Characterization of a Microbial Consortium Capable of Rapid and Simultaneous Dechlorination of 1,1,2

Jan 11, 03:21 AM

Current Headlines: By Jones, Elizabeth J P; Voytek, Mary A; Lorah, Michelle M; Kirshtein, Julie D

ABSTRACT

Mixed cultures capable of dechlorinating chlorinated ethanes and ethenes were enriched from contaminated wetland sediment at Aberdeen Proving Ground (APG) Maryland. The "West Branch Consortium" (WBC-2) was capable of degrading 1,1,2,2-tetrachloroethane (TeCA), trichloroethene (TCE), cis and trans 1,2-dichloroethene (DCE), 1,1,2- trichloroethane (TCA), 1,2-dichloroethane, and vinyl chloride to nonchlorinated end products ethene and ethane. WBC-2 dechlorinated TeCA, TCA, and cisDCE rapidly and simultaneously. A Clostridium sp. phylogenetically closely related to an uncultured member of a TCE- degrading consortium was numerically dominant in the WBC-2 clone library after 11 months of enrichment in culture. Clostridiales, including Acetobacteria, comprised 65% of the bacterial clones in WBC-2, with Bacteroides (14%), and epsilon Proteobacteria (14%) also numerically important. Methanogens identified in the consortium were members of the class Methanomicrobia, which includes acetoclastic methanogens. Dehalococcoides did not become dominant in the culture, although it was present at about 1% in the microbial population. The WBC-2 consortium provides opportunities for the in situ bioremediation of sites contaminated with mixtures of chlorinated ethenes and ethanes.

KEYWORDS anaerobic dechlorination, bioaugmentation of chlorinated ethanes, bioremediation, microbial consortium, mixed chlorinated ethenes and ethanes, 1,1,2,2-tetrachloroethane

INTRODUCTION

Bioaugmentation, site inoculation with a microbial mixed culture, is a proven approach for stimulating complete dechlorination of sites contaminated with chlorinated ethenes (e.g., Major et al., 2002; Lendvay et al., 2003). However, cultures have not previously been available for the large-scale treatment of chlorinated ethane contamination. Of additional concern, chlorinated ethanes can inhibit the degradation of chlorinated ethenes (Duhamel et al., 2002; Aulenta et al., 2005), and cultures are needed for remediation of sites with mixtures of these contaminants. Contamination of groundwater with chlorinated ethenes and ethanes is a serious problem due to widespread and historic commercial, industrial, and military use, relative resistance to degradation, and associated health hazards (Haggblom and Blossert, 2003). Under anaerobic conditions, chlorinated ethenes and ethanes can be partially reduced to less chlorinated compounds or completely degraded to nonchlorinated end products depending on the physiological capability of the indigenous microbial community (Ellis et al., 2000). Based on estimates of nonatmospheric release in the U.S. during the period 1987 to 1991 (http://www.epa.gov/safewater/ index.html; Chen et al., 1996), chlorinated ethanes comprise about 75% of the more than one million pounds of chlorinated ethenes and ethanes annually released to the environment. Yet relative to what has been learned regarding chlorinated-ethene degradation, the study of microorganisms that catalyze chlorinated ethane degradation is in its infancy. Isolates capable of reducing 1,2-dichloroethane (DCA) and 1,1,1-trichloroethane have been identified (Maym-Gatell et al., 1999; De Wildeman et al., 2003; Sun et al., 2002), and one isolate has been shown to reduce 1,1,2,2-tetrachloroethane (TeCA) to cis 1,2- dichloroethene (cisDCE) (Suyama et al., 2001). Recent research by Grostern and Edwards (2006) on a mixed culture demonstrated growth of Debalobacter sp. concomitant with the reduction of 1,1,2- trichloroethane (TCA) to vinyl chloride (VC).

TeCA was developed as a solvent prior to World War I, and large quantities are still used, primarily by the chemical industry. In addition, historic disposal practices and release of DNAPLs (dense non-aqueous phase liquids) have led to continuing groundwater contamination from subsurface sources. Natural attenuation of TeCA has been documented at Aberdeen Proving Ground (APG) Maryland, where contaminated groundwater discharges through anoxic wetland sediments at West Branch Canal Creek (Lorah and Olsen, 1999a, 1999b). This sediment provided source material for developing a dechlorinating culture. The TeCA degradation pathway (Figure 1) is primarily biotic, and includes both hydrogenolysis to less chlorinated ethanes and dichloroelimination to less chlorinated ethenes. Abiotic production of trichloroethene (TCE) from dehydrochlorination of TeCA generally accounts for less than 2% of TeCA removal at APG (Lorah and Olsen, 1999a), and chloroethane (CA), ethene and ethane have not been observed in the sediment Several intermediates of TeCA dechlorination are possible carcinogens and are listed as contaminants of concern by the U.S. Environmental Protection Agency. VC is a known human carcinogen that often accumulates at sites where dechlorination is slow or incomplete.

The purpose of the work described here was to develop a culture of microorganisms for bioaugmentation treatment of chlorinated- ethane contaminated groundwater at sites where dechlorination is incomplete or rates are too slow for effective remediation. In this paper, we describe a microbial consortium, West Branch Consortium (WBC-2), derived from organic-rich sediments collected in the wetland of West Branch Canal Creek (MD) at APG. We demonstrate the capability of WBC-2 to dechlorinate chlorinated ethanes and ethenes in culture, both individually and concurrently, and we present a preliminary analysis of its microbial composition.

MATERIALS AND METHODS

Development of the Microbial Consortium

Sediment was collected from two sites within the wetland at West Branch Canal Creek (WB23 and WB30), and prepared anaerobically using the same methods as for microcosms in previous studies (Lorah and Olsen, 1999a; Lorah et al., 2003b; Lorah and Voytek, 2004). APG sediments were collected in March 2003, sieved, slurried with groundwater (1:1.5), and incubated (19C) with TeCA (7 M) in 1-L serum bottles without headspace for 1 month. Most of the Fe(III) and sulfate (alternative electron acceptors) were depleted during this incubation period and methane was being produced. Aliquots (100 ml) of sediment slurries were then transferred to 120-ml serum bottles with a N^sub 2^/CO2 (95:5) headspace and amended with a daughter compound, cisDCE or TCA, for 1 to 2 months. In all sediment slurry enrichments, the electron donors were derived only from organic matter in the sediment. Sediment slurries (100 ml each of TCA- enriched WB23 and cisDCE-enriched WB30) were then transferred into anaerobic culture medium (1800 ml, see composition below) with sulfide (50 M) added as a reductant, and amended with target concentrations of TeCA (30 M) or a mixture of TeCA (25 M), TCA (50 M), and cisOCE (50 M). The electron donor for cultivation was selected in tests on the TeCA amended culture (see below) 3 weeks after inoculation from sediment slurry. Cultures were diluted over a 2-year period and contain about 0.1% sediment by volume.

Evaluation of Electron Donors

All electron donor tests were performed in duplicate on sub samples removed from TeCA-depleted stock culture. Because concentrations of intermediates often are low or undetectable during TeCA degradation by WBC2, TCA and cisVDCE were used as test compounds to ensure that the electron donor selected would support both chlorinated ethane and chlorinated ethene pathways. In addition, the ability of each electron donor to support the dechlorination of VC, a key compound for complete dechlorination of both chlorinated-ethenes and -ethanes, was evaluated. Aliquots of WBC-2 culture (10 ml) were transferred anaerobically to 28-ml pressure tubes (Bellco Glass, Vineland, NJ) filled with N2/CO2 (80:20). WBC-2 was evaluated for dechlorination of test compounds in the following electron donor treatments: propionate (10 mM); succinate (3 mM); lactate (3 mM); pyruvate (3 mM); benzoate (3 mM); formate (10 mM); acetate (10 mM); H2 (20 kPa overpressure, added three times during the incubation) with or without acetate (1 mM) added as a carbon source; whey (Sigma Chemical; from bovine milk, spray dried powder containing minimum 11% protein and approximately 65% lactose) (5 g/L); no electron donor added. The electron donors supplied electron equivalents (assuming complete oxidation) equal to about 100 times that required for the reduction of the chlorinated compounds, cisDCE or TCA (Supelco, Bellefonte, PA), which were added from aqueous emulsions (for a final concentration of approximately 1 and 0.75 mM, respectively) and monitored for VC and DCA production. The ability to dechlorinate VC was tested by adding VC (4.2 M) from a gaseous standard (Matheson, Twinsburg, OH) in a separate treatment. All treatments were incubated at 19C and monitored by sampling the headspace for analysis with a gas chromatograph (GC) with a flame ionization detector (FID), as described below.

Culture Medium and Maintenance

The anaerobic medium included (g/L deionized water): NaHCO3 (2.5), NHCl (0.5), NaPO (0.5), KC1 (0.1), 10 ml vitamin solution (Balch et al., 1979), and 10 ml trac\e mineral solution (below), with a gas phase of N2 and CO2 (80:20). The trace mineral solution contained (g/L): Nitrilotriacetic acid (1.5), MgSO 4-7H2O (3.0), MnSO 4-H 2O (0.5), NaCl (1.0), FeSO4-7H2O (0.1), CaCl2-2H2O (0.1), CoCl2-6H2O (0.1), ZnCl2 (0.13), CuSO4-5H2O (0.01), A1K(SO4)2-12H2O (0.01), H3BO3(0.01), Na2MoO4 (0.025), NiCl2-6H2O (0.024), Na 2WO4- 2H2O (0.025). Early in development, some batches were starved for periods as long as several months, but recovered activity when feeding was resumed. Once established, cultures were maintained either with lactate (1 mM) and TeCA (50 M) added from aqueous stocks once or twice weekly, or lactate (1.5 mM) and TeCA (25 M), TCA (50 M), and cisDCE (50 M). The ratio of electron equivalents for donors to acceptors was 30 and 17, respectively, for stocks maintained with TeCA and the chlorinated mixture. The chlorinated stocks were prepared by adding purified standards (Supelco, Bellefonte, PA) to sterile anaerobic deionized water. Chlorinated stock bottles were vigorously shaken prior to each amendment in order to emulsify any undissolved compound. The cisDCE contained approximately 1% transDCE.

WBC-2 was maintained in 2-L batches of medium from which culture volumes were removed for study. After 1 year, all cultures were restored to full 2 L volume by adding fresh medium. Under contract by the U.S. Army, samples of WBC-2 were given to SiREM Laboratories (Guelph, Ontario, Canada) for propagation of the culture to the large volumes required for field bioremediation tests conducted by the U.S. Geological Survey, using lactate and the chlorinated mixture described above (GeoSyntec Consultants Inc., 2004). WBC-2 microbial composition (clone analysis, see below) was assessed in a culture sample obtained from SiREM Laboratories that had been scaled up by transferring two times into fresh medium.

Gas Analysis

Culture headspace was sampled using a gas tight syringe and injected into one or both of the following GC-FID systems. For rapid analysis of TeCA, TCA, DCA, TCE, cisDCE, transDCE, and VC, we used a Hewlett-Packard model 5890 series II with isothermal separation at 100C on a VOCOL (Supelco, Bellefonte, PA) capillary column (30 m 0.53 mm). For separation of methane, ethene, and ethane, and for analysis of VC, transDCE, and cisDCE when interfering peaks were present, we used a Shimadzu model GC17A with separation on a Rt Q- Plot (Restek, Bellefonte, PA) column (30 m 0.32 mm) using a temperature program of 100C for 5 min, ramping to 200C at 20C/min. Aqueous standards of chlorinated compounds were prepared from highly purified neat calibration standards (Supelco, Bellefonte, PA). Standards of chlorinated compounds were prepared by adding 10 L of neat solution to 100 ml water, and preparing aqueous dilutions for headspace analysis in bottles sealed with Teflon coated stoppers (West, Lionville, PA). Dimensionless Henry's law constants (DHLCs), were used to calculate expected headspace concentrations from known liquid concentrations. Methane, ethene, and ethane standards were purchased as gas standards (Scott Specialty Gas). Concentrations of chlorinated compounds and non-chlorinated end products in samples of WBC-2 culture headspace were converted to dissolved values using DHLCs and total concentrations (per volume medium) were calculated. DHLCs have been measured empirically by many researchers, and vary widely. The chosen DHLCs fall in the midrange of published values. Nonetheless, the DHLCs are the greatest source of possible error in the concentrations reported here, and may exceed 10%. Errors between repeat injections are about 2%. The dimensionless Henry's law constants (DHLC) applied were 0.019 for TeCA, 0.556 for TCA, 0.1821 for DCA, 0.3056 for TCE, 0.1255 for mDCE, 0.3056 for transDCE, 0.9087 for VC (Gossett, 1987), 7.96 for ethene, 19.88 for ethane, and 28.5 for methane (Lampron et al., 1998). All detection limits were less than 0.01 M.

Cloning, RFLP Screening and Sequencing, TRFLP

DNA was extracted using the Bio-101 Fast DNA Spin Kit for Soil (MP Biomedicals, Irvine, CA) following manufacturers instructions, except that product recovery was maximized at each step. Bacterial and methanogen DNA were amplified using the polymerase chain reaction (PCR) in a Perkin Elmer Geneamp 2400 thermal cycler with 16S rDNA (46f and 519r) primers (Brunket al., 1996; Lane, 1991) and methyl coenzymeM reductase (mcrAf and mcrAr) primers (Luton et al., 2002), respectively. 16S rDNA PCR conditions (30 cycles) were denaturing at 94C (30 s), annealing at 56C (30 s), and extension at 720C (1 min). For mar, the conditions of Luton et al. (2002) were used. Microbial members of the consortium were characterized by cloning and sequencing the bacterial 16S rDNA and mcrA. amplicons. Amplicons were purified using the Wizard PCR purification kit (Promega, Madison, WI) and cloned using the TA cloning kit or the Topo TA cloning kit for sequencing according to manufacturer's instructions (Invitrogen, San Diego, CA). Colonies were picked and 16S rRNA and mcrA gene clone fragments (133 and 48, respectively) were recovered using vector primers and mcrA primers, respectively, using PCR. For the bacterial 16S rDNA characterization, the PCR products were reamplified using 46f and 519r primers. All PCR amplicons were digested with restriction enzymes (6 1 of PCR product with 2.5 U each of Mspl and HinPI) according to manufacturer's instructions (Promega, Madison, WI). Restriction fragments were analyzed by size separation on a 3.5% Metaphor (Cambrex, Rockland, ME) agarose gel, restriction fragment length polymorphism (RFLP) patterns were distinguished, and the frequency with which each pattern occurred was determined. It should be noted that the frequency of clones in the library may not correspond directly to relative phylotype numbers in the culture due to undefined differences in the number of 16S rDNA copies per cell. In addition, PCR and nucleic extraction biases may contribute to apparent differences in the abundance of RFLP patterns. Representative clones for each pattern were selected for sequencing. Amplicons to be sequenced were purified with the Wizard PCR purification system, and cycle sequencing was performed on both strands using Big Dye v3.1 (Applied Biosystems, Foster City, CA) and run on a ABI310 genetic analyzer. Sequences were edited and assembled using Autoassembler (Applied Biosystems, Foster City, CA). Closest phylogenetic relatives were determined by BLASTn search of the National Center of Bioinformatics (NCBI) database (http://www.ncbi.nlm. nih.gov/).

Terminal restriction fragment length polymorphism (TRFLP)-PCR was performed as described above, but using 46f primer with FAM label attached. A restriction digest of 6 l of PCR product was performed using 5U MnlI (New England Biolabs, Beverly, MA). Digested samples were precipitated with 0.1 volume of 3 M sodium acetate and 2 volumes of cold 100% ethanol and resuspended in 10 1 sterile water. A 2.5-1 aliquot of the digested sample was added to 12 1 of deionized formamide and 0.5 1 ROX500 standard (Applied Biosystems). Samples were denatured at 95C for 5 min. DNA fragments were separated using an ABI310 sequencer (Applied Biosystem). Terminal restriction fragments were detected using 310Genescan analytical software, version 2.1.1, resulting in a TRFLP profile for each sample.

Detection and Quantification of Specific Members by Quantitative PCR

Primers were used to detect organisms with abundances too low to be detected in the 16S clone library, including two known dechlorinators, Dehahcoccoides spp., andDesulfuromonas spp., and methanogens. DNA copy number in an extract of WBC-2 DNA was determined by quantitative PCR (qPCR) using the quantitect SYBR green real-time PCR kit (Qiagen, Chatsworth, CA.) and the Opticon real-time PCR system (MJ Research, now BioRad, Hercules, CA). The 16S rDNA based primers used to target Dehalococcoides were dhc730f, 5'-GCG GTT TTC TAG GTT GTC-3' and dhcl350r, 5'-CAC CTT GCT GAT ATG CGG-3' (Bunge et al., 2001). The Desulfuromonas primers (designed for specificity to Desulfuromonas sp. strain BBl and D. chloroethenica 16S rDNA) and conditions are previously described (Lffler et al., 2000). Methanogens were quantified using mcrA primers (Luton et al., 2002). A standard curve was determined using Ct values of serial dilutions of plasmid containing the dhc or mcrA amplified fragment, or the Desulfuromonas sp. strain BBl amplicon of known concentration (and thus copy number), and the samples were plotted against that curve to determine abundance. Calculations of cell numbers were based on one 16S rDNA copy per cell for Dehalococcoides (www.tigr.org), and 1 mcrA copy per cell for methanogens (Nunoura et al., 2006). For the purposes of calculating cell numbers, nucleic acid extractions were assumed to be perfect, because no measurement of extraction efficiency is available. For microscopic counts, culture samples were suspended in 0.01% Triton X- 100 and stained with 5 g/ml 4',6-diamidino-2-phenylindole (DAPI) in 1 phosphate-buffered saline (PBS), filtered onto a black Nuclepore filter (0.2 m), and viewed using epi-fluorescence (Porter and Feig, 1980).

RESULTS

Electron Donors Supporting WBC-2 Dechlorination

The electron donor for WBC-2 cultivation was selected by comparing dechlorination in electron donor treatments (summarized in Table 1) with dechlorination in a control with no added electron donor. Controls with no added electron donor exhibited decreases in added TCA and cisDCE characteristic of adsorption, with an initial decrease of 24% and 10%, respectively, followed by no further decrease. Less than 1% of the added TCA or cisDCE in the controls was reduced to VC and no DCA was produced. H2 did not stimulate reduction of TCA or cisDCE above that observed in the controls, whether or not acetate was added as a carbon source. The most complete \dechlorination was obtained in treatments with lactate and pyruvate. The pathway of TCA reduction (production of VC versus DCA from TCA) varied among electron donor treatments. Cultures with lactate produced more DCA relative to VC than did cultures with pyruvate. No DCA was produced in treatments with propionate, acetate, benzoate, or whey.

Dechlorination in Culture

WBC-2 was cultivated in batches amended with either lactate and TeCA or lactate and a mixture of TeCA, TCA, and cisDCE for 18 months. In the TeCA amended culture, WBC-2 in a 2-L batch culture completely dechlorinated TeCA (measured at 240 ) within 2 days (Figure 2A). The pathway of TeCA degradation could not be discerned by monitoring this culture because very little intermediate accumulation occurred. Less than 0.5 M (0.2% of the added TeCA) accumulated as VC, and this VC was degraded by day 2. The end products of dechlorination were ethene and ethane. The stoichiometry of TeCA degraded by WBC-2 to nonchlorinated end products (average of values from three stock cultures) was 2 1 moles TeCA to 1 mole [ethene + ethane]. In a 2-L batch culture maintained with the chlorinated mixture, TeCA, TCA, and cisDCE, WBC-2 rapidly, simultaneously, and completely reduced all three chlorinated compounds to the nonchlorinated end-products ethene and ethane (Figure 2B). Small amounts of transDCE and VC were observed as transient intermediates.

After dechlorination was completed, the fate of ethene was monitored in the cultures in order to determine if ethene reduction could account for the production of ethane. Ethene was degraded (6 day^sup -1^) with the production of ethane (Figure 3). After ethene was depleted, the ethane concentration also decreased, at a rate of 0.5 M day^sup -1^. In another set of treatments (data not shown), ethene (336 13 M) was added to the culture to achieve a higher starting ethene concentration. This ethene was completely degraded after 18 days with the production of ethane (188 40 M). The lack of stoichiometric accumulation of ethane suggests that ethane and ethene degradation can co-occur.

Pathways of intermediate degradation were determined by incubating subsamples of WBC-2 with TCA or cisDCE, WBC-2 cultures that had been maintained with (1) TeCA and (2) the chlorinated mixture (TeCA, TCA, cisDCE) were incubated to deplete chlorinated compounds and then compared with respect to their capabilities to degrade TCA (Figure 4A) and cisDCE (Figure 45). The two cultures were very similar in their abilities to degrade the two TeCA intermediates. Degradation products of intermediate dechlorination are shown for TeCA-maintained WBC-2 as a representative culture (Figure 5). As in treatments amended with TeCA, little intermediate accumulation was observed in TCA-amended treatments (Figure 5A). The rate of TCA degradation was 36 M day^sup -1^. The peak VC and DCA concentrations measured were 1.7% and 0.3% of the TCA added, respectively, and both intermediates were rapidly degraded. Chloroethane was not detected. cisDCE reduction was accompanied by the production of VC (peak accumulation, 24% of added cisDCE), ethene and ethane (Figure 5B). The rate of cis;DCE degradation in cultures amended with cisDCE was 54 M day^sup -1^.

When WBC-2 cultures were diluted (1:9) into fresh medium, dechlorination was initially slow enough to allow the observation of intermediates (Figure 6A, B). Diluted cultures amended with TeCA alone (84 M) accumulated measurable TCA (0.01 M) at one time point, and transDCE (6 M) and VC (0.25 M) were also both produced and degraded. Diluted cultures amended with TeCA, TCA, and cisDCE had a transient accumulation of VC (6% of the added chlorinated compounds) and transDCE (as much as 4% of added TeCA), and also the abiotic product, TCE (3% of the TeCA parent added). Within 2 to 4 weeks, dechlorination rates increased to a level comparable to that observed in undiluted cultures, with parent compounds degraded in as little as one day and little transient accumulation of intermediates (Figure 6 A, B).

Microbial Composition of the Dechlorinating Consortium

The bacterial community in the source sediments (APG sediments WB23 and WB30, Figure 7A and B) had shifted after a year under culture conditions with lactate as the sole electron donor and TeCA or a mixture of TeCA, TCA, and cisDCE as the electron acceptors. Both WBC-2 cultures (cultured with TeCA [Figure 7 C] or TeCA, TCA, and cisDCE [Figure 7D]) exhibited TRFLP profiles that were different from that of the source sediments and overall represented a different community than was present in the starting materials. This change reflects the selection pressures exerted on the community and individual members by the chlorinated compounds, such that the remaining peaks represent members tolerant of the chlorinated compounds and favored by the culturing conditions and perhaps directly or indirectly involved in the degradation process. The numerically dominant phylogenetic types in WBC-2 were identified by cloning and sequencing 16S rDNA and mcrA genes from a culture grown with a mixture of TeCA, TCA, and cisDCE. The frequency of phylotype occurrence in 16S rDNA and mcrA clone libraries was determined (Figure 8A and B), and phylogenetic placement was determined using a BLAST search for related sequences. Although most of the WBC-2 clones were not related to dechlorinating bacteria that have been studied in isolation, many were related to bacterial clones that have been observed at other dechlorinating sites (Table 2). The 16S rDNA library was dominated by Clostridiales (65%), including three phylotypes. The phylotype representing the greatest number of clones was a Clostridium sp., most closely related (99%) to an uncultured member of a TCE dechlorinating community (MacBeth et al., 2004). The second most prevalent phylotype was an Acetobacterium p. most closely related (97%) to uncultured clones from a 1,2- dichloropropane-dechlorinating enrichment (Ritalahti and Lffler, 2004), and 97% and 96% related to the homoacetogens Acetobactmum malicum and A. wieringae, respectively. Less prevalent was a third phylotype, 95% related to an uncultured clone from a TCE- dechlorinating community and 93% related to Dehalobacter restrictus, in the evaluated region between 46f and 519r. There was more variability among sequences of the Bacteroidetes (CFB group), which accounted for 14% bacterial clones. Many of these were related to uncultured clones from dechlorinating populations (see examples, Table 2). One clone (delta Proteobacteria) was most closely related (98%) to a Geobacter sp. from a chlorinated ethene enrichment culture, and 98% related to G. lovleyi, a recently described PCE- dechlorinating isolate (Sung et al., 2006). The gamma Proteobacteria were 99% related to the top 60 BLAST hits, including Pseudomonas stutzeri and Ps. chloridismutans, that is able to dechlorinate trichloroacetic acid (Wolterink et al., 2002). For the other phylotypes observed among the bacterial clones (Arcobacter sp. and Desulfobulbus sp., 14% and 2% of the clones, respectively) and all of the mcrA clones, the BLAST database did not reveal relatedness to organisms from dechlorinating populations.

No Debalococcoides clones were identified. However, Dehalococcoides numbers determined independently using qPCR and compared with the total number of cells by microscopic count indicated that about 1% of the total consortium population was comprised of Debalococcoides spp. Microscopic examination confirmed that cocci were rare, and the consortium population was composed almost entirely of rod-shaped cells. Absolute numbers of Dehalococcoides (evaluated by qPCR) in WBC-2 culture samples (generally 10^sup 5^ to 10^sup 6^ cells per ml) were higher than numbers measured in APG sediment (10^sup 3^ to 10^sup 4^ cells per ml), although the efficiency of nucleic acid extraction and the total microbial numbers are both likely lower for sediment than for culture. Desulfuromonas spp. were not detected by qPCR using the primers specific for that dechlorinating type.

WBC-2 mcrA clone library (Figure 85) was comprised of members of the class Methanomicrobia, and included both acetate- and H^sub 2^- utilizing methanogens (accession numbers DQ907209 to DQ907221). Members of the Methanosarcinaceae family are capable of utilizing acetate in the production of methane (Sowers, 1995). These include Metbanosarcina spp., which may utilize acetate, H^sub 2^, methanol, or methyl amines, and Methanosaeta spp., which are obligate acetate utilizers. The other Methanomicrobia are related to methanogens that utilize H^sub 2^ and formate as electron donors. The presence of Methanosaeta spp. in the WBC-2 culture indicates that acetate is being produced. Although the cultures were methanogenic, methanogens comprised a very small part of the total microbial population of WBC- 2. Total methanogens quantified using qPCR comprised 0.2% of the total WBC-2 microbial population.

DISCUSSION

WBC-2 TeCA Dechlorination: Pathway and Rates

In order to develop an effective mixed culture for bioremediation of TeCA contaminated sites, it is critical to support the microorganisms catalyzing all branches of the TeCA degradation pathway. Most of the intermediates (with the exception of chloroethane) shown in Figure 1 have been observed to accumulate in APG groundwater during TeCA degradation (Lorah et al., 2003b). Therefore the remediation of TeCA contaminated groundwater could require remediation of the chlorinated intermediates as well as mixtures of the parent with chlorinated ethene and ethane intermediates or cocontaminants. WBC-2 was enriched from APG sediments in which intermediates accumulated and were slowly degraded at rates of 0.1 to 0.6 M day^sup -1^ for TeCA, TCA, and cisDCE (Lorah et al., 2003a, 2003b; Jones et al., 2004). After microbial enrichment and 1 year in culture, the rate of WBC-2 dechlorination inc\reased (rates measured for TeCA, TCA, and dsDCE removal: 100, 36, and 54 M day^sup -1^ respectively) such that almost no intermediates were detected in WBC-2 cultures amended with TeCA or a mixture of TeCA, TCA, and cisDCE. TeCA (as much as 240 M) was converted to ethene and ethane within 2 days, a rate of dechlorination greater than that observed for another mixed culture reported to reduce TeCA (6 M day^sup -1^; Aulenta et al., 2005). Ten- times dilutions of WBC-2 culture in fresh medium resulted in the transient accumulation of small amounts of TCA and transDCE from TeCA degradation, but even in the diluted culture, 87% was converted "directly" to ethene and ethane without observation of intermediates. The rate of dechlorination in the diluted cultures increased to the same rate as the parent culture in about 2 weeks, indicating microbial growth. Although intermediates often did not accumulate, the ability of WBC-2 to degrade all known intermediates in the TeCA pathway was demonstrated in treatments with individual compounds (e.g., TCA and cisVDCE) and in mixtures (e.g., TeCA, TCA, cisDCE, and cis/trans DCE, data not shown).

We used a simple method for monitoring dechlorination, based on partitioning in the headspace. However, the stoichiometry of TeCA to its degradation products is difficult to determine using this method. Both adding and measuring the initial concentration of TeCA involved potentially large errors. We could not rely on dilution of the stock solution to determine the amount of TeCA added, because TeCA in the stock solution was not completely dissolved and possibly not completely homogenized by shaking. TeCA measurements were subject to two known sources of error. First, when calculating total concentrations from headspace values, errors are magnified for compounds with a very low DHLC. The DHLC for TeCA (0.019) was an order of magnitude lower than any other compound measured and therefore subject to the greatest error. Conversely, compounds such as methane, ethene and ethane are largely partitioned to the headspace and subject to the least error. Secondly, because there is some sediment as well as cell mass in the cultures, distribution of TeCA between the liquid and gas phases may be complicated by adsorption, resulting in a discontinuity between TeCA uptake and the appearance of end product. All of these errors may have contributed to differences between target TeCA additions and measured concentrations (e.g., target addition of 60 M versus measured concentration, 240 M, that is reported in Figure 2), as well as the error in stoichiometry between different bottles (i.e., 2 1). In spite of the large errors, it is evident that ethene and ethane are major products of TeCA degradation. The reduction of ethene to ethane demonstrated by WBC-2 during and after the depletion of chlorinated compounds and in ethene-amended culture offers an explanation for the low recovery of nonchlorinated end products (2:1 substrate: product). The lack of ethene and ethane accumulation is a common field observation during dechlorination at APG and other contaminated field sites. This fact has confounded the interpretation of field data with respect to the degradation mechanism and fate of less chlorinated compounds. One explanation for this, supported by the data presented here, is the degradation of nonchlorinated end products preventing their accumulation, such that ethene and ethane do not accumulate because their rate of removal exceeds their rate of production under most field conditions. Characterizing the consortium, WBC-2, has provided an opportunity to learn more about the TeCA degradation pathway and helped explain the lack of accumulation of end products at field sites, as a result of changes in the rates between degradation steps and in the microbial community associated with culturing and enrichment.

The reduction of ethene could account for the ethane observed during TeCA reduction. However, another possible source of ethane is directly through the hydrogenolysis of chlorinated ethanes (left half of Figure 1). 1,1,1-Trichloroethane was degraded to chloroethane and finally to ethane by an isolate related to Dehdobacter restrictus (Sun et al., 2002). However, at present there is no direct evidence for this pathway in WBC-2. Chloroethane, the immediate precursor for hydrogenolytic ethane, was never observed, indicating either that it was not produced, or that rapid degradation prevented its accumulation. Further research will be needed to definitively determine the source(s) of ethane in the culture during chlorinated ethane reduction.

Many individual reductive dechlorination reactions have been documented to occur in pure or mixed cultures of microorganisms. The unique contribution of WBC-2 for contaminant treatment is the ability of the mixed culture to handle a variety of compounds (both chlorinated ethenes and ethanes) without noticeable inhibition. The dechlorination rates observed in WBC-2 for chlorinated ethenes are similar to those observed in cultures used for bioremediation treatment of chlorinated ethene contamination (e.g., KB-1; Duhamel et al., 2002). The advantage offered by WBC-2 is its ability to degrade chlorinated ethenes and ethanes simultaneously. Although, not shown here, WBC-2 was also able to reduce tetrachloroethene (PCE) in the presence of TeCA (Jones et al., unpublished data). WBC- 2 may or may not contain dechlorinating organisms similar to those studied in pure or mixed cultures with chlorinated ethenes.

Evidence for a Distinct Dechlorinating Population

The TeCA dechlorination pathway includes reactions that are a part of the more thoroughly studied PCE-dechlorination pathway (see right half of Figure 1), and thus might be expected to support the growth of similar organisms. Indeed, the predominance of Clostridia and CFB in the WBC-2 population (Table 2) is also characteristic of TCE-dechlorinating communities (MacBeth et al., 2004, Richardson et al., 2002), and organisms similar to the ethene-dechlorinating Debedococcoides and Dehalobacter are observed in WBC-2. However, the dechlorinating abilities of WBC-2, enriched in the presence of chlorinated ethanes, are different than cultures enriched with chlorinated ethenes, both with respect to the response to added electron donors and in the relative importance of Dehalococcoides within the microbial population. Based on limited studies (this paper and Aulenta et al., 2005), the electron donor needs for chlorinated ethane-enriched cultures appear to be different from those of cultures enriched for chlorinated ethene reduction. Most bacterial isolates capable of reductive dechlorination of chlorinated ethenes, including Dehalococcoides and Dehalobacter, use H^sub 2^ as the preferred electron donor. Sun et al. (2002) showed that a Dehalobacter isolate required H^sub 2^ plus acetate to reduce 1,1,1-trichloroethane. Furthermore, He et al. (2002) suggested that a volatile fatty acid (VFA) such as propionate can provide a slow release of H^sub 2^ to support hydrogenotrophic dechlorinators at the expense of methanogenesis. The chlorinated ethane-enriched culture, WBC-2, was not stimulated to reduce cisDCE or TCA with H^sub 2^, H^sub 2^ plus acetate, or propionate added as the electron donor. The failure of H^sub 2^ to stimulate dechlorination suggests that the organisms involved are not the same as the chlorinated ethene-reducing organisms that have been studied in isolation. Aulenta et al. (2005) found that butyrate, a VFA that supported a PCE-dechlorinating culture, did not do so in the presence of TeCA. TeCA apparently inhibited organisms that could release H^sub 2^ from butyrate. Thus, another mechanism by which chlorinated ethanes may inhibit chlorinated ethene-enriched populations is through the inhibition of key consortium members that may be indirectly involved in dechlorination.

Dehalococcoides spp., which include the only isolated bacteria identified to completely biodegrade chlorinated ethenes, have been targeted as an indicator of dechlorinating capability (Hendrickson et al., 2002). A number of field studies support the idea that complete degradation of chlorinated alkenes is dependent upon the presence of specific microorganisms from the genus Dehalococcoides (e.g., Hendrickson et al., 2002; Lowe et al., 2002). Several laboratory mixed cultures enriched for chlorinated ethene reduction are reported to have populations of Dehalococcoides spp. representing more than 30% of the total bacteria (Gu et al., 2004; Richardson et al., 2002; Duhamel et al., 2004), and it is widely believed that Dehalococcoides is key to the dechlorination in these cultures. WBC-2 dechlorinated chlorinated ethenes at rates similar to these other laboratory mixed cultures (e.g., 54 m day^sup - 1^cisDCE versus approximately 30 M day^sup -1^ for KB-1 [Duhamel et al., 2002]). However, Dehabcoccoides spp. comprise only a minor part (about 1%) of the cell population in WBC-2. This, coupled with the presence of consortium members that appear to be closely related to clones of unknown function in other dechlorinating populations, suggests that organisms other than Dehalococcoides spp.may play a greater role in TeCA dechlorination.

These observations suggest that exposure to chlorinated ethanes results in the selection of a different population of organisms for chlorinated ethene reduction. Cultures that were enriched using chlorinated ethenes (i.e., in the absence of chlorinated ethanes) were inhibited in the presence of chlorinated ethanes (Duhamel et al., 2002; Aulenta et al., 2005). WBC-2 was able to degrade chlorinated ethanes and ethenes simultaneously with little VC accumulation. This capability makes the microbial consortiumWBC-2 a potentially valuable tool for bioremediation of sites contaminated with mixtures of chlorinated ethenes and ethanes. In addition, the simultaneous reduction of all componen\ts of the TeCA degradation pathway can reduce the total treatment time and help prevent transport of hazardous compounds out of the treatment zone.

Identifying Microbial Roles

Although we have identified numerically important components of the microbial consortium WBC-2, the specific roles of consortium members have yet to be determined. Although tentative roles could be assigned to Dehalococcoides (cisDCE, transDCE, VC, and DCA reduction), Dehalobacter (TCA reduction to VC, DCA reduction to ethene), and Acetobactmum (DCA reduction to ethene) based on studies of related organisms Puhamel et al., 2002; He et al., 2003; Grostern and Edwards, 2006; De Wildeman et al., 2003), WBC-2 exhibited some capabilites, such as TeCA reduction to transDCE, and TCA reduction to DCA, for which no organisms have been implicated. In addition, indirect roles, such as satisfying the "undefined nutritional needs" of dechlorinating organisms, may be critical to the consortium function. Additional research on WBC-2 is planned to identify the roles of individual consortium members.

The observation of closely related phylotypes in WBC-2 and other dechlorinating communities, such as Clostridium sp., Acetobacterium sp., and CFB provides some evidence for the involvement of previously unrecognized bacteria in dechlorination processes. However, enrichment of organisms in a dechlorinating system provides only circumstantial evidence for direct involvement. In WBC-2, lactate fermentation, homoacetogenesis, methanogenesis, sulfur cycling, syntrophy, and chemoautotrophy could support organisms in the culture without deriving energy from dechlorination. Thus, organisms could persist in mixed culture without playing a direct role in dechlorination, and further work would be needed to confirm whether or not each plays a significant role. For example, Acetobacteria are able to grow by converting H^sub 2^ + CO2 to acetate, but they may also be directly or indirectly involved in dechlorination. An Acetobacterium sp. has been isolated that can cometabolically reduce DCA to ethene (De Wildeman et al., 2003). This strain lost its ability to dechlorinate after about 10 transfers probably due to undefined nutritional requirements. Although the 16S rDNA of the dechlorinating Acetobacterium strain was > 99% related to A. wieringae, the type strain was unable to dechlorinate DCA (De Wildeman et al., 2003), illustrating the limitation of phylogenic identification alone. Acetobacteria could also support dechlorination indirectly, for example through the production of corrinoid factors, which play a role in some dechlorinating reactions (Magnuson et al., 1998; Hlscher et al., 2004), or in syntrophic relationships, through the production of acetate (He et al., 2002).

The possible role of methanogens in dechlorination by WBC-2 is not known. Although the presence and activity of methanogens are often considered to be inhibitory to or at least contraindicative of dechlorination activity, some evidence suggests that methanogens may play an important role in dechlorination. Dechlorination is associated with methanogenic environments (Vogel and McCarty, 1985) but is often thought to be a cometabolic process that is less efficient than dehalorespiration. Bromoethane sulfonic acid, a potent inhibitor of methanogenesis, inhibited VC degradation in APG sediments (Lorah et al., 2003b). Work with pure cultures of Metbanosarcina sp. and Methanosarcina mazei has shown that methanogens can dechlorinate PCE to TCE, and dechlorination occurs only during active methane production (Fathepure and Boyd, 1988). Methanogens also produce corrinoid factors and unique enzymes and vitamins that may support dechlorination. For example, extracellular factors produced by Methanosarcina have been shown to enhance dechlorination of carbon tetrachloride (Novak et al., 1998). Anaerobic methane oxidation ("reverse methanogenesis" carried out by methanogens) has been coupled to sulfate reduction (Boetius et al., 2000) and denitrification (Raghoebarsing et al., 2006). Although not yet shown to be coupled to dechlorination, such a reaction is theoretically possible. Production of methane by WBC-2 could also stimulate co-metabolic dechlorination by methane oxidizers in aerobic environments (Chang and Alvarez-Cohen, 1996) during actual field applications. WBC-2 presents an ideal platform for investigating the role of methanogens in dechlorination, because dechlorination and methanogenesis co-occur in WBC-2.

Bioremediation with WBC-2

Bioremediation is a promising treatment for sites contaminated with chlorinated ethanes or mixtures of chlorinated ethanes and ethenes. Until now, however, large-scale cultures capable of dechlorinating compounds such as TeCA and TCA have not been readily available. The microbial consortium WBC-2 was enriched from contaminated sediment at APG and is capable of rapid and complete dechlorination of TeCA under anaerobic conditions with almost no detectable intermediates. WBC-2 has been tested in small-scale bioaugmentation tests (Lorah et al., 2004), in which an addition of 5% (v/v) culture to sediment greatly decreased both the concentration of hazardous intermediates and the time required for complete dechlorination. In a field test at Aberdeen Proving Ground, application of a horizontal permeable barrier seeded with WBC-2 resulted in complete remediation of groundwater contaminated with chlorinated ethanes, ethenes and methanes prior to discharge to the land surface or creek (Lorah et al., 2005; Majcher et al., 2005). WBC-2 represents a useful addition to the bioremediation toolbox with potential uses for bioremediation of sites contaminated with chlorinated ethanes and mixed chlorinated compounds.

ACKNOWLEDGEMENTS

The development of WBC-2 was supported by the National Research Program of the U.S. Geological Survey and the U.S. Army Environmental Conservation and Restoration Division at Aberdeen Proving Ground (John Wrobel). The authors acknowledge SiREM Laboratory (Guelph, Canada) for providing DNA from the scaled-up WBC- 2 culture for use in cloning. They thank Frank Loffler for kindly providing Debalococcoides plasmid. Any use of trade, product, or firm names in this article is for descriptive purposes only and does not imply endorsement by the U.S. Government.

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Elizabeth J. P. Jones and Mary A. Voytek

U.S. Geological Survey, Reston, Virginia, USA

Michelle M. Lorah

U.S. Geological Survey, Baltimore, Maryland, USA

Julie D. Kirshtein

U.S. Geological Survey, Reston, Virginia, USA

Address correspondence to Elizabeth J. P. Jones, U.S. Geological Survey, 430 National Center, Reston, VA 20192, USA. E-mail: ejjones@usgs.gov

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