for 3 h. RNA was stored at −80°C. No DNA contamination was detected in RNA using DNase I-treated RNA as the template for qPCR (see below). The recovery of luciferase mRNA typically ranged between 5 and 20%.

RT.

RNA was quantified using a GeneQuant RNA/DNA calculator (Pharmacia Biosystems, Freiburg, Germany) at a wavelength of 260 nm. Within each experiment, equal amounts of sample RNA in the range of 0.1 to 1 μg were subjected to RT using random hexamer primers and the RevertAid H Minus first-strand cDNA synthesis kit (Fermentas) according to the manufacturer's recommendations.

PCR amplification of rdhA targets from DNA or cDNA.

Thirteen degenerate primer pairs were used to amplify specific fragments of all 32 rdhA genes of strain CBDB1 (see Table S1 in the supplemental material). The forward primers were labeled with 6-carboxyfluorescein (FAM). The PCR mixture (30 μl) contained 1 μl of DNA or cDNA, 1× reaction buffer, 3 mM MgCl2, 200 μM of each of the four deoxynucleoside triphosphates (Bioline, Luckenwalde, Germany), 210 nM of each primer (Metabion, Martinsried, Germany), and 0.1 U of HotStarTaq polymerase (Qiagen, Hildesheim, Germany). The PCR was performed using the following conditions: an initial incubation of 95°C for 15 min, followed by 36 to 50 cycles (for cDNA amplification) or 40 cycles (for DNA amplification) of 30 s at 94°C, 30 s at 45 to 58°C, 1 min at 72°C, and a final extension of 10 min at 72°C. The sensitivity of the PCR assay to detect cluster 3 and cluster 4b rdhA targets was analyzed using total DNA of strain CBDB1. First, the copy number of the genome in extracted total DNA of strain CBDB1 was determined as 8 × 105 copies μl−1 by qPCR targeting the 16S rRNA gene (see below). Total DNA was serially diluted from 8 × 105 to 100 genome copies μl−1, and 1 μl of each dilution served as the template for PCR. Cluster 3 and cluster 4b primers yielded PCR products from at least 80 and 800 copies per μl, respectively.

t-RFLP for the differentiation of dehalogenase gene transcripts.

The fluorescently labeled PCR products were purified using a QIAquick PCR purification kit (Qiagen) and analyzed on 1% (wt/vol) agarose gels. The amplicons of each degenerate primer pair were quantified by GeneQuant (GE Healthcare, Freiburg, Germany) and divided into aliquots. Each aliquot (100 to 500 ng of DNA) was digested with a single restriction enzyme (1 U of MspI, RsaI, AluI, MboI, FnuDII, PstI, or BsuRI; Fermentas) and the recommended restriction buffer for 5 h at 37°C in a total volume of 10 μl. A suitable set of restriction enzymes was applied to the products of each degenerate primer pair to obtain specific terminally labeled restriction fragments of all amplified dehalogenase genes or transcripts (Table (Table1).1). Digested PCR products were precipitated using standard techniques and vacuum dried using a SpeedVac SVC 100. The terminal restriction fragments (t-RFs) were size separated on either an ABI Prism 377 automated sequencer or an ABI Prism 3100 genetic analyzer (Applied Biosystems, Weiterstadt, Germany). By comparison with an internal size standard [ROX Genescan 500(−250)], the fragment length was determined in a range between 50 and 500 bp using the Genescan 2.1 software (PE Applied Biosystems). t-RFs were assigned to specific rdhA genes by a comparison to the results of computational digests (Clone 4.0 software; Scientific & Educational Software, Durham, NC) of the appropriate gene sequences (Table (Table11).

TABLE 1.
t-RFs based on a computational digest of the rdhA gene sequences of strain CBDB1

Quantification of genes and transcripts.

The copy numbers of the genes and corresponding transcripts of 10 putative reductive dehalogenases (CbdbA84=CbrA, CbdbA1624, CbdbA1453, CbdbA187, CbdbA1588, CbdbA1618, CbdbA1563, CbdbA243, CbdbA80, and CbdbA88), three predicted house-keeping proteins (RpoA, RpoB, and EF-Tu), and luciferase were measured by qPCR using the primers indicated in Table S1 in the supplemental material. The 16S rRNA gene copy number was quantified as previously reported (8). Reactions were performed in 20-μl volumes containing 10 μl QuantiTect SYBR green PCR master mix (Qiagen), 1.2 μl of each primer (5 pmol/μl), 6.6 μl of distilled water, and 1 μl of template DNA or cDNA. The PCR was performed using a Rotor Gene 6.0 qPCR machine (Corbett Research, Sydney, Australia). The thermocycling program was as follows: an initial step of 15 min at 95°C, followed by 40 to 44 cycles of 15 s at 94°C and 30 s at 55 to 59°C depending on the primer annealing temperature, and then 30 s at 72°C. Fluorescence data were collected after each elongation step. For the preparation of standard curves, PCR products of the genes of interest amplified with the primers specified in Table S1 in the supplemental material were purified using the QIAquick PCR purification kit (Qiagen), ligated into the pGEM T-Easy vector (Promega), and transformed into E. coli XL1-Blue MRF′ (Stratagene), and plasmids were prepared using the QIAprep Spin Miniprep kit (Qiagen). Plasmid-DNA (Table (Table2)2) was serially diluted from 109 to 101 copies μl−1 and measured in triplicate, resulting in a typical standard curve. In each run, three concentrations of the plasmid covering four orders of magnitude were included as external standards for quantification. Samples and external standards were analyzed in duplicate. The transcription level of the rdhA genes was calculated as the ratio of the luciferase mRNA-normalized copy number of the rdhA transcript and the copy number of the corresponding rdhA gene.

TABLE 2.
Plasmids used as qPCR standards

RESULTS

Dechlorination of 1,2,3- and 1,2,4-TCB.

The dechlorination of 1,2,3- or 1,2,4-TCB was studied with cells pregrown with 1,2,4-TCB. After 72 h, 1,2,3-TCB dechlorination led to the formation of 62 μM 1,3-DCB as the sole product (Fig. (Fig.1A),1A), while 1,2,4-TCB was dechlorinated to a mixture of 17 μM 1,3-DCB (34%) and 33 μM 1,4-DCB (66%) (Fig. (Fig.1B).1B). The TCBs were converted to levels below the detection limit after 72 h of cultivation (data not shown). The copy number of the 16S rRNA gene was monitored during the dechlorination process by qPCR and was relatively constant within the first 72 h of active dechlorination.

FIG. 1.
Reductive dechlorination of 65 μM 1,2,3-TCB (A) and 50 μM 1,2,4-TCB (B) and the copy number of the 16S rRNA gene (•). Dechlorination activity is demonstrated by the formation of the dechlorination products 1,3-DCB ([filled triangle]) and ...

Establishment of a t-RFLP protocol for detection and differentiation of rdhA genes and transcripts.

A t-RFLP approach was used to detect and distinguish all 32 rdhA sequences of Dehalococcoides sp. strain CBDB1 in DNA or cDNA. The attempt to design one degenerate primer pair for the amplification of all 32 rdhA targets failed because of the low overall sequence similarity. According to a phylogenetic tree (24), the rdhA gene sequences were grouped into four main clusters, which were further subdivided into 13 subgroups, each of which comprised one to five rdhA genes. One primer pair was designed for every subgroup. With one exception (cluster 2a-2_r), primers were degenerate (see Table S1 in the supplemental material). The reverse primers targeting cluster 4a and 4b were derived from the respective rdhB genes. For the detection and differentiation of single fluorescently labeled rdhA amplicons, two to four restriction enzymes were selected based on a computational digest of the sequences. The theoretically expected t-RFs were experimentally detected using genomic DNA of strain CBDB1 as a template for PCR and restriction analysis. Slight differences between the calculated and the practically determined sizes of the t-RFs (2 to 5 bp) occurred and presumably are caused by unpredictable sizing errors introduced by differences in the nucleotide composition of the internal size standard and the target sequences (42). The theoretical lengths of the t-RFs are given in Table Table11.

Transcription of rdhA genes during dechlorination of 1,2,3- and 1,2,4-TCB.

After a starvation phase of 72 h, several parallel cultures were spiked with 1,2,3- or 1,2,4-TCB. Total RNA was extracted from cultures at different times and subjected to RT and subsequent PCR with 13 degenerate primer pairs. At time zero, no or only minor amounts of RT-PCR products were detectable (Fig. (Fig.2).2). After 24 or 48 h at the latest, RT-PCR products were obtained with all primers except the cluster 3 primer pair, which did not yield PCR products from cDNA throughout repeated experiments. With genomic DNA as the template, a PCR product was formed with the cluster 3 primers, suggesting that the respective rdhA genes were not transcribed or were transcribed at a very low level. A low transcript level was confirmed later with primers specifically targeting the individual rdhA transcripts (see below). The intensity of bands representing different transcribed rdhA clusters was relatively constant between the data points after 24 and 72 h but showed a sharp decrease between 72 and 168 h after induction. This decrease of rdhA transcription level corresponded with the cessation of dechlorination activity in the cultures (Fig. (Fig.11).

FIG. 2.
Composite figure showing sections of gels with RT-PCR products obtained with 13 different degenerate primer pairs from cultures supplemented at time point 0 with 1,2,3-TCB (A) or 1,2,4-TCB (B). The approximate sizes of the respective amplicons according ...

t-RFLP analysis of the FAM-labeled RT-PCR products from cultures 48 h after induction revealed the presence of transcripts of all 29 rdhA genes captured by the 12 PCR product-forming cluster primer pairs, regardless of whether 1,2,3- or 1,2,4-TCB was the electron acceptor (Fig. (Fig.2C).2C). The t-RFLP profiles recognizing more than one rdhA fragment additionally were examined for differences in the relative peak areas by comparing the profiles obtained from cDNA of 1,2,3-TCB- and 1,2,4-TCB-grown cells and from genomic DNA. The t-RFLP profiles from genomic DNA reflected the amplification efficiency of the different rdhA targets by the respective primer pair. Examples are given in Fig. Fig.33 (also see Fig. S1 in the supplemental material). The relative abundance of three transcripts varied in response to the spiked TCBs, as indicated by the percentage of their t-RF peak areas relative to the total fluorescence intensity observed in the 50- to 500-nucleotide size range (Fig. (Fig.3).3). The genes cbdbA1453 and cbdbA187 were more strongly transcribed in the presence of 1,2,3-TCB, while cbdbA1624 was more strongly transcribed in the presence of 1,2,4-TCB. Furthermore, the transcript of cbrA (2) (=cbdbA84; cluster 2a_1 [24]) showed elevated relative peak areas in t-RFLP profiles of cDNA obtained from cells grown in both 1,2,3- and 1,2,4-TCB compared to levels shown by genomic DNA profiles. Using the cluster primer pair 2a_f/2a-1_r, fragments of cbrA and cbdbA1563 were coamplified. Whereas the ratio of relative peak areas of cbrA and cbdbA1563 t-RFs was, on average, 0.8:1 for genomic DNA (n = 2), the ratio was between 2:1 and 4:1 for cDNA obtained from cultures grown in 1,2,3-TCB (n = 4) or 1,2,4-TCB (n = 4) (see Fig. S1 in the supplemental material). cbrA recently was shown to code for a TCB-reductive dehalogenase (2). The t-RFLP approach did not visualize clear differential responses in other clusters of rdhA transcripts.

FIG. 3.
t-RFLP profiles obtained from genomic DNA of strain CBDB1 (A and B), from cDNA of 1,2,3-TCB-supplemented cultures (C and D), and from cDNA of 1,2,4-TCB-supplemented cultures (E and F) amplified with the degenerate primer pairs cluster 1_af/r (left) and ...

To determine the influence of the solvent acetone on rdhA gene expression, parallel cultures were treated with acetone and transcription was analyzed by RT-PCR and t-RFLP. RT-PCR products were amplified with two primer pairs (clusters 1a and 1c). An induction was not observed during the whole time period; in contrast, the amounts of the RT-PCR products decreased over time (data not shown).

rdhA gene transcript levels.

In the literature, two different standards were described for the normalization of the transcription level quantification of rdhA genes in Dehalococcoides spp. The copy number of the rdhA transcripts was related to the transcript level of rpoB (11, 32, 36) or to the copy number of the rdhA gene itself (19, 20, 25). To find an appropriate standard for the quantification of the temporal variability of rdhA gene transcription in Dehalococcoides sp. strain CBDB1, the transcription of three house-keeping genes (tuf, rpoA, rpoB, which code for translation elongation factor TU and two subunits of RNA polymerase, respectively) and of the 16S rRNA gene was determined by RT-qPCR in 1,2,3-TCB-fed cultures. Fig. S2 in the supplemental material shows that rpoB, rpoA, and tuf mRNA levels increased 10- to 100-fold within 24 h after 1,2,3-TCB was supplied. Starting from an initial level that was several orders of magnitude higher, the 16S rRNA level also increased, but only by 1.6-fold. The relatively stable copy number of the 16S rRNA gene (Fig. (Fig.1)1) suggested that the observed relative changes in the copy number of the house-keeping gene transcripts reflected differences in transcription rates rather than in cell numbers. Therefore, a cell number-based standard for the normalization of the transcription data was preferred in the current study. The copy number of each rdhA transcript was related to the copy number of the respective gene, thus allowing a direct comparison of the transcript levels of different rdhA genes despite the different amplification efficiencies with different primers and targets.

For qPCR studies of rdhA gene transcription, rdhA genes were selected that represented a specific response pattern to 1,2,3- and 1,2,4-TCB in the t-RFLP analyses. The rdhA genes cbdbA1624, cbdbA1453, and cbdbA187 were chosen because they showed a specific response to either 1,2,3- or 1,2,4-TCB. The transcription of cbrA was monitored by qPCR to prove its upregulation by both TCBs. Its partner gene in cluster 2a_1, cbdbA1563, also was analyzed to allow a direct comparison to the results of t-RFLP. Additionally, the transcription of cbdbA1588, a member of cluster 1c, was quantified with no obvious differential response. cbdbA1588 encodes an orthologue of a PCE-reductive and putative 2,3-dichlorophenol-reductive dehalogenase of strain 195 (11, 31).

All targets studied showed an increase of at least one order of magnitude in transcription levels within the first 24 h of culture and continued to show high expression levels (Fig. (Fig.4),4), and active dechlorination was observed (Fig. (Fig.1).1). For both 1,2,3- and 1,2,4-TCB, the highest transcription levels were observed for cbrA. The second highest expression level was observed with cbdbA1453 and cbdbA1624 in cultures with 1,2,3-TCB and 1,2,4-TCB, respectively, which is in accord with the t-RFLP results. As was also found by t-RFLP analysis, cbdbA187 transcripts reached a higher abundance with 1,2,3-TCB than with 1,2,4-TCB as the electron acceptor. However, it is interesting that the initial amounts of the different rdhA transcripts varied considerably. Whereas the transcripts of cbrA, cbdbA1624, and/or cbdbA1453 started at a similarly high level, the cbdbA187 and cbdbA1588 transcript levels were initially one to two orders of magnitude less abundant, and the cbdbA1563 level (not shown) was even three to four orders of magnitude less abundant. After 168 h, the transcript levels of all studied rdhA genes were reduced almost to their initial levels. The cbrA transcript remained at a comparatively high level, suggesting a mechanism that maintains a relatively high transcription level of this gene even after complete substrate conversion. To test the influence of the congener used to grow the preculture on rdhA gene expression in the analyzed cultures, similar experiments were set up in duplicate with cells pregrown with 1,2,3-TCB instead of 1,2,4-TCB. Fig. S3 in the supplemental material shows that similar results were obtained with this approach. In these experiments, cbdbA1618 was monitored instead of cbdbA1588, which also was clearly induced by both substrates but showed a relatively low transcript abundance level.

FIG. 4.
Quantitative transcription analysis of the five selected rdhA genes cbrA ([filled triangle]), cbdbA1624 (•), cbdbA1453 ([filled triangle]), cbdbA1588 ([filled square]), and cbdbA187 (♦) of strain CBDB1 in the presence of 1,2,3-TCB (A) or 1,2,4-TCB (B) or in ...

RT-qPCR also was used to study whether cluster 3 rdhA gene transcripts that were not found by the previous RT-PCR with degenerate primers were present or not in the mRNA pool. With the specific primers listed in Table S1 in the supplemental material, individual transcripts of cbdbA243, cbdbA80, and cbdbA88 were detected at a concentration of 10−3 to 10−4 transcripts per gene copy in the beginning (data not shown). All three genes were induced more than one to two orders of magnitude. After 48 h, they were determined at a concentration comparable to the low abundance of cbdbA1588 and cbdbA1563 transcripts (see Fig. S4 in the supplemental material).

To investigate a possible influence of the solvent acetone on transcript levels, control cultures were analyzed (Fig. (Fig.4C).4C). The levels of the transcription of cbrA and cbdbA1453 showed low temporal variations. A slight increase in the transcript levels was observed for cbdbA1624, cbdbA187, and cbdbA1588; however, these levels were low compared to those of the cultures fed with 1,2,3- and 1,2,4-TCB (Fig. 4A, B). It is interesting that the transcripts were detectable at a similarly low level even after 168 h of chlorobenzene starvation in these cultures, which succeeded even after an initial 72-h starvation in the preincubation phase.

DISCUSSION

The aim of this study was to describe the induction of rdhA genes in strain CBDB1 by TCBs. To allow the monitoring of all 32 genes, we established a new procedure based on the t-RFLP analysis of RT-PCR products. The t-RFLP approach is used frequently for the analysis of community structures of mixed cultures on the level of both 16S rRNA genes and transcripts (21, 35). In addition, functional genes have been used as a target for t-RFLP analysis to characterize physiological groups of microorganisms like ammonia-oxidizing bacteria, methanotrophs, methanogens, and nitrate-reducing or nitrogen-fixing bacteria (5, 17, 22, 27, 39). The rdhA genes encode products that share common features, like the presence of a TAT signal sequence and two motifs for the coordination of an iron-sulfur cluster (16, 24), which have been used as targets in primer design. In previous studies, a degenerate forward primer (RRF2) was derived from the TAT motif for the detection of different rdhA transcripts in strain BAV1 and a Dehalococcoides species-containing mixed culture KB1 (23, 44). However, the evaluation of the primer against the full genome sequence of strain CBDB1 revealed that RRF2 showed a perfect match to only four rdhA gene sequences, whereas the other rdhA sequences possessed one to six mismatches in the target region. In addition, the TAT consensus signal is not specific for rdhA gene products but is found in numerous other exported or membrane-targeted proteins (34), which prompted us to search for different primer locations, as has been suggested previously (37). The high overall sequence variability of rdhA genes (16) required the design of 13 different degenerate primer pairs, each recognizing one to five individual rdhA sequences of strain CBDB1 and producing fragments of 400 to 650 bp in length. As verified with genomic DNA, this method was appropriate to monitor the 32 different rdhA targets in strain CBDB1. However, primer degeneracy implies difficulties in detecting transcripts at low levels, as suggested by the failure of cluster 3 primers to amplify their targets from cDNA. Alternative methods like the cloning and sequencing of RT-PCR products (44) or RT-qPCR analyses of the whole set of rdhA genes (11) have been used previously. Compared to these methods, the t-RFLP approach was less labor-intensive and allowed a faster throughput of samples from different sampling times, leading to (i) the visualization of the induction of rdhA gene clusters by comparing band intensities of RT-PCR products in agarose gels (Fig. (Fig.2);2); (ii) the indication of specific transcripts in the mRNA pool; and (iii) the detection of differentially expressed rdhA genes by comparing relative peak areas of t-RFLP profiles obtained from cultures incubated with different electron acceptors (Fig. (Fig.3).3). Considering a biased PCR amplification of specific rdhA genes due to primer degeneracy (27), the t-RFLP approach provides a measure of transcript levels relative to that of profiles obtained from genomic DNA and allows only a semiquantitative comparison.

Using our RT-PCR/t-RFLP approach, we could find clear indications for 29 of the 32 rdhA genes in strain CBDB1 that they are transcribed in cultures with 1,2,3- or 1,2,4-TCB (Fig. (Fig.2).2). The expression of the complete, or a slightly reduced, set of rdhA genes recently was shown in D. ethenogenes strain 195 for growth on PCE and 2,3-dichlorophenol, respectively (11). The simultaneous transcription of some rdhA genes of a Dehalococcoides sp. strain in the KB1 mixed culture during the dechlorination of less chlorinated ethenes was reported previously (44). This complex transcriptional response suggests a global regulatory control that might be growth dependent. Canonical promoter sequences showing −10 and −35 regions of the σ70 type have been identified upstream of rdhA genes in strains 195 (11) and VS (33). Accordingly, the genomes of Dehalococcoides sp. strains 195 and CBDB1 both contain σ70 homologues (24, 38). Putative activator or repressor proteins of the MarR type or two-component regulatory protein family are encoded by genes that are in close proximity to the rdhA genes in strains CBDB1 and 195 and might be involved in the fine-tuning of transcription (24). Indeed, the observed transcript levels of different rdhA genes in cells of strain CBDB1 varied by several orders of magnitude. Assuming similar extraction efficiencies for DNA and RNA and one genome copy per cell, the normalization of rdhA transcript numbers by the number of each corresponding rdhA gene gives a rough estimate of transcript numbers per cell. Thus, at time zero, the 10 monitored rdhA transcripts varied between one transcript per 10,000 cells (cbdbA1563 and cbdbA243) to one transcript per cell (cbrA). Subsequently, all 10 rdhA transcripts increased in number, reaching a maximum before dechlorination was complete, followed by a slow decay in levels and finally approaching the initial level (Fig. 4A, B; also see Fig. S3 in the supplemental material). This general pattern of induction and mRNA turnover also was reported for tceA and three further rdhA genes in Dehalococcoides species-containing mixed cultures (19, 25, 36). The final low transcript level, which was reached several days after complete dehalogenation, proved to be very stable, as demonstrated for tceA (25) and in the current study for the controls with acetone not supplemented with a chlorinated substrate. Whether this apparent longevity of mRNA is due to constitutive expression at a low level or to the stabilization of the mRNA presently is not known.

In the present study, the highest transcript level (10 to a maximum of 22 transcripts per rdhA gene) was indicated for cbrA, which codes for the recently identified chlorobenzene-reductive dehalogenase in strain CBDB1 (2). Isolated CbrA reductively dehalogenated 1,2,3-TCB to 1,3-DCB and 1,2,3,4-tetrachlorobenzene to 1,2,4-TCB, but the latter was not further converted, leaving open the question of whether CbrA also is responsible for 1,2,4-TCB dechlorination in CBDB1. Our results demonstrate that both 1,2,3- and 1,2,4-TCB at least induced the transcription of cbrA to about 10-fold above the initial level and also above the level of most other rdhA gene transcripts. It would be of great interest to study other chlorinated benzenes or dibenzo-p-dioxins to determine whether they also act as efficient inducers or even constitute substrates of CbrA.

A similar correlation between a high transcript level (11) and the detectability of the corresponding protein with high coverage by mass spectrometry (31) has been demonstrated for D. ethenogenes strain 195 growing with PCE or 2,3-dichlorophenol. These findings also suggested a dual function for PceA as a PCE- and 2,3-dichlorophenol-reductive dehalogenase. In addition, CbdbA1588, the PceA homologue in strain CBDB1, was the dominant RdhA in membrane-enriched cell fractions after growth with 2,3-dichlorophenol (31). cbdbA1588 was not detected in cell extracts of strain CBDB1 after growth with 1,2,3-TCB (2), and the respective gene was transcribed only at a low level. Both findings suggest no direct involvement in TCB dechlorination.

Remarkably, transcripts of the cluster 3 rdhA genes cbdbA80, cbdbA88, and cbdbA243 were hardly detectable in mRNA of cells grown in TCB. Recently, the expression of CbdbA80 during growth on 1,2,3-TCB was observed when the preculture contained 1,2,4-TCB but not when the preculture was cultivated with a mixture of 1,2,3- and 1,2,4-TCB (2). In addition, CbdbA80 was detected in a 2,3-dichlorophenol-dechlorinating culture of strain CBDB1 (31). The homologue DET1559 in strain 195 was expressed in a PCE-dechlorinating culture (31). Thus, it seems that cbdbA80 is not induced by one specific chlorinated compound. The reductive dehalogenase CbdbA88 recently was detected in membrane-enriched protein extracts, albeit with a low protein sequence coverage, during the growth of strain CBDB1 on 2,3-dichlorophenol (31). It was not detected in extracts of cells grown in 1,2,3-TCB (2). Therefore, a specific role for CbdbA88 in TCB dechlorination most likely can be excluded.

Two rdhA genes, cbdbA1453 and cbdbA1624, showed an intermediate transcription level with initial and maximum levels similar to, or no more than 10-fold lower than, those of cbrA (Fig. (Fig.4;4; also see Fig. S3 in the supplemental material). Both proteins share a 91.7% identity and are the two most similar RdhA proteins in strain CBDB1. Orthologues are encoded by the genomes of Dehalococcoides sp. strains FL2 and VS but are absent in strain 195. The relative transcript level of cbdbA1453 could be shown reproducibly to exceed that of cbdbA1624 in the presence of 1,2,3-TCB, whereas this ratio was reversed when 1,2,4-TCB was supplied as the electron acceptor, suggesting that a specific regulation had taken place. The cbdbA1453 and cbdbA1624 genes are located in the vicinity of genes encoding MarR-type regulators (24). This contrasts with the situation for the highly expressed cbrA and cbdbA1588 genes, which are associated with two-component regulatory genes. MarR-type regulators are known to be involved in the regulation of the catabolism of aromatic compounds, frequently acting as repressors (45). Possibly, a relatively tight (auto)regulation prevents high transcript and protein levels, complicating the detection by a proteomic approach. Currently, the function of cbdbA1453 and cbdbA1624 gene products in the dechlorination of TCBs is unknown. It is interesting that in addition to the orthologues of cbdbA1453 and cbdbA1624, the cbrA orthologue also is absent from strain 195, which cannot dechlorinate 1,2,3- and 1,2,4-TCB in the absence of PCE as a cosubstrate. In addition, a further Dehalococcoides sp. strain was isolated recently (8) that is able to grow by respiratory dehalogenation with chlorinated dibenzo-p-dioxins and 1,2,3-TCB but not with 1,2,4-TCB. This strain has homologues of cbrA and cbdbA1453 but not of cbdbA1624 (unpublished results), further strengthening the correlation between the presence of cbdbA1624 and the ability of strain CBDB1 to dechlorinate 1,2,4-TCB.

If the function of a distinct RdhA protein is known, the corresponding gene and mRNA can serve as a biomarker for the characterization of a specific in situ dehalogenation potential or activity (19, 20). Recently, a correlation was demonstrated between vcrA and bvcA expression and the active in situ dechlorination of cis-DCE to ethene at a field site (26). The suggested function of CbrA as a chlorobenzene dehalogenase and its currently unique presence in the chlorobenzene-dehalorespiring strain CBDB1 suggest the use of cbrA as a functional marker for the natural potential of chlorobenzene dehalogenation at contaminated sites. Although more research is needed before a quantitative prediction of dechlorination rates can be derived, the presence of cbrA transcripts could be indicative of a capacity to dechlorinate chlorobenzenes.

Supplementary Material

[Supplemental material]

Acknowledgments

This work was supported by a grant to A.W., U.L., and J.R.A. from the Graduiertenkolleg 416 by the Deutsche Forschungsgemeinschaft.

We thank Gary Sawers for helpful comments on the manuscript.

Footnotes

[down-pointing small open triangle]Published ahead of print on 5 February 2009.

Supplemental material for this article may be found at http://aem.asm.org/.

REFERENCES

1. Adrian, L., S. K. Hansen, J. M. Fung, H. Görisch, and S. H. Zinder. 2007. Growth of Dehalococcoides strains with chlorophenols as electron acceptors. Environ. Sci. Technol. 41:2318-2323. [PubMed]
2. Adrian, L., J. Rahnenführer, J. Gobom, and T. Hölscher. 2007. Identification of a chlorobenzene reductive dehalogenase in Dehalococcoides sp. strain CBDB1. Appl. Environ. Microbiol. 73:7717-7724. [PMC free article] [PubMed]
3. Adrian, L., U. Szewzyk, J. Wecke, and H. Görisch. 2000. Bacterial dehalorespiration with chlorinated benzenes. Nature 408:580-583. [PubMed]
4. Bopp, R. F., M. L. Gross, H. Tong, H. J. Simpson, S. J. Monson, B. L. Deck, and F. C. Moser. 1991. A major incident of dioxin contamination: sediments of New Jersey estuaries. Environ. Sci. Technol. 25:951-956.
5. Bremer, C., G. Braker, D. Matthies, A. Reuter, C. Engels, and R. Conrad. 2007. Impact of plant functional group, plant species, and sampling time on the composition of nirK-type denitrifier communities in soil. Appl. Environ. Microbiol. 73:6876-6884. [PMC free article] [PubMed]
6. Bunge, M., L. Adrian, A. Kraus, M. Opel, W. G. Lorenz, J. R. Andreesen, H. Görisch, and U. Lechner. 2003. Reductive dehalogenation of chlorinated dioxins by an anaerobic bacterium. Nature 421:357-360. [PubMed]
7. Bunge, M., M. A. Kähkönen, W. Rämisch, M. Opel, S. Vogler, F. Walkow, M. Salkinoja-Salonen, and U. Lechner. 2007. Biological activity in a heavily organohalogen-contaminated river sediment. Environ. Sci. Pollut. R. 14:3-10. [PubMed]
8. Bunge, M., A. Wagner, M. Fischer, J. R. Andreesen, and U. Lechner. 2008. Enrichment of a dioxin-dehalogenating Dehalococcoides species in two-liquid phase cultures. Environ. Microbiol. 10:2670-2683. [PubMed]
9. Ewald, E. M., A. Wagner, I. Nijenhuis, H. H. Richnow, and U. Lechner. 2007. Microbial dehalogenation of trichlorinated dibenzo-p-dioxins by a Dehalococcoides-containing mixed culture is coupled to carbon isotope fractionation. Environ. Sci. Technol. 41:7744-7751. [PubMed]
10. Fennell, D. E., I. Nijenhuis, S. F. Wilson, S. H. Zinder, and M. M. Häggblom. 2004. Dehalococcoides ethenogenes strain 195 reductively dechlorinates diverse chlorinated aromatic pollutants. Environ. Sci. Technol. 38:2075-2081. [PubMed]
11. Fung, J. M., R. M. Morris, L. Adrian, and S. H. Zinder. 2007. Expression of reductive dehalogenase genes in Dehalococcoides ethenogenes strain 195 growing on tetrachloroethene, trichloroethene, or 2,3-dichlorophenol. Appl. Environ. Microbiol. 73:4439-4445. [PMC free article] [PubMed]
12. He, J., K. M. Ritalahti, K. L. Yang, S. S. Koenigsberg, and F. E. Löffler. 2003. Detoxification of vinyl chloride to ethene coupled to growth of an anaerobic bacterium. Nature 424:62-65. [PubMed]
13. He, J., Y. Sung, R. Krajmalnik-Brown, K. M. Ritalahti, and F. E. Löffler. 2005. Isolation and characterization of Dehalococcoides sp. strain FL2, a trichloroethene (TCE)- and 1,2-dichloroethene-respiring anaerobe. Environ. Microbiol. 7:1442-1450. [PubMed]
14. Holliger, C., G. Wohlfahrt, and G. Diekert. 1999. Reductive dechlorination in the energy metabolism of anaerobic bacteria. FEMS Microbiol. Rev. 22:383-398.
15. Hölscher, T., H. Görisch, and L. Adrian. 2003. Reductive dehalogenation of chlorobenzene congeners in cell extracts of Dehalococcoides sp. strain CBDB1. Appl. Environ. Microbiol. 69:2999-3001. [PMC free article] [PubMed]
16. Hölscher, T., R. Krajmalnik-Brown, K. M. Ritalahti, F. von Wintzingerode, H. Görisch, F. E. Löffler, and L. Adrian. 2004. Multiple nonidentical reductive-dehalogenase-homologous genes are common in Dehalococcoides. Appl. Environ. Microbiol. 70:5290-5297. [PMC free article] [PubMed]
17. Horz, H. P., J. H. Rotthauwe, T. Lukow, and W. Liesack. 2000. Identification of major subgroups of ammonia-oxidizing bacteria in environmental samples by T-RFLP analysis of amoA PCR products. J. Microbiol. Methods 39:197-204. [PubMed]
18. Jayachandran, G., H. Görisch, and L. Adrian. 2003. Dehalorespiration with hexachlorobenzene and pentachlorobenzene by Dehalococcoides sp. strain CBDB1. Arch. Microbiol. 180:411-416. [PubMed]
19. Johnson, D. R., P. K. Lee, V. F. Holmes, and L. Alvarez-Cohen. 2005. An internal reference technique for accurately quantifying specific mRNAs by real-time PCR with application to the tceA reductive dehalogenase gene. Appl. Environ. Microbiol. 71:3866-3871. [PMC free article] [PubMed]
20. Johnson, D. R., P. K. Lee, V. F. Holmes, A. C. Fortin, and L. Alvarez-Cohen. 2005. Transcriptional expression of the tceA gene in a Dehalococcoides-containing microbial enrichment. Appl. Environ. Microbiol. 71:7145-7151. [PMC free article] [PubMed]
21. Kittelmann, S., and M. W. Friedrich. 2008. Novel uncultured Chloroflexi dechlorinate perchloroethene to trans-dichloroethene in tidal flat sediments. Environ. Microbiol. 10:1557-1570. [PubMed]
22. Knauth, S., T. Hurek, D. Brar, and B. Reinhold-Hurek. 2005. Influence of different Oryza cultivars on expression of nifH gene pools in roots of rice. Environ. Microbiol. 7:1725-1733. [PubMed]
23. Krajmalnik-Brown, R., T. Hölscher, I. N. Thomson, F. M. Saunders, K. M. Ritalahti, and F. E. Löffler. 2004. Genetic identification of a putative vinyl chloride reductase in Dehalococcoides sp. strain BAV1. Appl. Environ. Microbiol. 70:6347-6351. [PMC free article] [PubMed]
24. Kube, M., A. Beck, S. H. Zinder, H. Kuhl, R. Reinhardt, and L. Adrian. 2005. Genome sequence of the chlorinated compound-respiring bacterium Dehalococcoides species strain CBDB1. Nat. Biotechnol. 23:1269-1273. [PubMed]
25. Lee, P. K., D. R. Johnson, V. F. Holmes, J. He, and L. Alvarez-Cohen. 2006. Reductive dehalogenase gene expression as a biomarker for physiological activity of Dehalococcoides spp. Appl. Environ. Microbiol. 72:6161-6168. [PMC free article] [PubMed]
26. Lee, P. K., T. W. Macbeth, K. S. Sorenson, Jr., R. A. Deeb, and L. Alvarez-Cohen. 2008. Quantifying genes and transcripts to assess the in situ physiology of Dehalococcoides spp. in a trichloroethene-contaminated groundwater site. Appl. Environ. Microbiol. 74:2728-2739. [PMC free article] [PubMed]
27. Lueders, T., and M. W. Friedrich. 2003. Evaluation of PCR amplification bias by terminal restriction fragment length polymorphism analysis of small-subunit rRNA and mcrA genes by using defined template mixtures of methanogenic pure cultures and soil DNA extracts. Appl. Environ. Microbiol. 69:320-326. [PMC free article] [PubMed]
28. Magnuson, J. K., M. F. Romine, D. R. Burris, and M. T. Kingsley. 2000. Trichloroethene reductive dehalogenase from Dehalococcoides ethenogenes: Sequence of tceA and substrate range characterization. Appl. Environ. Microbiol. 66:5141-5147. [PMC free article] [PubMed]
29. Magnuson, J. K., R. V. Stern, J. M. Gossett, S. H. Zinder, and D. R. Burris. 1998. Reductive dechlorination of tetrachloroethene to ethene by two-component enzyme pathway. Appl. Environ. Microbiol. 64:1270-1275. [PMC free article] [PubMed]
30. Maymó-Gatell, X., Y. Chien, J. M. Gossett, and S. H. Zinder. 1997. Isolation of a bacterium that reductively dechlorinates tetrachloroethene to ethene. Science 276:1568-1571. [PubMed]
31. Morris, R. M., J. M. Fung, B. G. Rahm, S. Zhang, D. L. Freedman, S. H. Zinder, and R. E. Richardson. 2007. Comparative proteomics of Dehalococcoides spp. reveals strain-specific peptides associated with activity. Appl. Environ. Microbiol. 73:320-326. [PMC free article] [PubMed]
32. Morris, R. M., S. Sowell, D. Barofsky, S. Zinder, and R. Richardson. 2006. Transcription and mass-spectroscopic proteomic studies of electron transport oxidoreductases in Dehalococcoides ethenogenes. Environ. Microbiol. 8:1499-1509. [PubMed]
33. Müller, J. A., B. M. Rosner, G. Von Abendroth, G. Meshulam-Simon, P. L. McCarty, and A. M. Spormann. 2004. Molecular identification of the catabolic vinyl chloride reductase from Dehalococcoides sp. strain VS and its environmental distribution. Appl. Environ. Microbiol. 70:4880-4888. [PMC free article] [PubMed]
34. Natale, P., T. Brüser, and A. J. Driessen. 2008. Sec- and Tat-mediated protein secretion across the bacterial cytoplasmic membrane—distinct translocases and mechanisms. Biochim. Biophys. Acta 1778:1735-1756. [PubMed]
35. Osborn, A. M., E. R. Moore, and K. N. Timmis. 2000. An evaluation of terminal-restriction fragment length polymorphism (T-RFLP) analysis for the study of microbial community structure and dynamics. Environ. Microbiol. 2:39-50. [PubMed]
36. Rahm, B. G., R. M. Morris, and R. E. Richardson. 2006. Temporal expression of respiratory genes in an enrichment culture containing Dehalococcoides ethenogenes. Appl. Environ. Microbiol. 72:5486-5491. [PMC free article] [PubMed]
37. Regeard, C., J. Maillard, and C. Holliger. 2004. Development of degenerate and specific PCR primers for the detection and isolation of known and putative chloroethene reductive dehalogenase genes. J. Microbiol. Methods 56:107-118. [PubMed]
38. Seshadri, R., L. Adrian, D. E. Fouts, J. A. Eisen, A. M. Phillippy, B. A. Methe, N. L. Ward, W. C. Nelson, R. T. Deboy, H. M. Khouri, J. F. Kolonay, R. J. Dodson, S. C. Daugherty, L. M. Brinkac, S. A. Sullivan, R. Madupu, K. E. Nelson, K. H. Kang, M. Impraim, K. Tran, J. M. Robinson, H. A. Forberger, C. M. Fraser, S. H. Zinder, and J. F. Heidelberg. 2005. Genome sequence of the PCE-dechlorinating bacterium Dehalococcoides ethenogenes. Science 307:105-108. [PubMed]
39. Shrestha, M., W. R. Abraham, P. M. Shrestha, M. Noll, and R. Conrad. 2008. Activity and composition of methanotrophic bacterial communities in planted rice soil studied by flux measurements, analyses of pmoA gene and stable isotope probing of phospholipid fatty acids. Environ. Microbiol. 10:400-412. [PubMed]
40. Smidt, H., A. D. Akkermans, J. van der Oost, and W. M. de Vos. 2000. Halorespiring bacteria—molecular characterization and detection. Enzyme Microb. Technol. 27:812-820. [PubMed]
41. Sung, Y., K. M. Ritalahti, R. P. Apkarian, and F. E. Löffler. 2006. Quantitative PCR confirms purity of strain GT, a novel trichloroethene-to-ethene-respiring Dehalococcoides isolate. Appl. Environ. Microbiol. 72:1980-1987. [PMC free article] [PubMed]
42. Takeshita, T., Y. Nakano, and Y. Yamashita. 2007. Improved accuracy in terminal restriction fragment length polymorphism phylogenetic analysis using a novel internal size standard definition. Oral Microbiol. Immunol. 22:419-428. [PubMed]
43. Verta, M., S. Salo, M. Korhonen, T. Assmuth, H. Kiviranta, J. Koistinen, P. Ruokojarvi, P. Isosaari, P. A. Bergqvist, M. Tysklind, I. Cato, J. Vikelsoe, and M. M. Larsen. 2007. Dioxin concentrations in sediments of the Baltic Sea—a survey of existing data. Chemosphere 67:1762-1775. [PubMed]
44. Waller, A. S., R. Krajmalnik-Brown, F. E. Löffler, and E. A. Edwards. 2005. Multiple reductive-dehalogenase-homologous genes are simultaneously transcribed during dechlorination by Dehalococcoides-containing cultures. Appl. Environ. Microbiol. 71:8257-8264. [PMC free article] [PubMed]
45. Wilkinson, S. P., and A. Grove. 2006. Ligand-responsive transcriptional regulation by members of the MarR family of winged helix proteins. Curr. Issues Mol. Biol. 8:51-62. [PubMed]
46. Wu, Q., C. E. Milliken, G. P. Meier, J. E. Watts, K. R. Sowers, and H. D. May. 2002. Dechlorination of chlorobenzenes by a culture containing bacterium DF-1, a PCB dechlorinating microorganism. Environ. Sci. Technol. 36:3290-3294. [PubMed]

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