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Applied and Environmental Microbiology, May 2009, p. 2684-2693, Vol. 75, No. 9
0099-2240/09/$08.00+0     doi:10.1128/AEM.02037-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Characterization of a Dehalobacter Coculture That Dechlorinates 1,2-Dichloroethane to Ethene and Identification of the Putative Reductive Dehalogenase Gene{triangledown}

Ariel Grostern1 and Elizabeth A. Edwards2*

Department of Botany,1 Department of Chemical Engineering, University of Toronto, Toronto, Ontario, Canada2

Received 2 September 2008/ Accepted 25 February 2009


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ABSTRACT
 
Dehalobacter and "Dehalococcoides" spp. were previously shown to be involved in the biotransformation of 1,1,2-trichloroethane (1,1,2-TCA) and 1,2-dichloroethane (1,2-DCA) to ethene in a mixed anaerobic enrichment culture. Here we report the further enrichment and characterization of a Dehalobacter sp. from this mixed culture in coculture with an Acetobacterium sp. Through a series of serial transfers and dilutions with acetate, H2, and 1,2-DCA, a stable coculture of Acetobacterium and Dehalobacter spp. was obtained, where Dehalobacter grew during dechlorination. The isolated Acetobacterium strain did not dechlorinate 1,2-DCA. Quantitative PCR with specific primers showed that Dehalobacter cells did not grow in the absence of a chlorinated electron acceptor and that the growth yield with 1,2-DCA was 6.9 (±0.7) x 107 16S rRNA gene copies/µmol 1,2-DCA degraded. PCR with degenerate primers targeting reductive dehalogenase genes detected three distinct Dehalobacter/Desulfitobacterium-type sequences in the mixed-parent culture, but only one of these was present in the 1,2-DCA-H2 coculture. Reverse transcriptase PCR revealed the transcription of this dehalogenase gene specifically during the dechlorination of 1,2-DCA. The 1,2-DCA-H2 coculture could dechlorinate 1,2-DCA but not 1,1,2-TCA, nor could it dechlorinate chlorinated ethenes. As a collective, the genus Dehalobacter has been show to dechlorinate many diverse compounds, but individual species seem to each have a narrow substrate range.


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INTRODUCTION
 
The estimated annual global production of 1,2-dichloroethane (1,2-DCA) is 1.6 x 107 tons (17), the highest of all synthetic chlorinated compounds. It is also a common groundwater contaminant; in the United States, 1,2-DCA has been found in at least 287 of 932 National Priorities List sites identified by the U.S. Environmental Protection Agency (search of the National Priorities List database [http://www.epa.gov/superfund/sites/query/advquery.htm] on 31 July 2008). While this potential human carcinogen (1) can be degraded by aerobic microorganisms (27), anaerobic dechlorination is often a more relevant process because aquifers can be depleted of oxygen. Several mixed anaerobic cultures (5-7, 9, 32, 48, 53) have been developed and shown to degrade 1,2-DCA, usually by dechlorination to ethene but also by reductive dechlorination to chloroethane. Cometabolic degradation of 1,2-DCA has also been shown for methanogenic and acetogenic isolated microorganisms (4, 10, 15, 24).

The use of 1,2-DCA as a terminal electron acceptor to support growth in the process known as dehalorespiration has also been demonstrated. This was first reported by Maymó-Gatell et al. (35), who found that the tetrachloroethene (PCE)-degrading isolate "Dehalococcoides ethenogenes" strain 195 could grow by oxidizing H2 and reducing 1,2-DCA primarily to ethene (and <1% to vinyl chloride). Growth with 1,2-DCA was subsequently reported for two other isolates, "Dehalococcoides" sp. strain BAV1 (20) and "Desulfitobacterium dichloroeliminans" strain DCA1 (8). Recently, Marzorati et al. (34) identified reductive dehalogenase homologue genes (rdh genes) in a mixed culture and in D. dichloroeliminans strain DCA1, whose gene product may be involved in dehalorespiration with 1,2-DCA. Those authors hypothesized that these reductive dehalogenases (RDases) are specifically adapted to 1,2-DCA dechlorination (34).

Dehalorespiration of chlorinated ethanes was also demonstrated in a set of enrichment cultures derived from a contaminated site in West Louisiana (WL) (19). Two of these ethanol (EtOH)-amended cultures (fed 1,2-DCA or 1,1,2-trichloroethane [1,1,2-TCA] and hereafter called WL DCA/EtOH and WL TCA/EtOH, respectively) contain both Dehalobacter and Dehalococcoides spp. A third EtOH-amended culture that was simultaneously fed 1,2-DCA and 1,1,2-TCA (WL DCA/TCA/EtOH) lost the Dehalococcoides population and therefore accumulates the intermediate vinyl chloride (VC) from 1,1,2-TCA dihaloelimination. In the current study, we report for the first time the identification of a Dehalobacter strain that utilizes 1,2-DCA dihaloelimination for growth in coculture with a nondechlorinating Acetobacterium sp. as well as the identification of a putative 1,2-DCA reductive dehalogenase gene from this organism.


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MATERIALS AND METHODS
 
Enrichment and substrate range study.
A series of transfer cultures were initiated using the previously described EtOH- and 1,2-DCA-amended WL DCA/EtOH enrichment culture (see Table 2) (19) as an inoculum. Serum bottles (160 ml) with black butyl rubber stoppers and containing 100 ml of sterile anaerobic mineral medium (14) with 5 mM sodium acetate were inoculated with 0.1 ml of culture using sterile disposable syringes and needles, and the bottles were amended with 40 µM 1,2-DCA and 5 ml of H2-CO2 (80%/20%). Upon the complete conversion of 1,2-DCA to ethene, the transfer was repeated. After 10 transfers, a series of 1/10 dilutions was prepared in 25-ml crimp-top tubes (Bellco Glass, Inc.) with 10 ml of medium. Dechlorination was repeatedly observed at the 10–6 dilution. One milliliter of the 10–6 dilution was then used to inoculate serum bottles with 100 ml of anaerobic mineral medium (5 mM acetate) and repeatedly amended with 1,2-DCA and H2/CO2. These and subsequent transfers were incubated in an anaerobic glove box (Coy) at room temperature and are referred hereafter as WL DCA/H2.


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  		TABLE 2. Characteristics of WL culturesa

For substrate range experiments, 5 ml of WL DCA/H2 was diluted in 5 ml of fresh mineral medium with 5 mM sodium acetate in crimp-top tubes with butyl rubber stoppers. The test chlorinated compounds included 1,2-DCA, 1,1,2-TCA, PCE, trichloroethene (TCE), cis-dichloroethene (cDCE), and VC, which were added from saturated aqueous stocks or a gaseous stock (VC) to a concentration of 10 mg/liter (60 to 100 µM) and subsequently amended with 3 ml of H2-CO2. Uninoculated negative controls were also prepared.

16S rRNA gene cloning.
The UltraClean soil DNA kit (Mo Bio Laboratories, Inc., Solana Beach, CA) was used to extract total genomic DNA from 40 ml of culture pelleted at 6,600 x g for 20 min according to the manufacturer's alternative protocol for maximum yields. Bacterial 16S rRNA genes were selectively amplified from the purified DNA by PCR using the primer set 8f/1541r (Table 1). PCR was performed in triplicate 50-µl reaction mixtures containing 1x ThermoPol PCR buffer (New England Biolabs, Mississauga, Canada), 0.4 µM of each primer, 0.4 mM deoxynucleoside triphosphates (dNTPs), 1.5 U of Taq DNA polymerase, and 1 to 50 ng of DNA. The conditions used for PCR amplification were as follows: an initial denaturation step at 94°C for 5 min and then 25 cycles of denaturation at 94°C for 30 s, primer annealing at 52°C for 30 s, and chain extension for 1.5 min at 72°C, followed by a final extension step at 72°C for 10 min. A PTC-200 thermocycler (MJ Research, Inc., Waltham, MA) was used for PCR. Triplicate reaction mixtures were combined and cloned with the TOPO TA cloning kit (Invitrogen Corp., Carlsbad, CA) according to the manufacturer's protocol. Thirty-two clones were chosen to be sequenced with primers M13r and T7f, and the closest sequence match was then identified with the blastn utility of GenBank (http://blast.ncbi.nlm.nih.gov/Blast.cgi).


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  		TABLE 1. Primers used in this study

To test for the presence of archaea, the general archaeal primer pair 1Af/1100Ar (Table 1) was used to amplify genomic DNA from WL DCA/H2 using the above PCR protocol with an annealing temperature of 55°C and 35 cycles. To test for the presence of Dehalococcoides, the Dehalococcoides-specific primers Dhc1f and Dhc264r were used (Table 1).

Isolation of Acetobacterium and testing for dechlorination.
Shake tubes containing sodium formate (44 mM), methanol (0.04%, vol/vol), yeast extract (0.04%, wt/vol), and agar (1.5%) in anaerobic mineral medium were used to isolate Acetobacterium sp. cells from the WL DCA/H2 coculture. One milliliter of a 10-fold dilution series of WL DCA/H2 coculture in mineral medium was used to inoculate 9 ml of shake tube medium kept at 45°C. After solidification of the agar in a cool-water bath, 3 ml of H2-CO2 (80:20, vol/vol) was added to each tube with a gas-tight syringe, and the tubes were stored at room temperature. Following growth of colonies (~10 days), a single colony was picked with a syringe containing anaerobic medium and transferred into a serum bottle (160 ml) containing 50 ml of anaerobic mineral medium amended with sodium formate (44 mM), methanol (0.04%, vol/vol), and yeast extract (0.04%, wt/vol). The culture was grown statically at room temperature.

The ability of the Acetobacterium sp. isolate to dechlorinate 1,2-DCA was tested in serum bottles (160 ml) with mineral medium containing either (i) sodium formate (44 mM), methanol (0.04%, vol/vol), and yeast extract (0.04%, wt/vol); (ii) sodium formate (44 mM) and sodium acetate (5 mM); or (iii) sodium acetate (5 mM) and H2-CO2 (5 ml). Medium (45 ml) was inoculated with 5 ml of formate-methanol-yeast extract-grown cells, amended with 100 µM 1,2-DCA, and incubated at room temperature with shaking at 150 rpm.

Cloning of reductive dehalogenase genes.
rdh fragments were amplified from WL DCA/H2 genomic DNA using the degenerate PCR primer set RRF2/RD7r, ceRD2Sf/RD7r, or ceRD2Lf/RD7r (Table 1). PCR components were the same as those used for 16S rRNA gene cloning except that 0.16 µg/µl bovine serum albumin was added into each reaction mixture. The thermocycling conditions used for PCR amplification were similar to those used for 16S rRNA gene cloning. PCR products were cloned with the TOPO TA cloning kit, and positive clones were sequenced with the vector primers following restriction fragment length polymorphism (RFLP) selection. Sequences were assembled and aligned using Bioedit (http://www.mbio.ncsu.edu/BioEdit/BioEdit.html), and the sequences’ closest matches were identified with the tblastx utility of GenBank.

Two rounds of inverse PCR were used obtain the 5' and 3' flanking sequences for the partial rdh sequence obtained from the WL DCA/H2 culture. In the first round, 175 ng of genomic DNA was digested overnight in 50-µl reaction mixtures with Eco31I or NdeI restriction enzymes at 37°C. Self-ligation was performed by the addition of 5 U of T4 DNA ligase and 10x ligase buffer and incubation at room temperature overnight. Inverse PCR with primers WLInv1stL/WLInv1stR (Table 1) was then performed according to the PCR protocol described above except that the annealing temperature was 57°C and the extension step was performed for 3 min. PCR products were cloned and sequenced as described above. To obtain more of the 5' flanking region, a second round of inverse PCR was performed similarly but with the restriction enzymes BamHI, EcoRI, and NcoI; primers WLInv2ndL and WLInv2ndR (Table 1); and an annealing temperature of 56°C.

Time course experiments.
Growth of Dehalobacter and Acetobacterium spp. during the degradation of 1,2-DCA was monitored. Serum bottles (160 ml) were filled with 100 ml of anaerobic mineral medium containing 5 mM sodium acetate, closed with black butyl rubber stoppers, and autoclaved. Triplicate bottles were amended with neat 1,2-DCA (initial aqueous concentration of 0.5 mM) and 5 ml of H2-CO2.Triplicate bottles were amended with only H2-CO2 as no-electron-acceptor controls. The WL DCA/H2 culture was added at a 1% (vol/vol) inoculum, and 1.5 ml of culture was immediately removed for DNA extraction. DNA was subsequently extracted from 1.5-ml samples after 3, 4, and 5 days. DNA was extracted from no-electron-acceptor bottles in parallel to 1,2-DCA bottles. After the 1,2-DCA was completely dechlorinated, DNA was extracted from all bottles (day 5), the bottles were reamended with H2-CO2, and 1,2-DCA was added to a concentration of 1 mM. Following the second complete dechlorination, DNA was again extracted (day 13), the bottles were reamended with H2-CO2, and 1,2-DCA was added to a concentration of 2 mM. Final DNA extractions were performed on day 36.

For DNA extractions, the culture was transferred into microcentrifuge tubes (1.7 ml) and centrifuged at 16,000 x g for 10 min at 4°C. Most of the supernatant was removed, and the remaining 100 µl was used to resuspend and transfer the pellet into an UltraClean soil DNA kit bead-beating tube. DNA was then extracted according to the manufacturer's protocol except that the DNA was finally eluted with 5 mM Tris-Cl (pH 8.0). The copies of Dehalobacter and Acetobacterium 16S rRNA genes in the extracted DNA were then enumerated using quantitative PCR (qPCR) (see below).

qPCR.
qPCR for enumerating specific genes was conducted with 20-µl reaction mixtures containing 10 µl of SYBR green JumpStart Taq ReadyMix (Sigma), 7.2 µl of sterile water, 2 µl of DNA template, and 0.5 µM each of the forward and reverse primers. The thermocycling program was as follows: an initial denaturation step for 3 min at 94°C; 45 cycles of denaturation at 94°C for 20 s, annealing at 63°C for 20 s, extension at 72°C for 30 s, and fluorescence reading; and a final melting-curve analysis from 72°C to 95°C, measuring fluorescence every 0.5°C. Primer set Dhb477f/Dhb647r was used for the detection of Dehalobacter 16S rRNA genes, while primer set Aceto572f/Aceto791r was used for the detection of Acetobacterium 16S rRNA genes (Table 1). Specific primer sets for WL rdh genes were also designed and are listed in Table 1.

Calibration for the Acetobacterium qPCR was performed with serial dilutions of a known quantity of an M13r/T7f-amplified fragment of an Acetobacterium 16S rRNA gene-carrying plasmid generated through the cloning experiment described above. For Dehalobacter 16S rRNA gene and reductive dehalogenase gene homologue qPCRs, a vector carrying the 16S rRNA gene and three culture WL-derived rdh fragments was constructed (see below) (Fig. 1). The entire four-gene insert was amplified with M13r and T7f, and serial dilutions of this PCR product were used as the standard for qPCR, thereby ensuring accurate relative quantification. The dynamic range for qPCR for both targeted 16S rRNA genes was 2 x 103 to 4 x 108 16S rRNA gene copies/reaction. DNA concentrations were determined by UV absorbance measurement with a NanoDrop ND-1000 apparatus (NanoDrop Technologies, Wilmington, DE).


Figure 1
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  		FIG. 1. Schematic of assembly of the four-gene plasmid. The starting Dehalobacter 16S rRNA gene plasmid (1), intermediate three-gene plasmid (2), and final four-gene plasmid (3) are shown. Positions of genes, relevant restriction enzyme sites, and primer binding positions are relative to the pCR2.1 numbering convention. Kan, kanamycin resistance cassette.

Development of a four-gene qPCR calibration standard.
A pCR2.1 vector carrying the Dehalobacter 16S rRNA gene was used as the backbone for the new vector (Fig. 1). Note that all of the subsequent PCRs were performed in 50-µl reaction mixtures containing 1x HF or GC buffer (Finnzymes protocol), 200 µM dNTPs, 0.5 µM each primer, 1 U of Phusion DNA polymerase (Finnzyme Oy), and 1 µl of template. First, a 236-bp fragment of the WL rdhA2 gene was added simultaneously with a 352-bp fragment of the WL rdhA3 gene between the 16S rRNA fragment and the plasmid T7f sequence by using the unique NotI and XhoI restriction enzyme plasmid target sites. The WL rdhA2 gene fragment had a 5' NotI site and a 3' NcoI site engineered by PCR with primers WLrdhA2NotIf and WLrdhA2NcoIr, while the WL rdhA3 fragment had a 5' NcoI site and a 3' XhoI site engineered with primers WLrdhA3NcoIf and WLrdhA3XhoIr (Table 1). Next, a 484-bp fragment of WL rdhA1 was cloned between the plasmid M13r sequence and the 16S rRNA gene fragment using the unique SacI and BamHI sites. The WL rdhA1 fragment was amplified with primers WLrdhA1SacIf and WLrdhA1BamHIr (Table 1) to introduce a 5' SacI site and a 3' BamHI site. Following restriction digestion and ligation, the sequence of the four-gene plasmid was verified by DNA sequencing.

Reverse transcriptase PCR (RT-PCR).
The WL DCA/H2 culture was purged with N2-CO2 (80%/20%) to remove volatile compounds, and 40 ml was then transferred into two 120-ml serum bottles. The bottles were left for 5 days in the dark in an anaerobic glove box, and each bottle was then amended with 5 ml of H2-CO2. 1,2-DCA (0.43 mM) was also added to one of the bottles. After one-quarter of the 1,2-DCA had been degraded, 40 ml of each culture was centrifuged at 10,000 x g for 20 min at 4°C. RNA was then extracted and cDNA was generated according to methods described previously by Waller et al. (51).

To test for the transcription of WL rdhA1, PCR using primers WLrdhA1f and WLrdhA1r (Table 1) and cDNA generated as described above was performed in 50-µl reaction mixtures containing 1x ThermoPol PCR buffer, 0.4 µM of each primer, 0.4 mM dNTPs, 1.5 U of Taq DNA polymerase (New England Biolabs), and 1 to 2 µl of cDNA. The conditions used for PCR amplification were as follows: an initial denaturation step at 94°C for 5 min and then 35 cycles of denaturation 94°C for 20 s, primer annealing for 20 s at 63°C, and chain extension for 45 s at 72°C, followed by a final extension step at 72°C for 10 min. RT-PCR products were analyzed by gel electrophoresis. The cotranscription of rdhA1 and rdhB1 was tested by RT-PCR as described above except that primers WLrdhA1f and WLrdhB1r (Table 1) were used with an annealing temperature of 57°C and an extension time of 1.5 min. PCRs with non-reverse-transcribed RNA were run in parallel.

Analytical procedures.
For culture maintenance, time course, and substrate range experiments, chlorinated ethanes, ethenes, methane, and ethene were measured by injecting a 300-µl-headspace sample onto a Hewlett-Packard 5890 Series II gas chromatograph fitted with a GSQ column (J&W Scientific), as previously described by Grostern and Edwards (19) for chlorinated ethanes and Duhamel et al. (13) for chlorinated ethenes.

Nucleotide sequence accession numbers.
The cloned Acetobacterium 16S rRNA gene and rdh sequences identified in the WL cultures were deposited in GenBank with the following accession numbers: FJ157998 for Acetobacterium sp. strain WL, J010189 for WL rdhAB1, FJ010190 for WL rdhA2, and FJ010191 for WL rdhA3.


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RESULTS
 
Culture enrichment and characterization of coculture.
The parent WL DCA/EtOH mixed culture rapidly degrades 1,2-DCA stoichiometrically to ethene while also producing methane (Table 2). After 10 0.1% transfers of this culture into anaerobic medium containing H2 as an electron donor, acetate as a carbon source, and 1,2-DCA, methanogenesis ceased, but dechlorinating activity persisted at a rate of 150 µM/day. This was substantially higher than the dechlorination rate of 80 µM/day in the parent WL DCA/EtOH culture and was evidence of enrichment of a 1,2-DCA-dechlorinating organism.

The WL DCA/EtOH culture contains Dehalococcoides and Dehalobacter spp. that were previously shown to grow only in the presence of a chlorinated substrate (19). Growth experiments demonstrated that both species were likely to be involved in the degradation of 1,2-DCA (19). Given the strict electron donor (H2) and acceptor (a chlorinated compound) requirements of other species in these genera, enrichment with H2 and 1,2-DCA was expected to simultaneously select for both the Dehalococcoides and Dehalobacter strains in this culture. However, after 18 transfers, four dilution series, and a further transfer, the WL DCA/H2 culture was dominated microscopically by gram-positive short rods occurring in pairs or as single cells, similar to the morphology of other Dehalobacter sp., while no small circular cells typical of Dehalococcoides strains were visible. After several more feedings, a thin, long filament that was pink in Gram stains and stained with DAPI (4',6'-diamidino-2-phenylindole) could also be observed (Fig. 2).


Figure 2
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  		FIG. 2. DAPI-stained image of the WL DCA/H2 culture.

Cloning and sequencing of bacterial 16S rRNA genes from the WL DCA/H2 culture with general primers revealed the presence of only Dehalobacter (28 of 32 clones) and Acetobacterium (4 of 32 clones) spp. but no Dehalococcoides cells. PCR using WL DCA/H2 genomic DNA as a template did not produce a product with Dehalococcoides-specific primers (data not shown), confirming the loss of Dehalococcoides through transfers. Additionally, no PCR product was detected with general archaeal primers. On the basis of this information, the transfers were considered to be a coculture of Dehalobacter and Acetobacterium spp.

In order to eliminate the homoacetogenic Acetobacterium sp., several isolation methods were tried unsuccessfully. Roll tubes inoculated with serial dilutions of the coculture and amended with acetate, H2, and 1,2-DCA resulted in substantial ethene production from 1,2-DCA but no distinct colony formation. To exclude CO2 that is required by homoacetogens for growth, a CO2-free culture using HEPES buffer (instead of sodium bicarbonate) and an N2-purged headspace was tested, but no 1,2-DCA dechlorination was observed upon the addition of acetate and pure H2. Finally, a selenium- and tungsten-free trace-metals solution was tested, since it was through the exclusion of these metals that Dehalobacter restrictus was isolated from a contaminating homoacetogen (23). Four transfers in this medium resulted in a Dehalobacter-enriched population, but Acetobacterium remained present (data not shown).

An alternative to eliminating the Acetobacterium sp. is to isolate it from the Dehalobacter and then investigate its ability to dechlorinate 1,2-DCA. Using a medium containing formate, methanol, and yeast extract in shake tubes, distinct cream disc-shaped colonies that are 1 to 2 mm in diameter appeared in the 10–6-dilution tube after 8 days of incubation. Two subsequent shake-tube transfers ensured the purity of the strain, and its identity as an Acetobacterium strain was confirmed by sequencing of the 16S rRNA gene amplified from DNA from three colonies. The ability of the isolate to dechlorinate 1,2-DCA was tested in medium containing formate-methanol-yeast extract, formate-acetate, or acetate-H2-CO2. No 1,2-DCA depletion or ethene production was observed with any of the media tested after 15 days despite growth of the culture as monitored by the optical density at 595 nm (data not shown).

Correlation of Dehalobacter growth and 1,2-DCA dechlorination.
To verify the involvement of Dehalobacter in 1,2-DCA degradation, the WL DCA/H2 coculture was inoculated into fresh medium and amended with H2, acetate, and 1,2-DCA, and degradation was then monitored by headspace sampling, while growth was monitored by species-specific 16S rRNA gene qPCR of DNA samples extracted during degradation. qPCR samples were compared to donor-only controls that were not amended with 1,2-DCA. After a 2-day lag, the 100x-diluted culture degraded 25 µM 1,2-DCA to stoichiometric amounts of ethene in 5 days (Fig. 3A). Through refeeding with 50 µM and then with 100 µM 1,2-DCA, 1,2-DCA degradation remained relatively constant, with a dechlorination rate of 150 µM/day.


Figure 3
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  		FIG. 3. Dechlorination of 1,2-DCA by the WL DCA/H2 culture. (A) 1,2-DCA dechlorination profile with a 100x-diluted culture. Each curve shows the mean values of data from triplicate bottles, and error bars are the standard deviations. (B) Dehalobacter and Acetobacterium growth during 1,2-DCA degradation. Circles represent Dehalobacter, and triangles represent Acetobacterium. Open symbols represent controls (amended with H2 only); closed symbols represent bottles amended with H2 and 1,2-DCA. Gene copy values are mean values of data from triplicate bottles and are expressed as copies/ml of culture. Error bars are the standard deviations of data from triplicate bottles. Numbers on the right represent the net increase ("+") or decrease ("–") in gene copies/ml from the start to the end of the experiment for each target gene with each treatment.

In the presence of 1,2-DCA, Dehalobacter cultures grew by almost 25-fold to a density of 9.8 (±3.0) x 107 16S rRNA gene copies/ml (Fig. 3B). The perception of a lack of growth from days 14 to 36 stems from the inherent difficulty of using qPCR to detect small changes in already concentrated cell populations (19). In a culture at 1 x 107 copies/ml, a change of 1 x 107 copies/ml (equal to what was observed in the first 14 days) is not easily detectable. Based on the exponential growth phase from days 0 to 5 as depicted in Fig. 3B, a doubling time of 0.7 days was estimated. In the absence of 1,2-DCA, Dehalobacter concentrations decreased threefold, illustrating its requirement for the chlorinated electron acceptor for growth (Fig. 3B). For Acetobacterium, levels of growth were similar in both 1,2-DCA-amended and electron-donor-only bottles, illustrating that the growth of this organism is independent from 1,2-DCA. A yield of Dehalobacter from 1,2-DCA dechlorination was calculated, assuming that all 1,2-DCA was degraded by Dehalobacter. The yield of 6.9 (±0.7) x 107 16S rRNA genes copies/µmol 1,2-DCA degraded is only one-quarter that observed during the dechlorination of 1,1,2-TCA to VC (a process requiring equal electron equivalents) in the WL TCA/EtOH culture (19).

Dechlorination of 1,1,2-TCA, PCE, TCE, cDCE, and VC was tested with WL DCA/H2. In 5 weeks, no dechlorination products were detected for any of these potential substrates. It was previously established that the WL culture does not dechlorinate 1,1,1-TCA or 1,1-DCA (19).

Dehalogenase cloning and transcription.
Degenerate PCR was used to identify putative reductive dehalogenase homologues in the WL DCA/H2 culture. Primer sets ceRD2Sf/RD7r and RRF2/RD7r yielded positive PCR products, and after RFLP of 86 clones, 16 clones were sequenced. Of these, seven were identified by tblastx as being rdh genes by matches to rdh clones in GenBank, and alignment of these clones showed that they represent the same sequence. Two rounds of inverse PCR were used to increase the known sequence from 1,000 bp to 2,255 bp, allowing the complete WL rdhA1 and WL rdhB1 sequences to be obtained (Fig. 4). rdhA1 encodes a 551-amino-acid protein that has 95% similarity to PceA of Dehalobacter restrictus (33) and 92% similarity to the putative 1,2-DCA RDase of D. dichloroeliminans strain DCA1 (34). It also has TAT signal peptide and Fe-S cluster binding sequences identical to those of PceA and DcaA. rdhB1 encodes a 105-amino-acid putative membrane protein with 99% identity to PceB of D. restrictus.


Figure 4
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  		FIG. 4. Map of the 2,255-bp WL rdh-1 gene cluster. (A) Location of rdh genes in the 2,255-bp cloned region. TAT, twin-arginine translocation signal sequence; Fe-S, iron-sulfur cluster binding sequence. (B) Approximate position and direction of primers used for detection by qPCR or RT-PCR (see text and Table 1 for primers). (C) Approximate position and direction of primers used in two rounds of inverse PCR.

To examine the diversity of Dehalobacter-type rdh gene sequences in the WL cultures, primer set RRF2/RD7r was used to amplify rdh-like sequences from the more complex parent culture WL DCA/TCA/EtOH (Table 2) (19). Sixteen positive clones were analyzed by RFLP, seven clones were sequenced, and two distinct sequences were obtained: WL rdhA2 and WL rdhA3. WL rdhA1 was not detected by this method. These two new sequences were translated and compared with rdhA1 and found to have 74% and 96% identity, respectively, over 360 amino acids.

To determine the prevalence of the rdh gene sequences in various WL cultures, qPCR targeting of each rdh was used. The standard for qPCR was a construct containing the Dehalobacter strain WL 16S rRNA gene sequence as well as 236 to 484 bp representing each of the three identified rdh sequences (Fig. 1). Table 3 shows that rdhA1 was the only sequence detected in DNA from the WL DCA/H2 culture, whereas all three sequences were detected in the more complex WL DCA/EtOH and WL DCA/TCA/EtOH cultures although in differing rdh copies. None of the rdh genes were detected in DNA from the MS culture, a Dehalobacter-containing culture used as a control that degrades 1,1,1-TCA but not 1,2-DCA (18).


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  		TABLE 3. Occurrence of 16S rRNA and WL rdh copies in various Dehalobacter-containing cultures

The copy numbers for the rdh genes were never more than one-quarter of the copy numbers of Dehalobacter 16S rRNA genes detected by qPCR (Table 3). To determine if this was an artifact of extracting DNA from stationary cultures or if this represented a difference in the chromosomal copies of 16S rRNA genes and rdh genes, the samples from the above-described WL DCA/H2 culture growth experiment were reexamined for the presence of rdhA1. As shown in Fig. 5A, an increase in numbers rdhA1 gene copies during dechlorination correlated with an increase in Dehalobacter 16S rRNA gene copies; the ratio of the increase in the total copy number of rdhA1 to that of the 16S rRNA gene increase was 1:4. Also, in the donor-only controls, both rdhA1 and Dehalobacter 16S rRNA gene copy numbers decreased over time, although the ratio between the two templates was more variable.


Figure 5
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  		FIG. 5. Growth and transcription of WL rdh-1 during 1,2-DCA dechlorination. (A) Growth of Dehalobacter 16S rRNA and WL rdhA1 gene copies during dechlorination of 1,2-DCA. Circles represent Dehalobacter 16S rRNA genes, and squares represent WL rdhA1 genes. Open symbols represent controls (amended with H2 only); closed symbols represent bottles amended with H2 and 1,2-DCA. Gene copy values are mean values of data from triplicate bottles and are expressed as copies/ml of culture. Error bars are the standard deviations of data from triplicate bottles. Numbers on the right represent the net increase ("+") or decrease ("–") in gene copies/ml from the start to the end of the experiment for each target gene with each treatment. (B) Transcription of WL rdh-1 during dechlorination of 1,2-DCA. The left gel is the RT-PCR result using WL rdhA1-specific primers. The right gel is the RT-PCR result with primers for the detection of cotranscription of WL rdhA1 and rdhB1. For both gels, the first two lanes used cDNA as the PCR template (see text), the third lane used WL DCA/H2 genomic DNA as the template, and the fourth lane is the PCR water control. No PCR products were detected with no-RT culture RNA (data not shown). Images are a negative of ethidium bromide-stained agarose gels.

The actual transcription of rdhAB1 was tested by RT-PCR using RNA extracted from an active WL DCA/H2 culture. The culture was starved for 5 days and then amended with either 1,2-DCA and H2 or H2 only. After 17 h, RNA was extracted, and cDNA was generated and probed for transcription of rdh-1. Transcription of rdhA1 was observed only in the presence of 1,2-DCA (Fig. 5B). Additionally, by using primers that simultaneously amplify a portion of rdhA1 and rdhB1, it is evident that these two genes are cotranscribed. For both RT-PCRs, control reaction mixtures lacking reverse transcriptase did not produce a PCR product (data not shown).


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DISCUSSION
 
Dehalobacter dechlorinates 1,2-DCA.
In this work, the dechlorination of 1,2-DCA by a Dehalobacter sp. in the WL DCA/H2 coculture was confirmed because Dehalobacter cultures grew in the presence, but not in the absence, of 1,2-DCA. The dependence on a chlorinated substrate for growth is a trademark of only two known genera, Dehalobacter and Dehalococcoides, both of which were present in the parent WL DCA/EtOH culture (19). Marzorati et al. (34) recently described a 6VS culture, a 1,2-DCA-degrading enrichment culture that contained Desulfitobacterium and Dehalobacter spp. In the 6VS culture, Dehalobacter was found at about 3 x 106 16S rRNA gene copies/ng of total culture DNA, a concentration similar to that found for Dehalobacter strains in the various WL cultures (Table 3). Although Marzorati et al. (34) did not explicitly prove that 1,2-DCA was a growth substrate of the Dehalobacter strain, the organism became enriched after several transfers on 1,2-DCA. Therefore, it is clear that Dehalobacter can dechlorinate 1,2-DCA efficiently by dechlorination to ethene. Vinyl chloride, a by-product of 1,2-DCA dechlorination by D. ethenogenes strain 195 (35), was not detected in these Dehalobacter cultures. The 1,2-DCA-dependent growth of these Dehalobacter strains makes these organisms ideal candidates for use in the bioremediation of 1,2-DCA-contaminated sites.

The competition for electron donors among microorganisms is a well-known phenomenon in dechlorinating mixed cultures, where dechlorinators, methanogens, and acetogens vie for hydrogen use (11). Among dechlorinators, competition for electron acceptors can also occur, for example, between Geobacter and Dehalococcoides for TCE in the KB-1 culture (11, 12), between different strains of Dehalococcoides for chlorinated ethenes (47), or between Dehalococcoides and Dehalobacter for 1,2-DCA in WL DCA/EtOH (19). In the WL DCA/EtOH culture, the batch mode of maintenance and low rates of culture dilution allowed the competitors to coexist. However, under conditions of higher dilutions, differences in growth rate and cell yield become important in competition for the electron acceptor and for population maintenance. Previously, it was shown that when starved for 3 months, Dehalococcoides in the WL DCA/EtOH culture persisted at a higher concentration than did Dehalobacter (19); however, upon reamendment with 1,2-DCA, the Dehalobacter population rebounded at a much higher rate. It follows that deep dilutions and multiple serial transfers as carried out in this study would favor the faster-growing Dehalobacter over Dehalococcoides.

Acetobacterium persisted in the culture despite an aggressive enrichment process. The Acetobacterium strain was isolated and was found to be unable to dechlorinate 1,2-DCA, unlike the Acetobacterium sp. described previously by De Wildeman et al. (10). No dechlorination of 1,2-DCA was observed with the WL Acetobacterium isolate in any of three medium growth conditions tested, including the acetate-H2 medium used to grow the coculture, over a period of 2 weeks (whereas the coculture completely dechlorinated 1.5 mM of 1,2-DCA to ethene in the same time frame). Acetobacterium may simply compete with Dehalobacter for reducing equivalents (H2), and the batch style of culturing allows Acetobacterium to coexist with Dehalobacter, or Acetobacterium may provide some essential growth factors to Dehalobacter. The latter possibility was the hypothesis presented for the obligatory presence of Sedimentibacter in coculture with a β-hexachlorocyclohexane-dechlorinating Dehalobacter sp. (49). Another dechlorinating organism, D. dichloroeliminans strain DCA1, was initially enriched in coculture with Enterococcus casseliflavus (8); further work showed that the latter species produced a menaquinone essential for the growth of strain DCA1, and the addition of menaquinones led to the isolation of strain DCA1 (8). Acetobacterium spp. are known for vitamin B12 production (2, 3, 26, 45); however, the ability of Dehalobacter to synthesize or salvage corrinoids is unknown. More work is required to determine how the WL Acetobacterium culture is in fact assisting Dehalobacter, if at all.

Multiple Dehalobacter strains exist in the WL culture.
A decreased substrate range and the loss of rdh sequences following enrichment are evidence that, at minimum, two strains of Dehalobacter exist in the parent WL cultures. Substrate range experiments showed that the WL DCA/H2 culture had lost the parent WL culture's ability to degrade 1,1,2-TCA, PCE, TCE, cDCE, and VC. The loss of 1,1,2-TCA dechlorination activity is especially interesting given the past finding that Dehalobacter was responsible for the dechlorination of 1,1,2-TCA in the WL TCA/EtOH culture (19). Likely, the strain responsible for 1,1,2-TCA degradation is different from the 1,2-DCA-dechlorinating strain enriched here. The presence of three rdh sequences in the parent WL DCA/EtOH culture compared to only one in the enriched DCA/H2 culture also suggests the presence of at least two strains of Dehalobacter in the parent culture and that enrichment on H2 selected for one of those strains. rdhA1 was found to be a relatively minor proportion of Dehalobacter rdh genes in the WL DCA/TCA/EtOH culture (amended with both chloroethanes), whereas the gene constituted a much larger proportion in the WL DCA/EtOH culture (amended with only 1,2-DCA). This, coupled with the transcription of WL rdhA1 in the presence of 1,2-DCA, supports the hypothesis that rdhA1 represents a putative 1,2-DCA (and not 1,1,2-TCA) reductive dehalogenase gene. The occurrence of rdhA2 and rdhA3 in similar abundances in the 1,1,2-TCA-amended culture suggests that these sequences are present in a second Dehalobacter strain that uses 1,1,2-TCA and, possibly to a lesser extent, 1,2-DCA.

The quantitative comparison of rdh and 16S rRNA gene copy numbers to discriminate strains was previously reported for Dehalococcoides (25, 40, 47), but no such work for Dehalobacter- or Desulfitobacterium-containing cultures has been reported. This approach is a powerful tool to determine whether a population, as defined by a 16S rRNA gene sequence, is in fact composed of multiple strains. With qPCR, it is feasible, as demonstrated with Dehalococcoides (47) and in this report, to assess the population strain complexity in field samples and in enrichments and isolates, particularly those that are achieved through serial dilution and transfers rather than colony picking. Caution is warranted, however, because multiple copies of the 16S rRNA gene in a genome can exist (see below); a concatenated qPCR calibration standard including both ribosomal and functional genes ensures the accurate relative quantification essential for this approach.

Dehalobacter carries multiple rRNA operons.
qPCR of rdh genes also provided evidence that Dehalobacter contains multiple rRNA (rrn) operons. As of February 2009, no genomic data for any Dehalobacter strains exist despite the significant study of the biochemical properties of the Dehalobacter restrictus PceA reductive dehalogenase (33, 43, 44) and the apparent widespread presence of this genus at contaminated sites worldwide. Additionally, there have been no reports examining the link between cell counts and 16S rRNA gene copies, which is important if qPCR of Dehalobacter 16S rRNA gene copies is used to assess biostimulated or bioaugmented sites. The standard method for determining rrn operon copy number is Southern blotting (e.g., see reference 31), but PCR methods (e.g., see reference 30) have also been used (genome sequencing gives the clearest answer, of course). By simultaneously analyzing changes in 16S rRNA gene copies and RDase gene copies in a growing culture, we found that there is a near-constant ratio of 4 16S rRNA gene copies:1 rdhA1 gene copy. Whereas all four sequenced Dehalococcoides strains have one rrn operon in their genome, two organisms that are more phylogenetically related to Dehalobacter, Desulfitobacterium hafniense Y51 and Desulfitobacterium hafniense DCB-2, have six (36) and five (search of rrnDB) (28) rrn operons, respectively. Therefore, the presence of four rrn operons in the Dehalobacter strain in the DCA/H2 culture is quite feasible. It would be useful to do this kind of analysis with other Dehalobacter strains to determine if there is a consistent rrn operon copy number across Dehalobacter strains.

A novel putative Dehalobacter 1,2-DCA reductive dehalogenase gene was identified.
This work identified a 1,2-DCA reductive dehalogenase gene from a Dehalobacter strain. A putative 1,2-DCA rdh operon was cloned, its abundance was shown to correlate with Dehalobacter growth and dechlorination of 1,2-DCA, and it was transcribed specifically upon exposure to 1,2-DCA. WL rdhA1 is distinct from the putative 1,2-DCA rdh recently identified by Marzorati et al. in their 6VS enrichment (34). Although the 6VS enrichment culture was reported to contain both Dehalobacter and Desulfitobacterium spp. (34), it is not clear which organism harbored the cloned RDase gene. However, an rdh sequence with high similarity (98% at the amino acid level) to the 6VS culture sequence was cloned from a pure culture of Desulfitobacterium dichloroeliminans DCA1, indicating that the 6VS putative 1,2-DCA rdh was most likely from Desulfitobacterium rather than Dehalobacter. The three novel sequences (WL rdhA1, rdhA2, and rdhA3) are the only Dehalobacter reductive dehalogenase homologues that have been reported in the literature, aside from those obtained from D. restrictus (33, 38). Given the involvement of Dehalobacter in the transformation of a number of different chlorinated compounds, including chlorinated ethanes (19, 41) and ethenes (22, 52), polychlorinated biphenyls (37), β-hexachlorocyclohexane (49), trichlorobenzenes (50), and 1,2-dichloropropane (42), there are few rdh genes identified from this genus. Dehalobacter strains are obligate dechlorinators; therefore, reductive dehalogenases necessarily play a key role in the organism's metabolism. In the absence of genomic information, study of how strains differ in their complements of rdh genes can give insight into strain relatedness and substrate range potential. A larger effort is required to understand the physiology and diversity of this widespread and environmentally important dechlorinating genus.

In summary, we have identified a Dehalobacter sp. that dechlorinates 1,2-DCA to ethene and have discovered a likely 1,2-DCA reductive dehalogenase gene. This work adds a body of research describing different Dehalobacter strains and their substrate ranges. Dehalobacter restrictus (22) and strain TEA (52) dechlorinate only PCE and TCE, while Dehalobacter sp. strains TCA1 (46) and MS (18) dechlorinate only 1,1,1-TCA and 1,1,-DCA but not other chlorinated ethanes or ethenes. Similarly, Dehalobacter sp. strain E1-E3 dechlorinates β-hexachlorocyclohexane but not chlorinated ethanes or ethenes (49). The Dehalobacter strain described herein was found to dechlorinate only 1,2-DCA. These findings are in contrast to previous observations on individual Dehalococcoides strains that tend to dechlorinate chlorinated ethanes, ethenes, propenes, and aromatic compounds. Thus, Dehalobacter strains collectively dechlorinate a broad spectrum of substrates, while the substrate range of individual strains appears to be quite restricted.


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ACKNOWLEDGMENTS
 
We thank Winnie Chan for help in clone library construction, Cheryl Devine for help in the Acetobacterium isolation, and Melanie Duhamel and Cheryl Devine for constructive comments on the manuscript.

The research was funded by U.S. Department of Defense Strategic Environmental Research and Development Program (SERDP). A. Grostern was funded by a Natural Science and Engineering Research Council Canada graduate scholarship.


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FOOTNOTES
 
* Corresponding author. Mailing address: University of Toronto, 200 College St., Toronto, ON M5S 3E5, Canada. Phone: (416) 946-3506. Fax: (416) 978-8605. E-mail: elizabeth.edwards{at}utoronto.ca Back

{triangledown} Published ahead of print on 6 March 2009. Back


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REFERENCES
 
    1
  1. Agency for Toxic Substances and Disease Registry. 2004. ToxFAQs for 1,2-dichloroethane. Centers for Disease Control and Prevention, Atlanta, GA. http://www.atsdr.cdc.gov/tfacts38.html.
    2. 2
  2. Bainotti, A. E., B. Estebanez, H. Nagadomi, and N. Nishio. 2000. Production of vitamin B12 in an upflow anaerobic filter continuous reactor using Acetobacterium sp. Biotechnol. Lett. 22:503-508.[CrossRef]
    3. 3
  3. Bainotti, A. E., K. Yamaguchi, Y. Nakashimada, and N. Nishio. 1998. Kinetics and energetics of Acetobacterium sp. in chemostat culture on methanol-CO2. J. Ferment. Bioeng. 85:223-229.[CrossRef]
    4. 4
  4. Belay, N., and L. Daniels. 1987. Production of ethane, ethylene, and acetylene from halogenated hydrocarbons by methanogenic bacteria. Appl. Environ. Microbiol. 53:1604-1610.[Abstract/Free Full Text]
    5. 5
  5. Bouwer, E. J., and P. L. McCarty. 1983. Transformations of 1- and 2-carbon halogenated aliphatic organic compounds under methanogenic conditions. Appl. Environ. Microbiol. 45:1286-1294.[Abstract/Free Full Text]
    6. 6
  6. Chang, Y. C., M. Hatsu, K. Jung, Y. S. Yoo, and K. Takamizawa. 1998. Degradation of a variety of halogenated aliphatic compounds by an anaerobic mixed culture. J. Ferment. Bioeng. 86:410-412.[CrossRef]
    7. 7
  7. Chen, C., J. A. Puhakka, and J. F. Ferguson. 1996. Transformations of 1,1,2,2-tetrachloroethane under methanogenic conditions. Environ. Sci. Technol. 30:542-547.
    8. 8
  8. De Wildeman, S., G. Diekert, H. Van Langenhove, and W. Verstraete. 2003. Stereoselective microbial dehalorespiration with vicinal dichlorinated alkanes. Appl. Environ. Microbiol. 69:5643-5647.[Abstract/Free Full Text]
    9. 9
  9. De Wildeman, S., N. Hendrik, H. Van Langenhove, and W. Verstraete. 2001. Reductive biodegradation of 1,2-dichloroethane by methanogenic granular sludge in lab-scale UASB reactors. Adv. Environ. Res. 6:17-27.[CrossRef]
    10. 10
  10. De Wildeman, S., A. Neumann, G. Diekert, and W. Verstraete. 2003. Growth-substrate dependent dechlorination of 1,2-dichloroethane by a homoacetogenic bacterium. Biodegradation 14:241-247.[CrossRef][Medline]
    11. 11
  11. Duhamel, M., and E. A. Edwards. 2007. Growth and yields of dechlorinators, acetogens, and methanogens during reductive dechlorination of chlorinated ethenes and dihaloelimination of 1,2-dichloroethane. Environ. Sci. Technol. 41:2303-2310.[Medline]
    12. 12
  12. Duhamel, M., and E. A. Edwards. 2006. Microbial composition of chlorinated ethene-degrading cultures dominated by Dehalococcoides. FEMS Microbiol. Ecol. 58:538-549.[CrossRef][Medline]
    13. 13
  13. Duhamel, M., S. Wehr, L. Yu, H. Rizvi, D. Seepersad, S. Dworatzek, E. E. Cox, and E. A. Edwards. 2002. Comparison of anaerobic dechlorinating enrichment cultures maintained on tetrachloroethene, trichloroethene, cis-1,2-dichloroethene and vinyl chloride. Water Res. 36:4193-4202.[Medline]
    14. 14
  14. Edwards, E. A., and D. Grbic-Galic. 1994. Anaerobic degradation of toluene and o-xylene by a methanogenic consortium. Appl. Environ. Microbiol. 60:313-322.[Abstract/Free Full Text]
    15. 15
  15. Egli, C., R. Scholtz, A. M. Cook, and T. Leisinger. 1987. Anaerobic dechlorination of tetrachloromethane and 1,2-dichloroethane to degradable products by pure cultures of Desulfobacterium sp. and Methanobacterium sp. FEMS Microbiol. Lett. 43:257-261.[CrossRef]
    16. 16
  16. Embley, T. M., B. J. Finlay, R. H. Thomas, and P. L. Dyal. 1992. The use of rRNA sequences and fluorescent probes to investigate the phylogenetic positions of the anaerobic ciliate Metopus palaeformis and its archaeobacterial endosymbiont. J. Gen. Microbiol. 138:1479-1487.[Abstract/Free Full Text]
    17. 17
  17. Field, J. A., and R. Sierra-Alvarez. 2004. Biodegradability of chlorinated solvents and related chlorinated aliphatic compounds. Rev. Environ. Sci. Biotechnol. 3:185-254.[CrossRef]
    18. 18
  18. Grostern, A., and E. A. Edwards. 2006. A 1,1,1-trichloroethane-degrading anaerobic mixed microbial culture enhances biotransformation of mixtures of chlorinated ethenes and ethanes. Appl. Environ. Microbiol. 72:7849-7856.[Abstract/Free Full Text]
    19. 19
  19. Grostern, A., and E. A. Edwards. 2006. Growth of Dehalobacter and Dehalococcoides spp. during degradation of chlorinated ethanes. Appl. Environ. Microbiol. 72:428-436.[Abstract/Free Full Text]
    20. 20
  20. He, J., K. M. Ritalahti, K. 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.[CrossRef]
    21. 21
  21. Hendrickson, E. R., J. Payne, R. M. Young, M. G. Starr, M. P. Perry, S. Fahnestock, D. E. Ellis, and R. C. Ebersole. 2002. Molecular analysis of Dehalococcoides 16S ribosomal DNA from chloroethene-contaminated sites throughout North America and Europe. Appl. Environ. Microbiol. 68:485-495.[Abstract/Free Full Text]
    22. 22
  22. Holliger, C., D. Hahn, H. Harmsen, W. Ludwig, W. Schumacher, B. Tindall, F. Vazquez, N. Weiss, and A. J. B. Zehnder. 1998. Dehalobacter restrictus gen. nov. and sp. nov., a strictly anaerobic bacterium that reductively dechlorinates tetra- and trichloroethene in an anaerobic respiration. Arch. Microbiol. 169:313-321.[CrossRef][Medline]
    23. 23
  23. Holliger, C., G. Schraa, A. J. M. Stams, and A. J. B. Zehnder. 1993. A highly purified enrichment culture couples the reductive dechlorination of tetrachloroethene to growth. Appl. Environ. Microbiol. 59:2991-2997.[Abstract/Free Full Text]
    24. 24
  24. Holliger, C., G. Schraa, A. J. M. Stams, and A. J. B. Zehnder. 1990. Reductive dechlorination of 1,2-dichloroethane and chloroethane by cell suspensions of methanogenic bacteria. Biodegradation 1:253-261.[CrossRef][Medline]
    25. 25
  25. Holmes, V. F., J. He, P. K. H. Lee, and L. Alvarez-Cohen. 2006. Discrimination of multiple Dehalococcoides strains in a trichloroethene enrichment by quantification of their reductive dehalogenase genes. Appl. Environ. Microbiol. 72:5877-5883.[Abstract/Free Full Text]
    26. 26
  26. Inoue, K., S. Kageyama, K. Miki, T. Morinaga, Y. Kamagata, K. Nakamura, and E. Mikami. 1992. Vitamin B12 production by Acetobacterium sp. and its tetrachloromethane-resistant mutants. J. Ferment. Bioeng. 73:76-78.[CrossRef]
    27. 27
  27. Janssen, D. B., and W. De Koning. 1995. Development and application of bacterial cultures for the removal of chlorinated aliphatics. Water Sci. Technol. 31:237-247.
    28. 28
  28. Klappenbach, J. A., P. R. Saxman, J. R. Cole, and T. M. Schmidt. 2001. Rrndb: the ribosomal RNA operon copy number database. Nucleic Acids Res. 29:181-184.[Abstract/Free Full Text]
    29. 29
  29. Krajmalnik-Brown, R., T. Holscher, 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.[Abstract/Free Full Text]
    30. 30
  30. Lee, C., S. Lee, S. G. Shin, and S. Hwang. 2008. Real-time PCR determination of rRNA gene copy number: absolute and relative quantification assays with Escherichia coli. Appl. Microbiol. Biotechnol. 78:371-376.[CrossRef][Medline]
    31. 31
  31. Linton, D., M. Moreno, R. J. Owen, and J. Stanley. 1992. 16S rrn gene copy number in Helicobacter pylori and its application to molecular typing. J. Appl. Bacteriol. 73:501-506.[Medline]
    32. 32
  32. Löffler, F. E., J. E. Champine, K. M. Ritalahti, S. J. Sprague, and J. M. Tiedje. 1997. Complete reductive dechlorination of 1,2-dichloropropane by anaerobic bacteria. Appl. Environ. Microbiol. 63:2870-2875.[Abstract]
    33. 33
  33. Maillard, J., W. Schumacher, F. Vazquez, C. Regeard, W. R. Hagen, and C. Holliger. 2003. Characterization of the corrinoid iron-sulfur protein tetrachloroethene reductive dehalogenase of Dehalobacter restrictus. Appl. Environ. Microbiol. 69:4628-4638.[Abstract/Free Full Text]
    34. 34
  34. Marzorati, M., F. de Ferra, H. Van Raemdonck, S. Borin, E. Allifranchini, G. Carpani, L. Serbolisca, W. Verstraete, N. Boon, and D. Daffonchio. 2007. A novel reductive dehalogenase, identified in a contaminated groundwater enrichment culture and in Desulfitobacterium dichloroeliminans strain DCA1, is linked to dehalogenation of 1,2-dichloroethane. Appl. Environ. Microbiol. 73:2990-2999.[Abstract/Free Full Text]
    35. 35
  35. Maymó-Gatell, X., T. Anguish, and S. H. Zinder. 1999. Reductive dechlorination of chlorinated ethenes and 1,2-dichloroethane by "Dehalococcoides ethenogenes" 195. Appl. Environ. Microbiol. 65:3108-3113.[Abstract/Free Full Text]
    36. 36
  36. Nonaka, H., G. Keresztes, Y. Shinoda, Y. Ikenaga, M. Abe, K. Naito, K. Inatomi, K. Furukawa, M. Inui, and H. Yukawa. 2006. Complete genome sequence of the dehalorespiring bacterium Desulfitobacterium hafniense Y51 and comparison with Dehalococcoides ethenogenes 195. J. Bacteriol. 188:2262-2274.[Abstract/Free Full Text]
    37. 37
  37. Oh, K. H., E. B. Ostrofsky, and Y. C. Cho. 2008. Molecular characterization of polychlorinated biphenyl-dechlorinating populations in contaminated sediments. J. Microbiol. 46:165-173.[CrossRef][Medline]
    38. 38
  38. 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.[CrossRef][Medline]
    39. 39
  39. Reysenbach, A. L., G. S. Wickham, and N. R. Pace. 1994. Phylogenetic analysis of the hyperthermophilic pink filament community in Octopus Spring, Yellowstone National Park. Appl. Environ. Microbiol. 60:2113-2119.[Abstract/Free Full Text]
    40. 40
  40. Ritalahti, K. M., B. K. Amos, Y. Sung, Q. Z. Wu, S. S. Koenigsberg, and F. E. Löffler. 2006. Quantitative PCR targeting 16S rRNA and reductive dehalogenase genes simultaneously monitors multiple Dehalococcoides strains. Appl. Environ. Microbiol. 72:2765-2774.[Abstract/Free Full Text]
    41. 41
  41. Rossetti, S., F. Aulenta, M. Majone, G. Crocetti, and V. Tandoi. 2008. Structure analysis and performance of a microbial community from a contaminated aquifer involved in the complete reductive dechlorination of 1,1,2,2-tetrachloroethane to ethene. Biotechnol. Bioeng. 100:240-249.[CrossRef][Medline]
    42. 42
  42. Schlotelburg, C., C. von Wintzingerode, R. Hauck, F. von Wintzingerode, W. Hegemann, and U. B. Gobel. 2002. Microbial structure of an anaerobic bioreactor population that continuously dechlorinates 1,2-dichloropropane. FEMS Microbiol. Ecol. 39:229-237.[Medline]
    43. 43
  43. Schumacher, W., and C. Holliger. 1996. The proton/electron ration of the menaquinone-dependent electron transport from dihydrogen to tetrachloroethene in "Dehalobacter restrictus." J. Bacteriol. 178:2328-2333.[Abstract/Free Full Text]
    44. 44
  44. Schumacher, W., C. Holliger, A. J. B. Zehnder, and W. R. Hagen. 1997. Redox chemistry of cobalamin and iron-sulfur cofactors in the tetrachloroethene reductase of Dehalobacter restrictus. FEBS Lett. 409:421-425.[CrossRef][Medline]
    45. 45
  45. Stupperich, E., H. J. Eisinger, and B. Krautler. 1988. Diversity of corrinoids in acetogenic bacteria. P-cresolylcobamide from Sporomusa ovata, 5-methoxy-6-methylbenzimidazolylcobamide from Clostridium formicoaceticum and vitamin B12 from Acetobacterium woodii. Eur. J. Biochem. 172:459-464.[Medline]
    46. 46
  46. Sun, B., B. N. Griffin, H. L. Ayala-del-Rio, S. A. Hashsham, and J. M. Tiedje. 2002. Microbial dehalorespiration with 1,1,1-trichloroethane. Science 298:1023-1025.[Abstract/Free Full Text]
    47. 47
  47. 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.[Abstract/Free Full Text]
    48. 48
  48. Tandoi, V., T. D. DiStefano, P. A. Bowser, J. M. Gossett, and S. H. Zinder. 1994. Reductive dehalogenation of chlorinated ethenes and halogenated ethanes by a high-rate anaerobic enrichment culture. Environ. Sci. Technol. 28:973-979.
    49. 49
  49. van Doesburg, W., M. H. A. van Eekert, P. J. M. Middeldorp, M. Balk, G. Schraa, and A. J. M. Stams. 2005. Reductive dechlorination of β-hexachlorocyclohexane (β-HCH) by a Dehalobacter species in coculture with a Sedimentibacter sp. FEMS Microbiol. Ecol. 54:87-95.[CrossRef][Medline]
    50. 50
  50. von Wintzingerode, F., B. Selent, W. Hegemann, and U. B. Gobel. 1999. Phylogenetic analysis of an anaerobic, trichlorobenzene-transforming microbial consortium. Appl. Environ. Microbiol. 65:283-286.[Abstract/Free Full Text]
    51. 51
  51. 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.[Abstract/Free Full Text]
    52. 52
  52. Wild, A., R. Hermann, and T. Leisinger. 1996. Isolation of an anaerobic bacterium which reductively dechlorinates tetrachloroethene and trichloroethene. Biodegradation 7:507-511.[CrossRef][Medline]
    53. 53
  53. Wild, A. P., W. Winkelbauer, and T. Leisinger. 1995. Anaerobic dechlorination of trichloroethene, tetrachloroethene and 1,2-dichloroethane by an acetogenic mixed culture in a fixed-bed reactor. Biodegradation 6:309-318.[CrossRef][Medline]

Applied and Environmental Microbiology, May 2009, p. 2684-2693, Vol. 75, No. 9
0099-2240/09/$08.00+0     doi:10.1128/AEM.02037-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.




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