This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rowe, A. R.
Right arrow Articles by Richardson, R. E.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rowe, A. R.
Right arrow Articles by Richardson, R. E.
Agricola
Right arrow Articles by Rowe, A. R.
Right arrow Articles by Richardson, R. E.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, November 2008, p. 6709-6719, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.00445-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Characterization of the Community Structure of a Dechlorinating Mixed Culture and Comparisons of Gene Expression in Planktonic and Biofloc-Associated "Dehalococcoides" and Methanospirillum Species{triangledown} ,{dagger}

Annette R. Rowe,1 Brendan J. Lazar,2,{ddagger} Robert M. Morris,2,§ and Ruth E. Richardson2*

Field of Microbiology,1 School of Civil and Environmental Engineering, Hollister Hall, Cornell University, Ithaca, New York 148532

Received 23 February 2008/ Accepted 27 August 2008


arrow
ABSTRACT
 
This study sought to characterize bacterial and archaeal populations in a perchloroethene- and butyrate-fed enrichment culture containing hydrogen-consuming "Dehalococcoides ethenogenes" strain 195 and a Methanospirillum hungatei strain. Phylogenetic characterization of this microbial community was done via 16S rRNA gene clone library and gradient gel electrophoresis analyses. Fluorescence in situ hybridization was used to quantify populations of "Dehalococcoides" and Archaea and to examine the colocalization of these two groups within culture bioflocs. A technique for enrichment of planktonic and biofloc-associated biomass was developed and used to assess differences in population distribution and gene expression patterns following provision of substrate. On a per-milliliter-of-culture basis, most D. ethenogenes genes (the hydrogenase gene hupL; the highly expressed gene for an oxidoreductase of unknown function, fdhA; the RNA polymerase subunit gene rpoB; and the 16S rRNA gene) showed no statistical difference in expression between planktonic and biofloc enrichments at either time point studied (1 to 2 and 6 h postfeeding). Normalization of transcripts to ribosome (16S rRNA) levels supported that planktonic and biofloc-associated D. ethenogenes had similar gene expression profiles, with one notable exception; planktonic D. ethenogenes showed higher expression of tceA relative to biofloc-associated cells at 6 h postfeeding. These trends were compared to those for the hydrogen-consuming methanogen in the culture, M. hungatei. The vast majority of M. hungatei cells, ribosomes (16S rRNA), and transcripts of the hydrogenase gene mvrD and the housekeeping gene rpoE were observed in the biofloc enrichments. This suggests that, unlike the comparable activity of D. ethenogenes from both enrichments, planktonic M. hungatei is responsible for only a small fraction of the hydrogenotrophic methanogenesis in this culture.


arrow
INTRODUCTION
 
Anaerobic dechlorination of chlorinated organic compounds is an important mechanism for the remediation of common groundwater pollutants (11, 58). It is now accepted that members of the "Dehalococcoides" group play a crucial role in the remediation of compounds such as chloroethenes, chlorobenzenes, chloroalkanes, chlorophenols, dioxins, and polychlorinated biphenyls, in some cases dechlorinating these compounds to nontoxic end products (1, 2, 6, 16, 18, 26, 32, 35). While researchers have been able to isolate and perform pure culture studies of these organisms, there is significant evidence that reductive dechlorination in environmental systems and in the most robust laboratory cultures is the work of microbial consortia (4, 11). All cultured representatives of the Dehalococcoides group require hydrogen as an electron donor (often supplied by syntrophic fermentation) and a halogenated organic as an electron acceptor. In addition, Dehalococcoides grow robustly in mixed cultures, likely due to currently undetermined growth factors from other community members (11, 18, 31, 34, 44). Though reductive dechlorination is an energetically favorable process under syntrophic conditions with low hydrogen partial pressures (19, 57, 58), other, less favorable metabolic reactions such as methanogenesis and acetogenesis often occur in these communities, especially when excess hydrogen or a donor fermented at high hydrogen partial pressures is available (19, 20, 24, 38). Many methanogens depend on acetate and/or H2, which are both utilized by Dehalococcoides. This suggests that competition for resources is an important interaction within dechlorinating microbial communities containing both methanogenic and Dehalococcoides populations.

Several studies have looked at dechlorinating microbial communities derived from both enrichment cultures and environmental systems (8, 12, 17, 21, 22, 28, 30, 33, 43, 54, 55, 64). Several distinct lineages of microorganisms, representing a variety of metabolic capabilities, are commonly found in these consortia, supporting the potential complexity of community dynamics. The D2 enrichment culture, which has been studied previously (19, 20, 46, 48, 52, 53), is derived from the same consortium from which "Dehalococcoides ethenogenes" strain 195 was isolated (13, 14, 44, 45). In this study, the phylogenetic community structure of the D2 enrichment culture including D. ethenogenes was assessed from phylogenetic analysis of bacterial and archaeal 16S rRNA gene libraries created from community DNA.

Within this heterogeneous enrichment culture, two distinct cellular attachment phases were observed: planktonic cells (individual suspended cells) and cells associated with bioflocs (suspended cell aggregates). In the D2 culture, bioflocs (typically 10 to 100 µm in diameter) tended to contain multiple species and form around mineral precipitates from the medium (see Movie S1 in the supplemental material for a three-dimensional [3D] micrograph). Planktonic and biofloc-associated growth forms are common in environmental microbial communities (i.e., activated sludge, marine, sediments, and groundwater) (5, 10, 41, 60). In this study, a technique for physical enrichment of these two cell attachment phases via low-speed centrifugation was developed. Fluorescence in situ hybridization (FISH) with 16S rRNA-targeting probes was used to estimate the distribution of D. ethenogenes populations between plankton and bioflocs and to examine colocalization of D. ethenogenes and methanogenic Archaea within the bioflocs.

Potential differences in gene expression between the two attachment forms were determined for both D. ethenogenes and the hydrogenotrophic methanogen present in the culture, Methanospirillum hungatei, using quantitative reverse transcription-PCR. This method was also used to compare expression of housekeeping and hydrogenase genes between these organisms. Understanding the distribution and difference in gene expression of the two D. ethenogenes cell attachment phases not only is important for elucidating the ecology of these organisms; it also has implications for the use of DNA and RNA as bioindicators of Dehalococcoides activity. A groundwater sample, while easier and less expensive to obtain, would predominantly sample planktonic Dehalococcoides. Therefore, it is important to establish whether the populations and activities of the planktonic phase reflect those of the community as a whole.


arrow
MATERIALS AND METHODS
 
Chemicals and analysis of chloroethenes.
Butyric acid (99%; Acros Organics) and perchloroethene (PCE) (99%; Alfa Aesar) were used as culture substrates. PCE, trichloroethene (TCE), cis-1,2-dichloroethene, vinyl chloride (VC), and ethene standards were constructed as previously described (53). Methane, ethene, and chlorinated ethenes were measured using the gas chromatography-flame ionization detector temperature program and standard construction as described by Rahm et al. (53). Methane was measured by use of a gas chromatograph equipped with a thermal conductivity detector as described by Fennell et al. (20).

Enrichment culture.
An enrichment culture (D2) containing Dehalococcoides ethenogenes strain 195 has been maintained for over 10 years on a low-PCE/butyrate feeding regimen described previously(20, 53). Under this regimen, the mean cell residence time in the reactors averages 80 days. Briefly, the culture is grown in a 9.1-liter stirred reactor containing 5.7 liters of culture at 30°C. PCE (110 µM) and butyric acid (440 µM) are added at a 2:1 ratio of H2-electron equivalents (assuming that each mole of butyrate is fermented to 2 moles of H2 and 2 moles of acetate).

Culture sampling and cell attachment phase enrichment.
Liquid culture samples of 30 to 50 ml were collected via a stainless-steel valve at the reactor mouth (20, 24). Duplicate 2-ml bulk culture samples (control) for DNA and RNA were pelleted at 21,000 x g for 5 min at 4°C and stored at –20°C or –80°C for DNA and RNA, respectively. For all FISH and enrichment samples, large-orifice pipette tips and gentle pipetting techniques were used to minimize biofloc disruption. Enrichments for biofloc or planktonic cells were done using low-speed centrifugation (100 x g, 500 x g, and 1,000 x g). Alternate separation methods were tested, including gravity settling (settled for 24 h) and selective filtering through 5-µm and 12-µm polycarbonate filters.

To enrich for biofloc-associated and planktonic cells, at various time points culture samples (2-ml aliquots) were distributed into microcentrifuge tubes (eight for RNA, eight for DNA, and two for FISH). Enriched plankton and biofloc regions were created by centrifugation at 1,000 x g for 10 min at 4°C. For each sample type (RNA, DNA, and FISH), the uppermost 0.5 ml (plankton enriched) was pooled to create a 4-ml (for DNA and RNA) or 1-ml (for FISH) plankton-enriched sample. An additional 0.5 ml was discarded, leaving a 1-ml biofloc-enriched region in each tube. Two of these samples were pooled. The biofloc enrichment concentrated bioflocs from 2 ml of culture into 1 ml. In order to convert back to values on a per-milliliter culture basis, data from biofloc-enriched samples were multiplied by a correction factor (0.5). All pooled enrichment samples for DNA and RNA extraction were immediately pelleted at 21,000 x g for 5 min at 4°C and stored at –20°C for DNA or at –80°C for RNA. FISH samples for each sample (control, plankton enriched, and biofloc enriched) were immediately fixed in an equal amount of filter-sterilized phosphate-buffered saline (PBS)-buffered (pH 7) 4% paraformaldehyde solution (EM Sciences) for 6 to 12 h at 4°C (27, 47). After fixation, a subsample was removed for assessment of attachment phase separation (see below). In order to disrupt biofloc structures for ease of counting, samples were briefly sonicated with a sonic dismembrator (Fisher Scientific model no. 100 at a setting of 5 for 5 half-second pulses). Twenty-microliter fixed samples were dispersed in 25 ml of sterile PBS and were then vacuum filtered onto black polycarbonate membrane filters of known filtration area (diameter, 25 mm; pore size, 0.22 µm) (type Poretics; Osmonics, Inc.) supported by binder-free glass fiber support filters (25 mm, 1 µm; type A/B extra thick; Pall).

Assessment of cell attachment phase separation.
For each sample type (i.e., control, plankton enriched, or biofloc enriched), a 20-µl sample was spotted onto coated slides as described below in order to obtain a qualitative assessment of biofloc prevalence. The quality of enrichment using the low-speed centrifugation method was based on biofloc prevalence in at least 100 randomly selected fields of DAPI (4',6'-diamidino-2-phenylindole)-stained cells from each enrichment. Quality and reproducibility of separation were based on nine replicate separation experiments.

Nucleic acid extraction.
For the first bacterial clone library construction, DNA was extracted using the bead-beating, phenol-chloroform protocol of Dojka et al. (15) without the addition of poly(A). Raw DNA was passed through a Chromaspin 1000 column (Clontech) to remove DNA fragments smaller than 1,000 bp. All subsequent DNA extractions were performed using a microbial DNA isolation kit (Mo Bio Laboratories) according to the manufacturers' instructions.

RNA extractions were performed within 48 h of sampling using the bacterial protocol of the RNeasy minikit (Qiagen) with modifications and DNase treatments as previously described (53). Luciferase RNA was added to samples to be used as a measure of overall recovery efficiency as described previously (36). RNA was quantified using the RNA 6000 nanoassay on the Agilent 2100 bioanalyzer (Agilent Technologies).

16S rRNA gene amplification, clone library construction, and sequencing.
One archaeal and three bacterial rRNA gene clone libraries were developed from DNA extracted from the D2 enrichment culture. Bacterial 16S rRNA gene primers 8F (5'-AGA GTT TGA TCC TGG CTC AG) and 1492R (5'-GC[C/T] TAC CTT GTT ACG ACT T) were used as previously reported (22, 54) with annealing temperatures of 53°C and 55°C. Archaeal 16S rRNA gene primers 1Af (5'-TCY GKT TGA TCC YGS CRG AG) and 1100Ar (5'TGG GTC TCG CTC GTT G) were used as previously described (29). 16S rRNA gene clone library construction and restriction fragment length polymorphism type screening were performed as described previously (22, 54). All cloning was performed using a TOPO TA cloning kit with DH5{alpha}-T1 chemically competent cells (Invitrogen). Clones for each bacterial restriction fragment length polymorphism type were sequenced using the M13 forward and reverse primers. In some cases, a third internal primer (the bacterial 515F primer, 5'-GTG CCA GC[A/C] GCC GCG GTA A) was employed in order to obtain full sequences (22). Sequencing reactions were carried out using BigDye terminator chemistry according to the manufacturer's instructions (Applied Biosystems) and analyzed at the Cornell Biotechnology Resource Center using an Applied Biosystems automated 3730 DNA analyzer. Sequence assembly was performed using SeqBuilder software (Lasergene). ChimeraCheck through the Ribosomal Database Project (http://rdp8.cme.msu.edu/html/analyses.html) or through the Bellerophon server (http://foo.maths.uq.edu.au/~huber/bellorophon.pl) was performed on the assembled sequences. BLAST searches (http://www.ncbi.nlm.nih.gov/BLAST/) were run to obtain putative phylogenetic affiliations and assign more informative names to the sequences. Retrieved sequences were then aligned with a 16S rRNA gene database maintained by the Ribosomal Database Project (http://rdp.cme.msu.edu) using ARB (http://www.arb-home.de) (42).

For assessment of lineage or classification of publicly available sequences from chloroethene-reducing enrichment cultures, microcosms, and field studies, as well as the D2 enrichment culture, the Classifier function available through the Ribosomal Database Project (http://rdp.cme.msu.edu/classifier/classifier.jsp) was used. This function, a naïve Bayesian rRNA classifier, was developed to assess taxonomy from domain to genus with confidence estimates for each assignment (62). A 90% confidence level was used as a cutoff for determination of phylogenetic assignment.

DGGE.
To ensure completeness of the community composition observed in clone library analysis, denaturing gradient gel electrophoresis (DGGE) was performed in triplicate on DNA samples used for clone library construction. PCRs (40-µl reaction mixtures) were carried out as described by Nakatsu et al. (49) using bacterial primers PRBA338F (amended with a 40-bp GC region at the 5' end) and PRUN518R, designed to target the V3 region. Archaeal DGGE primers PARCH340F (with a 5' GC tag) and PARCH519R, designed to target the archaeal V3 region, were also used (49). Reaction programs consisted of 9 min at 94°C; followed by 30 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 30 s; followed by a 7-min extension period at 72°C. DGGE was carried out using the D-code system (Bio-Rad). PCR products were resolved on an 8% (wt/vol) polyacrylamide gel using 1x Tris-acetate-EDTA buffer with a denaturing gradient from 35% to 55% denaturant (40% [wt/vol] formamide and 7 M urea). Gels were visualized using Sybr green (Molecular Probes). Bands excised for sequencing were run on a second DGGE gel under the same conditions to ensure purity. Sequences were obtained using DGGE primers without GC flanking regions. Sequencing reactions were carried out as described for the 16S rRNA gene clone library, which provided 150 to 300 bp of sequence information.

FISH, Dehalococcoides probe analysis, and fluorescence microscopy.
Sixty-six Dehalococcoides and 8,794 total 16S rRNA sequences were aligned using the ARB sequence analysis package (42) to design two new Dehalococcoides specific probes (see Table S2 in the supplemental material). Dehalococcoides-targeting probes (two from this study along with Dhe1259degR [63] and Dhe201R [50]) were evaluated with a mechanistic FISH model based on the thermodynamics of nucleic acid hybridization as described by Yilmaz et al. (65). All probes were labeled with Cy3. Probe intensity was tested for both individual and probe combinations. Cell images were captured with an Olympus BX61 fluorescence microscope equipped with a Cooke SensiCam high-performance charge-coupled device digital camera, filter sets appropriate for DAPI and Cy3, and Intelligent Imaging Innovations Slidebook software, version 3.0.10.15. Exposure times were 1 second for probed samples (Cy3) and 100 milliseconds for DAPI.

Hybridization reactions were performed as described previously (27, 47) with the following modifications. Air-dried filters were stored with desiccant at –20°C prior to hybridization. Quarter membrane sections were pretreated by dipping filters in 0.2 M HCl for 10 minutes, followed by a 2-min wash with PBS to reduce background fluorescence(51). Hybridizations were then performed at 37°C for 16 h in hybridization buffer with the desired probe(s). Control hybridization reactions, stringency washes, and DAPI counterstaining were employed as described previously (47), except for a 48°C dissociation temperature. Hybridization and wash buffers were made as described by Yang and Zeyer (63); however, 20% formamide (EM Science) was used in all hybridizations. All probes were filter sterilized through 0.2-µm-pore filters to a final concentration of 2 ng/µl. Cell counts were performed on images captured via fluorescence microscopy with a minimum of 10 fields (field area of 5,292 µm2) per duplicate samples. Cell count microscopy was carried out on an Olympus BX-50 at the Cornell Microscope Imaging Facility at a magnification of x630.

Multiplex FISH for visualizing bioflocs.
The FISH protocol described above was amended to facilitate observation of biofloc architecture and associations. Ethanol-cleaned slides were coated by being dipped in a warmed (70°C) gelatin-chromium solution of 0.1% gelatin and 0.01% chromium(III) potassium sulfate (Aldrich) (3). Twenty microliters of fixed samples was spotted with large-orifice pipette tips onto gelatin-chromium-coated slides over a circular area of approximately 1 cm in diameter. Cell spots were allowed to air dry and then dipped sequentially in 50%, 80%, and 95% ethanol for 3 minutes each. Slides were air dried again and either hybridized as described above, DAPI stained, or stored at –20°C until hybridization. Multiplexing with differentially labeled probes (Cy3 for Dhe1259/Dhe201 and fluorescein for ARCH915) was employed in the hybridization of preserved bioflocs. A Leica confocal TCS SP2 microscope system was used to collect z-series stacks (z-step size of 0.12 µm) from 60 images of both Cy3 (excitation range, 510 to 560 nm; emission range, >590 nm) and fluorescein (excitation range, 460 to 500 nm; emission range, 512 to 542 nm) channels. Leica and Leica Lite software was employed in creation of 3D projections and visualization of individual bioflocs.

Reverse transcription and qPCR.
RNA and DNA were extracted from two replicate separation experiments. From the replicate RNA pools, cDNA was synthesized from 0.2 µg of RNA using the iScript cDNA synthesis kit (Bio-Rad) with random hexamers as primers according to the instructions of the supplier. Primers targeting D. ethenogenes strain 195 genes rpoB (DET0603 on the D. ethenogenes genome), hupL (DET0110), fdhA (DET0187), and tceA (DET0079) were developed previously (25, 53). D. ethenogenes primers targeting the 16S rRNA genes (2) were also used. Quantitative PCR (qPCR) primers were designed for the 16S rRNA genes of the two D2 methanogens obtained from clone library sequencing, Methanospirillum hungatei (MHU16S F/R, 5'-AGT AAC ACG TGG ACA ATC TGC CCT and 5'-ACT CAT CCT GAA GCG ACG GAT CTT) and a Methanosaeta sp. (Ms16S F/R, 5'-GGG GTA GGG GTG AAA TCT TGT AAT CCT and 5'-CGG CGT TGA ATC CAA TTA AAC CGC A), using PrimerQuest available through IDT (Coralville, IA).

Expression of D. ethenogenes and M. hungatei hydrogenase and RNA polymerase subunit genes was used for comparison between organisms. Because the genome sequence of the culture-specific Methanospirillum population was not known, in order to determine hydrogenase sequences for the Methanospirillum sp. present in our culture, degenerate hydrogenase primers for methyl viologen-reducing hyrdrogenases (mvrD) in methanogens were designed using orthologous mvrD sequences. PCR products were cloned and sequenced (as described for the 16S rRNA gene clone library) and used to design primers appropriate for qPCR for the mvrD gene present in the D2 culture (D2mvrh F/R, 5'-TGT TCG TAT GCA GGT GCT GAC CTT-3' and 5'-ACC ATC TGC ACC CTC AAC AAA TGC-3' (accession no. EU498366). Using the sequence available for the Methanospirillum hungatei JF-1 rpoE gene (YP_504275), a qPCR primer set was designed using PrimerQuest as described above (Msp rpoE-F/R, 5'-TCA GTC TTG GAC CGA TTG ATG CGA-3' and 5'-TCA CGA GGT TCA CGT TCG TTG AGA-3').

All qPCRs were performed using the iCycler iQ Multicolor real-time PCR detection system (Bio-Rad). Triplicate qPCRs for each sample were constructed along with standard curves (log DNA concentration versus cycle threshold) as described by Rahm et al. (53). Methanospirillum hungatei JF-1 and Methanosaeta thermophila CALS-1 pure-culture DNA samples were quantified using PicoGreen assays (Invitrogen) and converted to genome copies using molecular weights of the published genomes. Dilutions were used as standards for the 16S rRNA gene copies of D2 methanogens as well as Methanospirillum sp. rpoE and mvrD copies using the iCycler method. Luciferase DNA stock (Promega) was used to generate standard curves for quantification of recovered transcripts as described by Johnson et al. (36). qPCR conditions and melt curve analysis were described previously (53). Methanogen primer sets used the same qPCR program as described for D. ethenogenes with a 60°C annealing temperature.

Statistical analysis.
Statistical tests were performed using JMP statistical software. The statistical significance of gene expression data was determined using an unpaired t test in addition to analysis of variance to determine experimental effects on these values. F tests were done to ensure that there was no statistically significant difference in variance between the gene expression values being compared.

Nucleotide sequence accession numbers.
The sequences determined in this study have been submitted to GenBank under accession numbers EU498367 to EU498393.


arrow
RESULTS
 
Phylogenetic analysis of the D2 enrichment culture.
The phylogenetic structure of the D2 enrichment culture was determined through the classification of archaeal and bacterial 16S rRNA gene clone libraries and DGGE profiling (Fig. 1). Cumulative analysis of clone libraries suggests that there are around 18 bacterial and archaeal operational taxonomic units present in the culture. In order to ensure that we were observing all the major members of the D2 enrichment culture, we employed a second community characterization method, DGGE, on replicate DNA extractions. Replicate extracts produced a consistent profile (Fig. 1). Sequences obtained from dominant bands (bands a through l in Fig. 1) matched sequences from the D2 bacterial clone libraries.


Figure 1
View larger version (38K):
[in this window]
[in a new window]

  		FIG. 1. Phylogenetic tree (left) containing sequences from a D2 enrichment clone library (sequences designated D2CL_) and from cultured representatives. This unrooted tree was constructed from a 770-bp alignment using ARB with a Kimura correction parameter. Brackets indicate higher-order taxonomic groupings. Lowercase letters correspond to matching sequences obtained from DGGE bands (right). Certain bands matched more than one sequence (i.e., b matched multiple deltaproteobacteria).

In the archaeal clone library, Methanosarcinales and Methanomicrobiales were the only two phylogenetic groups represented. The nearest cultured organisms were an acetotrophic methanogen (a Methanosaeta sp.) (97% identity over 1,060 bp) and a hydrogenotrophic methanogen (Methanospirillum hungatei JF-1) (98% identity over 1015 bp). DGGE using Archaea-specific primers resulted in two bands. One band matched a Methanosaeta sp., while the other produced a short sequence similar to sequences in many organisms in the Methanomicrobia, including Methanospirillum (data not shown).

Dehalococcoides probe comparison.
In order to enumerate populations of Dehalococcoides ethenogenes strain 195, we aimed to improve FISH for this organism. The previously published probe Dhe1259degR (64) produced the highest value for average fluorescence intensity, followed by Dhe201R, Dhe137R, and Dhe619R (see Table S2 in the in the supplemental material). With actively dechlorinating cells, attempts to increase signal intensity through probe combinations did not improve signal intensity above that of Dhe1259degR used alone; however, incorporation of a second probe, Dhe201, did improve detection of starved cells (percent D. ethenogenes to DAPI, 21% ± 15% with Dhe1259degR alone versus 65% ± 8% with Dhe1259 and Dhe201R combined). This combination was used subsequently for cell enumeration.

Enumeration of specific populations via FISH.
FISH with DAPI counterstaining was employed as a method to simultaneously visualize and quantify both Dehalococcoides (D. ethenogenes in this culture) and archaeal (methanogens in this culture) populations. Counts for three time points in a batch feeding cycle (prefeeding, 6 h [active dechlorination; PCE to VC], and 16 h [cometabolic dechlorination; VC to ethene]) (53) showed that the total culture population ranged from 6.3 x 108 to 9.9 x 108 cells per ml, with an average of 7.9 x 108 cells per ml across all time points. D. ethenogenes and archaeal cells averaged 60.1% ± 18.1% and 10.9% ± 5.7% of the total cells, respectively. Based on these techniques, any growth observed during one dechlorination cycle (110 µM PCE) is within the error of the measurements.

In addition to FISH with dispersed culture samples, multiplex FISH was employed on spotted culture samples to examine localization of the D. ethenogenes and Archaea within the bioflocs that are common in this culture (Fig. 2). Archaeal and D. ethenogenes cells were consistently observed in close association around black medium precipitates. A full 3D projection of this biofloc is available in Movie S1 in the supplemental material.


Figure 2
View larger version (11K):
[in this window]
[in a new window]

  		FIG. 2. Microscopic field of view with a typical biofloc multiplexed with both Arch915R (green) and Dhe201R/1259degR (red). The 3D rotation of this biofloc is available as Movie S1 in the supplemental material.

Separation and enumeration of cells in plankton-enriched and biofloc-enriched samples.
When observing the bioflocs using FISH on bulk culture, it was noted that some cells were located outside of these bioflocs (planktonic cells). We describe these as two distinct cellular attachment phases: planktonic and biofloc associated. A low-speed centrifugation separation technique was developed to separate D2 culture samples into plankton-enriched and biofloc-enriched samples. Quality of enrichment was assessed through microscopy on DAPI-stained spotted cells. Of the centrifuge speeds tested (100 x g, 500 x g, and 1,000 x g), 1,000 x g for 10 min produced the most complete separations. Other techniques were not as successful at enriching these two phases: the gravity settling separation technique yielded poorer separation of bioflocs, and size exclusion filtering attempts were unsuccessful because bioflocs became sheared under the vacuum pressure (data not shown). Biofloc prevalence counts over the course of nine centrifugation separation experiments suggested that this technique produces a significant and consistent reduction in the number of bioflocs in plankton-enriched samples. The average percentages of fields with bioflocs for the control, plankton-enriched, and biofloc-enriched samples were 76% ± 13%, 17% ± 8%, and 88% ± 8%, respectively. Qualitatively, the bioflocs detected in the plankton-enriched samples were smaller than those in the bulk (control) or biofloc-enriched samples (data not shown). However, since separation was not absolute, the terms "biofloc enriched" and "plankton enriched" are used throughout this report.

Cell counts via DAPI staining and D. ethenogenes counts via use of Dhe201R/Dhe1259degR were determined for samples of bulk culture (control), plankton-enriched, and biofloc-enriched samples at 2 hours and 6 hours postfeeding (Fig. 3). In terms of percentage of total cells, the plankton-enriched samples were enriched for D. ethenogenes (78% ± 6%), while the biofloc-enriched samples (53.4% ± 1%) contained a lower percentage of D. ethenogenes relative to control samples (64.2% ± 2%). qPCR was also used as a technique for enumerating D. ethenogenes populations. Both FISH and qPCR suggest that D. ethenogenes cells in the culture are equally distributed between cell locations (Fig. 3 and 4). Direct comparison of FISH and qPCR numbers on duplicate samples showed an average ratio of qPCR numbers to FISH numbers of 1.8 ± 1.5. Removal of a qPCR outlier two standard deviations above the mean resulted in a ratio of 1.3 ± 0.6. In subsequent studies examining gene expression in the different cell attachment phases, qPCR was used for quantification of populations due to the simplicity of the assay relative to FISH. Also, the processing and quantification biases are similar to those of quantitative reverse transcription-PCR (the method employed for assaying gene expression levels).


Figure 3
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 3. Summary of DAPI and Dehalococcoides (DET)-specific cell counts for control (untreated) culture, planktonic enrichments, and biofloc enrichments at 2 and 6 h postfeeding. Error bars represent standard deviations.


Figure 4
View larger version (25K):
[in this window]
[in a new window]

  		FIG. 4. 16S rRNA gene copies present in each enrichment (control, planktonic, or biofloc) of each replicate experiment. (A) Dehalococcoides ethenogenes strain 195 16S rRNA gene copies; (B) Methanospirillum hungatei rpoE gene copy numbers; (C) Methanosaeta sp. 16S rRNA gene copies. Panels A and B represent known single-copy genes, whereas panel C potentially has multiple copies per genome. Error bars represent standard errors of values from two separate DNA extractions.

During gene expression experiments, the D. ethenogenes population numbers determined by qPCR of 16S rRNA gene copies for the control culture over two time points averaged 6.7 x 108 and 1.3 x 108 for replicates A and B, respectively (Fig. 4A). In general, the qPCR estimates for all enrichments in replicate A were higher than those for in replicate B (Fig. 4). As in earlier studies with FISH and qPCR, an equal distribution of D. ethenogenes cells between biofloc and planktonic enrichments (averaging 47 and 53%, respectively) was noted on both of these dates. Unlike D. ethenogenes, methanogenic cells (both Methanosaeta sp. and Methanospirillum) were present predominantly in biofloc enrichments (Fig. 4B and C). This matched previously observed FISH assays on bulk culture, where few archaeal cells were observed outside bioflocs (data not shown).

Gene expression in planktonic and biofloc-associated Dehalococcoides.
Gene expression levels in planktonic and biofloc-associated D. ethenogenes were compared at two time points after provision of PCE and butyrate, specifically, 1 to 2 h and 6 h (Fig. 5). The 6-hour time point was selected based on recent studies in our laboratory showing that this time point is during active dehalorespiration of the chlorinated ethenes and is also a point at which a variety of respiratory genes were upregulated in whole-culture samples (53). The general expression trends in the control culture as well as enrichments showed that the hydrogenase gene hupL, an annotated formate dehydrogenase gene (fdhA), and an RNA polymerase gene (rpoB) were more highly expressed at early (1 to 2 h) time points postfeeding, while expression levels of the reductive dehalogenase gene tceA was higher at 6 h postfeeding. Comparing planktonic and biofloc enrichments, there was no statistically significant difference in the relative expression of rpoB, hupL, or fdhA between the enrichments at either time point (Fig. 5A, C, and D). However, expression of the reductive dehalogenase gene tceA at 6 hours was higher in planktonic enrichments (1.5 x 109 ± 5.6 x 108) than in biofloc enrichments (7.5 x 108 ± 1.1 x 108) (Fig. 5B) (P = 0.0004). Testing for the effect of experiment by analysis of variance suggested an effect of experimental replicate on per-milliliter culture data (P < 0.0001), though this did not detract from the significant difference in the tceA expression mean mentioned above. The effect of experiment is likely a result of minor differences in the starting culture between different separation experiments.


Figure 5
View larger version (22K):
[in this window]
[in a new window]

  		FIG. 5. Average gene expression values for Dehalococcoides ethenogenes strain 195 on a per-milliliter enrichment basis (control, planktonic, or biofloc) at 1 to 2 h and 6 h postfeeding. (A) rpoB; (B) tceA; (C) hupL; (D) fdhA; (E) 16S rRNA copies (ribosome copies). Error bars represent standard errors of values from replicate experiments.

In order to look more closely at any differences in relative expression of different genes in the two attachment phases, transcript data were normalized to 16S rRNA copies from the same RNA pool (Fig. 6). Using rRNA as an internal normalizing factor minimized any variability due to slight differences in cell density or cell lysis across replicates; the effect test suggested no effect of experiment for rRNA normalized values (P = 1.00). Normalized values support the trends noted earlier, i.e., twofold higher relative expression of tceA at 6 h in planktonic cells (0.36 tceA transcript per ribosome, versus 0.16 tceA transcript per ribosome for biofloc-associated cells [P < 0.0001]). Only slight differences in ribosome-normalized transcript abundance were observed between planktonic and biofloc-enriched D. ethenogenes for rpoB or fdhA at either time point. On a per-cell basis, the numbers of 16S rRNA (ribosome) copies per DNA gene copies (48 ± 32 per cell) were not statistically higher or lower in either enrichment.


Figure 6
View larger version (29K):
[in this window]
[in a new window]

  		FIG. 6. 16S rRNA normalized gene expression for Dehalococcoides ethenogenes strain 195 at 1 to 2 h postfeeding (A) and 6 h postfeeding (B) for each enrichment (control, planktonic, or biofloc). Error bars represent standard errors of replicate experiments.

Planktonic and biofloc-associated Methanospirillum.
As a hydrogenotrophic methanogen, Methanospirillum hungatei competes with Dehalococcoides for electron equivalents (19). Transcripts of M. hungatei functional genes in RNA pools from the same enrichments were also studied. Primers for the well-conserved delta subunit of the methyl viologen-reducing hydrogenase (mvrD) were designed. This coenzyme F420 hydrogenase has been shown to interact with (provide electrons to) heterodisulfide reductase, an enzyme involved in the last step in methanogenesis (9). The sequences recovered from clone sequencing (EU498366) were most closely related to Methanospirillum hungatei JF-1 (YP_503279) (100% identity over 186 bp of mvrD).

Given that the majority of M. hungatei cells were present in biofloc enrichments (Fig. 4B), detection of larger numbers of transcripts for the housekeeping gene rpoE and the hydrogenase gene mvrhD in the biofloc enrichment was expected (Fig. 7). For mvrD but not rpoE, a very strong dependence on time was observed, with mvrD showing a strong increase (~10-fold) between the early and late time points. A lag time in the onset of mvrD expression has been previously observed in our lab and corresponds to a lag in the onset of methane accumulation in batch cultures (data not shown). Ribosome-normalized expression data did not show any difference between enrichments (data not shown). On a per-cell basis, plankton-enriched M. hungatei contained 8.5 x 103 ± 2.5 x 103 ribosomes and biofloc-enriched cells contained significantly more, with 1.4 x 104 ± 3.6 x 103 ribosomes per DNA copy. M. hungatei in both attachment phases contained ribosome densities over 2 orders of magnitude higher than those in D. ethenogenes cells on a per-cell basis.


Figure 7
View larger version (18K):
[in this window]
[in a new window]

  		FIG. 7. Average gene expression values for Methanospirillum hungatei on a per-milliliter enrichment basis (control, planktonic, or biofloc) at 1 to 2 h and 6 h postfeeding. (A) rpoE; (B) mvrD; (C) 16S rRNA copies (ribosome copies). Error bars represent standard errors of values from replicate experiments.


arrow
DISCUSSION
 
The D2 enrichment culture containing Dehalococcoides ethenogenes strain 195 has been the subject of many previously published studies. This report presents the first phylogenetic characterization of the bacterial and archaeal groups in D2. From the bacterial libraries, representatives were found from the Firmicutes, Delta-/Epsilonproteobacteria, Bacteriodetes, Nitrospirales, Thermotogales, Spirochetes, and Chloroflexi. Each of these taxa has been previously observed in chloroethene-reducing communities. However, considering the diversity within many of these phyla and classes, we sought to assess the similarity within representatives of these groups, which showed conservation at the family level and even the genus level for certain operational taxonomic units (see Table S1 in the supplemental material). The Clostridia were represented in all studied clone libraries. Within the class certain families were favored, especially the Clostridiaceae. While phylogeny cannot completely inform function and many of these representative groups are physiologically diverse, the similar phylogenetic structure of these communities suggests that the potential roles of these operational taxonomic units in chloroethene-degrading communities are likely conserved. For the D2 enrichment culture, it has previously been shown that butyrate is converted to H2 and acetate (19, 20). This conversion is a microbially mediated process which involves one if not several organisms in the culture. Candidate syntrophic fermenters are different members of the Firmicutes, the Bacteroidetes, and some of the Delta-/Epsilonproteobacteria (7).

Though we cannot yet determine the specific organisms responsible for all metabolic processes occurring in this culture, two functions that we know occur and have been able to associate with the appropriate phylogenies are methanogenesis (via hydrogenotrophic Methanospirillum and acetoclastic Methanosaeta) and reductive dechlorination (via D. ethenogenes). The detection of a single D. ethenogenes phylotype in our culture is consistent with previous Dehalococcoides-specific DGGE analyses (18). While the clone library results illuminate which archaeal and bacterial taxa are present, they do not provide insights into the relative or absolute populations.

The distribution and localization of D. ethenogenes and archaeal (methanogenic) populations was studied using both FISH and qPCR. Methanogens tended to form long strands of cells around the exteriors of bioflocs, while D. ethenogenes cells were more evenly distributed within bioflocs (Fig. 2) as well as between biofloc and planktonic phases. It is unknown how or why D. ethenogenes associates in these different phases, though each growth form is likely to have benefits, with planktonic cells having more access to soluble nutrients and biofloc cells better access to nutrients provided by other organisms. It has been postulated that spatial orientation is important in syntrophic communities, as it facilitates the transfer of metabolites between organisms (in this case between butyrate fermenters and H2 consumers) (59). As such, different environmental conditions may favor one attachment phase over another; however, further study of culturing conditions and growth of these organisms in different phases would help determine the factors controlling attachment phase. Additional multiplex FISH studies targeting putative fermenters will further illuminate the association of the various syntrophic populations in bioflocs.

A direct comparison of absolute numbers of D. ethenogenes cells suggested that, in general, qPCR values were higher and more variable than FISH estimates, i.e., 1.8 ± 1.5 times higher, or 1.3 ± 0.6 times higher excluding an outlier (2.5 x 109), which is twice the highest total cell density observed by DAPI during these experiments. While the errors for these ratios are large (stemming from the fact they are a combination of two values, each with large associated errors), they suggest that in general qPCR reports higher values than FISH. Both qPCR and FISH are subject to their own biases. qPCR is a measure of DNA copies and can both over- and underestimate viable populations. Overestimation can be the result of individual cells containing multiple copies just before cell division as well as contributions from dead or inactive cells. On the other hand, poor cell lysis or DNA recovery can lead to an underestimation of populations via qPCR. FISH does not specifically discriminate between live and dead cells, but inactive cells are likely difficult to detect using FISH due to a low ribosome content. The discrepancies between these two methods, as they are commonly used to measure biomass, suggest that further investigation of factors that affect variability in these methods is needed. It also suggests that the method of quantification should be taken into account when considering reported values.

In this study, we showed that even active D. ethenogenes cells contain a small number of ribosomes (48 ± 32 ribosomes per cell) compared to M. hungatei (14,000 ± 3600 ribosomes per cell). Given that these values are a ratio of assays for two different nucleotides (RNA/DNA), each of which is subject to different biases and associated errors, the large standard deviations are not surprising. Even considering these large standard deviations, there is a dramatic difference in per-cell ribosome content between M. hungatei and D. ethenogenes. These differences are likely a function of size. D. ethenogenes cells are flattened cocci approximately 0.5 µm in diameter and 0.2 µm thick (44), giving a biovolume of 40 nm3 (based on cylindrical volume {pi}r2h). M. hungatei JF1 cells are long spiral chains of 0.4 to 0.5 µm in diameter and can range from 7.4 to 10 µm in length (7), with an approximate biovolume of 1,400 to 1,900 nm3, or approximately 40 times the volume of D. ethenogenes. Though cell populations given by DNA copies suggest that methanogens are far less numerous organisms, in terms of ribosomes per milliliter of control culture (Fig. 5E and 7C), there are approximately six times more Methanospirillum ribosomes than D. ethenogenes ribosomes. This prevalence of methanogens in terms of biovolume is not surprising considering that this culture is given butyrate in excess of what is required to reduce the supplied PCE to ethene. In a typical feeding cycle, a similar number of moles of electron equivalents go to the production of methane (approximately 250 µmol methane per liter of culture from both Methanosaeta and Methanospirillum) as go to reducing PCE (approximately 288 µmol VC plus ethene per liter culture), specifically, 2.0 and 2.3 milli-electron equivalents per liter of culture based on respective end product productions (data not shown). Population numbers given by qPCR may suggest that methanogens are only minor members of the community. However, in terms of biovolume in the culture, ribosomes per milliliter, and portion of electron equivalents that are converted to end products, they are on par with the dechlorinators.

Overall gene expression levels and ribosome contents in D. ethenogenes cells from each cell attachment phase were similar for most targets investigated. One exception was the planktonic D. ethenogenes's higher expression of tceA, which encodes the TCE reductive dehalogenase in D. ethenogenes and has shown promise as a specific bioindicator of TCE respiration at contaminated field sites (36, 37, 39, 40, 61). Given that no other differences in gene expression were observed between the D. ethenogenes in planktonic and biofloc enrichments (similar overall transcriptional activity), this result suggests differential expression of tceA in different cell locations. Though specific factors controlling reductive dehalogenase expression have yet to be elucidated, the differences in expression trends for different reductive dehalogenases suggest tight transcriptional control (25, 40, 52, 53, 61). In the case of bioflocs, mass transfer of an exogenously supplied chlorinated substrate or other small molecules (e.g., vitamins) may explain the resulting lower expression of tceA in the biofloc-associated cells relative to the planktonic cells. As bioflocs are presumed to be a major site of butyrate fermentation to H2, one would expect biofloc-associated D. ethenogenes to experience higher H2 partial pressures. However, no statistical difference in hydrogenase expression between planktonic and biofloc associated D. ethenogenes was noted. This could be due to a lack of sensitivity in our assay (incomplete separation and/or variation in the method) obscuring true differences. However, previous studies with D. ethenogenes showed very little difference in expression of hupL in D. ethenogenes grown in 0.1 atm versus 10–4 to 10–5 atm H2 partial pressures (46).

The strong temporal trend for the M. hungatei hydrogenase gene mvrD (but not for the housekeeping gene rpoE) may be explained by the nature of hydrogen production in this culture via interspecies hydrogen transfer. In this culture, butyrate is fermented to hydrogen and acetate, with H2 levels stabilizing near 10–5 atmospheres after 2 to 3 h (19, 20, 57). These studies have reported that H2 thresholds for methanogens are higher than those for D. ethenogenes. The delayed expression of mvrD may be tied to the accumulation of H2 above a threshold level for gene induction at somewhere between 2 and 6 h postfeeding. Under different culturing conditions, differences in distribution of populations and/or gene expression between enrichments may become more or less pronounced for either D. ethenogenes or M. hungatei.

For the M. hungatei populations, the biofloc-associated cells were the predominant contributors to overall gene transcript levels. In terms of number of transcripts per ml, M. hungatei had 1 or 2 orders of magnitude fewer transcripts than D. ethenogenes, depending on the gene (Fig. 6). The corresponding ratios of transcripts to ribosomes for M. hungatei (all bellow 10–3) were also at least an order of magnitude lower than those for D. ethenogenes. The greater number of ribosomes and fewer transcripts per milliliter of enrichment for M. hungatei compared to D. ethenogenes suggest that these two organisms may have different translational strategies. As M. hungatei harbors more ribosomes than D. ethenogenes, it may require fewer transcripts to maintain its protein pool. Further work that examines the relationship between transcript abundance and ribosome content in combination with the quantification of protein pools will further our understanding of the expression differences between these organisms.

The activity of planktonic Dehalococcoides populations along with their small size and tolerance for low hydrogen partial pressures may explain why organisms from Dehalococcoides-containing enrichment cultures, such as the commercially available KB-1, are so successful for bioaugmentation. Though growth of Dehalococcoides organisms in mixed communities is more robust, dispersal in an aquifer is likely dominated by planktonic rather than biofloc-associated Dehalococcoides. To assess the activity of Dehalococcoides in an environmental setting, it has been suggested that DNA copies or mRNA transcripts could be useful bioindicators of in situ dehalorespiration. Since these organisms can be found in association with other organisms (biofloc) or as planktonic cells in environmental settings, it is important to establish what differences may exist in gene copies and gene expression between these attachment phases. Our data suggest that, though some differences in gene expression exist, no cell attachment phase appears to be significantly more or less transcriptionally active than the other. This suggests that the activity of planktonic Dehalococcoides cells in groundwater samples, which are easier and less costly to obtain than soil cores, should reflect broader activity of Dehalococcoides in the subsurface. However, additional studies where mRNA transcript levels in groundwater and soil samples are directly compared at active bioremediation sites are needed.


arrow
ACKNOWLEDGMENTS
 
We sincerely thank L. Safak Yilmez, Daniel R. Noguera, and the members of their lab for their analysis of probe hybridization free energies displayed in Table S1 in the supplemental material. For assistance with lab techniques and troubleshooting, we thank Laura Jennings, Emily Warren, and especially Brian Rahm. Special thanks go to Janice E. Thies for use of laboratory equipment supplies and to Angelika Rumberger for her technical expertise and help with DGGE. Thanks go to Steve Zinder for supplying methanogen pure cultures and general assistance and advice throughout the project.

This work was supported through a Biogeochemistry and Environmental Biocomplexity IGERT small grant (DGE 0221658), by Department of Defense Army Research Office grant W911NF-07-1-0249, and by the National Science Foundation's Graduate Research Fellowship.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: 220 Hollister Hall, Cornell University, Ithaca, NY 14853. Phone: (607) 255-3233. Fax: (607) 255-9004. E-mail: rer26{at}cornell.edu Back

{triangledown} Published ahead of print on 5 September 2008. Back

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

{ddagger} Present address: TRC Environmental Corporation, 57 East Willow Street, Millburn, NJ 07041. Back

§ Present address: University of Washington, Center for Environmental Genomics, Seattle, WA 98105. Back


arrow
REFERENCES
 
    1
  1. Adrian, L., and H. Goerisch. 2002. Microbial transformation of chlorinated benzenes under anaerobic conditions. Res. Microbiol. 153:131-137.[Medline]
    2. 2
  2. Adrian, L., S. K. Hansen, J. M. Fung, H. Gorisch, and S. H. Zinder. 2007. Growth of Dehalococcoides strains with chlorophenols as electron acceptors. Environ. Sci. Technol. 41:2318-2323.[Medline]
    3. 3
  3. Amann, R. I., L. Krumholz, and D. A. Stahl. 1990. Fluorescent-oligonucleotide probing of whole cells for determinative phylogenetic and environmental studies in microbiology. J. Bacteriol. 172:762-770.[Abstract/Free Full Text]
    4. 4
  4. Becker, J. G., G. Berardesco, B. E. Rittmann, and D. A. Stahl. 2005. The role of syntrophic associations in sustaining anaerobic mineralization of chlorinated organic compounds. Environ. Health Perspect. 113:310-316.[Medline]
    5. 5
  5. Becker, P., W. Hufnagle, G. Peters, and M. Herrmann. 2001. Detection of differential gene expression in biofilm-forming versus planktonic populations of Staphylococcus aureus using microrepresentational-difference analysis. Appl. Environ. Microbiol. 67:2958-2965.[Abstract/Free Full Text]
    6. 6
  6. Bedard, D. L., K. A. Ritalahti, and F. E. Loffler. 2007. The Dehalococcoides population in sediment-free mixed cultures metabolically dechlorinates the commercial polychlorinated biphenyl mixture aroclor 1260. Appl. Environ. Microbiol. 73:2513-2521.[Abstract/Free Full Text]
    7. 7
  7. Boone, D. R., R. W. Castenholz, and G. M. Garrity (ed.). 2001. Bergey's manual of systematic bacteriology. Springer, New York, NY.
    8. 8
  8. Bowman, K. S., W. M. Moe, B. A. Rash, H. S. Bae, and F. A. Rainey. 2006. Bacterial diversity of an acidic Louisiana groundwater contaminated by dense nonaqueous-phase liquid containing chloroethanes and other solvents. FEMS Microbiol. Ecol. 58:120-133.[CrossRef][Medline]
    9. 9
  9. Brodersen, J., G. Gottschalk, and U. Deppenmeier. 1999. Membrane-bound F420H2-dependent heterodisulfide reduction in Methanococcus voltae. Arch. Microbiol. 171:115-121.[CrossRef][Medline]
    10. 10
  10. Busch, P. L., and W. Stumm. 1968. Chemical interactions in the aggregation of bacteria bioflocculation in waste treatment. Environ. Sci. Technol. 2:49-53.[Medline]
    11. 11
  11. Chen, G. 2004. Reductive dehalogenation of tetrachloroethylene by microorganisms: current knowledge and application strategies. Appl. Microbiol. Biotechnol. 63:373-377.[CrossRef][Medline]
    12. 12
  12. Dennis, P. C., B. E. Sleep, R. R. Fulthorpe, and S. N. Liss. 2003. Phylogenetic analysis of bacterial populations in an anaerobic microbial consortium capable of degrading saturation concentrations of tetrachloroethylene. Can. J. Microbiol. 49:15-27.[CrossRef][Medline]
    13. 13
  13. DiStefano, T. D., J. M. Gossett, and S. H. Zinder. 1992. Hydrogen as an electron donor for dechlorination of tetrachloroethene by an anaerobic mixed culture. Appl. Environ. Microbiol. 58:3622-3629.[Abstract/Free Full Text]
    14. 14
  14. DiStefano, T. D., J. M. Gossett, and S. H. Zinder. 1991. Reductive dechlorination of high concentrations of tetrachloroethene to ethene by an anaerobic enrichment culture in the absence of methanogenesis. Appl. Environ. Microbiol. 57:2287-2292.[Abstract/Free Full Text]
    15. 15
  15. Dojka, M. A., P. Hugenholtz, S. K. Haack, and N. R. Pace. 1998. Microbial diversity in a hydrocarbon- and chlorinated-solvent-contaminated aquifer undergoing intrinsic bioremediation. Appl. Environ. Microbiol. 64:3869-3877.[Abstract/Free Full Text]
    16. 16
  16. 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]
    17. 17
  17. 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]
    18. 18
  18. Duhamel, M., K. Mo, and E. A. Edwards. 2004. Characterization of a highly enriched Dehalococcoides-containing culture that grows on vinyl chloride and trichloroethene. Appl. Environ. Microbiol. 70:5538-5545.[Abstract/Free Full Text]
    19. 19
  19. Fennell, D. E., and J. M. Gossett. 1998. Modeling the production of and competition for hydrogen in a dechlorinating culture. Environ. Sci. Technol. 32:2450-2460.
    20. 20
  20. Fennell, D. E., J. M. Gossett, and S. H. Zinder. 1997. Comparison of butyric acid, ethanol, lactic acid, and propionic acid as hydrogen donors for the reductive dechlorination of tetrachloroethene. Environ. Sci. Technol. 31:918-926.
    21. 21
  21. Flynn, S. J., F. E. Loffler, and J. M. Tiedje. 2000. Microbial community changes associated with a shift from reductive dechlorination of PCE to reductive dechlorination of cis-DCE and VC. Environ. Sci. Technol. 34:1056-1061.
    22. 22
  22. Freeborn, R. A., K. A. West, V. K. Bhupathiraju, S. Chauhan, B. G. Rahm, R. E. Richardson, and L. Alvarez-Cohen. 2005. Phylogenetic analysis of TCE-dechlorinating consortia enriched on a variety of electron donors. Environ. Sci. Technol. 39:8358-8368.[Medline]
    23. 23
  23. Reference deleted.
    24. 24
  24. Freedman, D. L., and J. M. Gossett. 1989. Biological reductive dechlorination of tetrachloroethylene and trichloroethylene to ethylene under methanogenic conditions. Appl. Environ. Microbiol. 55:2144-2151.[Abstract/Free Full Text]
    25. 25
  25. Fung, J. M., R. M. Morris, L. Adrian, and S. H. Zinder. 2007. Expression of reductive dehalogenase genes in Dehalococcoides ethenogenes strain 195 growing on tetrachloroethene, trichloroethene, or 2,3-dichlorophenol. Appl. Environ. Microbiol. 73:4439-4445.[Abstract/Free Full Text]
    26. 26
  26. Futamata, H., N. Yoshida, T. Kurogi, S. Kaiya, and A. Hiraishi. 2007. Reductive dechlorination of chloroethenes by Dehalococcoides-containing cultures enriched from a polychlorinated-dioxin-contaminated microcosm. Isme J. 1:471-479.[CrossRef][Medline]
    27. 27
  27. Gloeckner, F. O., R. Amann, A. Alfreider, J. Pernthaler, R. Psenner, K. Trebesius, and K. H. Schleifer. 1996. An in situ hybridization protocol for detection and identification of planktonic bacteria. Syst. Appl. Microbiol. 19:403-406.
    28. 28
  28. Gu, A. Z., B. P. Hedlund, J. T. Staley, S. E. Strand, and H. D. Stensel. 2004. Analysis and comparison of the microbial community structures of two enrichment cultures capable of reductively dechlorinating TCE and cis-DCE. Environ. Microbiol. 6:45-54.[CrossRef][Medline]
    29. 29
  29. Hales, B. A., C. Edwards, D. A. Ritchie, G. Hall, R. W. Pickup, and J. R. Saunders. 1996. Isolation and identification of methanogen-specific DNA from blanket bog feat by PCR amplification and sequence analysis. Appl. Environ. Microbiol. 62:668-675.[Abstract]
    30. 30
  30. Harkness, M. R., A. A. Bracco, M. J. Brennan, K. A. Deweerd, and J. L. Spivack. 1999. Use of bioaugmentation to stimulate complete reductive dechlorination of trichloroethene in dover soil columns. Environ. Sci. Technol. 33:1100-1109.
    31. 31
  31. He, J. Z., V. F. Holmes, P. K. H. Lee, and L. Alvarez-Cohen. 2007. Influence of vitamin B-12 and cocultures on the growth of Dehalococcoides isolates in defined medium. Appl. Environ. Microbiol. 73:2847-2853.[Abstract/Free Full Text]
    32. 32
  32. Hoelscher, T., H. Goerisch, and L. Adrian. 2003. Reductive dehalogenation of chlorobenzene congeners in cell extracts of Dehalococcoides sp. strain CBDB1. Appl. Environ. Microbiol. 69:2999-3001.[Abstract/Free Full Text]
    33. 33
  33. Hohnstock-Ashe, A. M., S. M. Plummer, R. M. Yager, P. Baveye, and E. L. Madsen. 2001. Further biogeochemical characterization of a trichloroethene-contaminated fractured dolomite aquifer: electron source and microbial communities involved in reductive dechlorination. Environ. Sci. Technol. 35:4449-4456.[Medline]
    34. 34
  34. Holmes, V. F., J. Z. 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]
    35. 35
  35. Holoman, T. R. P., M. A. Elberson, L. A. Cutter, H. D. May, and K. R. Sowers. 1998. Characterization of a defined 2,3,5,6-tetrachlorobiphenyl-ortho-dechlorinating microbial community by comparative sequence analysis of genes coding for 16S rRNA. Appl. Environ. Microbiol. 64:3359-3367.[Abstract/Free Full Text]
    36. 36
  36. Johnson, D. R., P. K. H. Lee, V. F. Holmes, and L. Alvarez-Cohen. 2005. An internal reference technique for accurately quantifying specific mRNAs by real-time PCR with a application to the tceA reductive dehalogenase gene. Appl. Environ. Microbiol. 71:3866-3871.[Abstract/Free Full Text]
    37. 37
  37. Johnson, D. R., P. K. H. Lee, V. F. Holmes, A. C. Fortin, and L. Alvarez-Cohen. 2005. Transcriptional expression of the tceA gene in a Dehalococcoides-containing microbial enrichment. Appl. Environ. Microbiol. 71:7145-7151.[Abstract/Free Full Text]
    38. 38
  38. Kassenga, G., J. H. Pardue, W. M. Moe, and K. S. Bowman. 2004. Hydrogen thresholds as indicators of dehalorespiration in constructed treatment wetlands. Environ. Sci. Technol. 38:1024-1030.[Medline]
    39. 39
  39. Krajmalnik-Brown, R., Y. Sung, K. M. Ritalahti, F. M. Saunders, and F. E. Loffler. 2007. Environmental distribution of the trichloroethene reductive dehalogenase gene (tceA) suggests lateral gene transfer among Dehalococcoides. FEMS Microbiol. Ecol. 59:206-214.[CrossRef][Medline]
    40. 40
  40. Lee, P. K. H., D. R. Johnson, V. F. Holmes, J. Z. He, and L. Alvarez-Cohen. 2006. Reductive dehalogenase gene expression as a biomarker for physiological activity of Dehalococcoides spp. Appl. Environ. Microbiol. 72:6161-6168.[Abstract/Free Full Text]
    41. 41
  41. Logan, B. E., and J. R. Hunt. 1987. Advantages to microbes of growth in permeable aggregates in marine systems. Limnol. Oceanogr. 32:1034-1048.
    42. 42
  42. Ludwig, W., O. Strunk, R. Westram, L. Richter, H. Meier, Yadhukumar, A. Buchner, T. Lai, S. Steppi, G. Jobb, W. Forster, I. Brettske, S. Gerber, A. W. Ginhart, O. Gross, S. Grumann, S. Hermann, R. Jost, A. Konig, T. Liss, R. Lussmann, M. May, B. Nonhoff, B. Reichel, R. Strehlow, A. Stamatakis, N. Stuckmann, A. Vilbig, M. Lenke, T. Ludwig, A. Bode, and K. H. Schleifer. 2004. ARB: a software environment for sequence data. Nucleic Acids Res. 32:1363-1371.[Abstract/Free Full Text]
    43. 43
  43. Macbeth, T. W., D. E. Cummings, S. Spring, L. M. Petzke, and K. S. Sorenson. 2004. Molecular characterization of a dechlorinating community resulting from in situ biostimulation in a trichloroethene-contaminated deep, fractured basalt aquifer and comparison to a derivative laboratory culture. Appl. Environ. Microbiol. 70:7329-7341.[Abstract/Free Full Text]
    44. 44
  44. Maymo-Gatell, X., Y. T. Chien, J. M. Gossett, and S. H. Zinder. 1997. Isolation of a bacterium that reductively dechlorinates tetrachloroethene to ethene. Science 276:1568-1571.[Abstract/Free Full Text]
    45. 45
  45. Maymo-Gatell, X., V. Tandoi, J. M. Gossett, and S. H. Zinder. 1995. Characterization of an H-2-utilizing enrichment culture that reductively dechlorinates tetrachloroethene to vinyl chloride and ethene in the absence of methanogenesis and acetogenesis. Appl. Environ. Microbiol. 61:3928-3933.[Abstract]
    46. 46
  46. Morris, R. M., S. Sowell, D. Barofsky, S. Zinder, and R. Richardson. 2006. Transcription and mass-spectroscopic proteomic studies of electron transport oxidoreductases in Dehalococcoides ethenogenes. Environ. Microbiol. 8:1499-1509.[CrossRef][Medline]
    47. 47
  47. Morris, R. M., M. S. Rappe, E. Urbach, S. A. Connon, and S. J. Giovannoni. 2004. Prevalence of the Chloroflexi-related SAR202 bacterioplankton cluster throughout the mesopelagic zone and deep ocean. Appl. Environ. Microbiol. 70:2836-2842.[Abstract/Free Full Text]
    48. 48
  48. Morris, R. M., J. M. Fung, B. G. Rahm, S. Zhang, D. L. Freedman, S. H. Zinder, and R. E. Richardson. 2007. Comparative proteomics of Dehalococcoides spp. reveals strain-specific peptides associated with activity. Appl. Environ. Microbiol. 73:320-326.[Abstract/Free Full Text]
    49. 49
  49. Nakatsu, C. H., V. Torsvik, and L. Ovreas. 2000. Soil community analysis using DGGE of 16S rDNA polymerase chain reaction products. Soil Sci. Soc. Am. J. 64:1382-1388.[Abstract/Free Full Text]
    50. 50
  50. Nielson, R. B. 1999. PhD Thesis. University of California, Berkeley, Berkeley, CA.
    51. 51
  51. Pernthaler, A., and R. Amann. 2004. Simultaneous fluorescence in situ hybridization of mRNA and rRNA in environmental bacteria. Appl. Environ. Microbiol. 70:5526-5533.
    52. 52
  52. Rahm, B. G., and R. E. Richardson. 2008. Correlation of respiratory gene expression levels and pseudo-steady-state PCE respiration rates in Dehalococcoides ethenogenes. Environ. Sci. Technol. 42:416-421.[Medline]
    53. 53
  53. Rahm, B. G., R. M. Morris, and R. E. Richardson. 2006. Temporal expression of respiratory genes in an enrichment culture containing Dehalococcoides ethenogenes. Appl. Environ. Microbiol. 72:5486-5491.[Abstract/Free Full Text]
    54. 54
  54. Richardson, R. E., V. K. Bhupathiraju, D. L. Song, T. A. Goulet, and L. Alvarez-Cohen. 2002. Phylogenetic characterization of microbial communities that reductively dechlorinate TCE based upon a combination of molecular techniques. Environ. Sci. Technol. 36:2652-2662.[Medline]
    55. 55
  55. Rossetti, S., L. L. Blackall, M. Majone, P. Hugenholtz, J. J. Plumb, and V. Tandoi. 2003. Kinetic and phylogenetic characterization of an anaerobic dechlorinating microbial community. Microbiology 149:459-469.[Abstract/Free Full Text]
    56. 56
  56. Reference deleted.
    57. 57
  57. Smatlak, C. R., J. M. Gossett, and S. H. Zinder. 1996. Comparative kinetics of hydrogen utilization for reductive dechlorination of tetrachloroethene and methanogenesis in an anaerobic enrichment culture. Environ. Sci. Technol. 30:2850-2858.
    58. 58
  58. Smidt, H., and W. M. de Vos. 2004. Anaerobic microbial dehalogenation. Annu. Rev. Microbiol. 58:43-73.[Medline]
    59. 59
  59. Stams, A. J. M. 1994. Metabolic interactions between anaerobic-bacteria in methanogenic environments. Antonie van Leeuwenhoek Int. J. Gen. Mol. Microbiol. 66:271-294.[CrossRef]
    60. 60
  60. Thiele, J. H., M. Chartrain, and J. G. Zeikus. 1988. Control of interspecies electron flow during anaerobic digestion: role of floc formation in syntrophic methanogenesis. Appl. Environ. Microbiol. 54:10-19.[Abstract/Free Full Text]
    61. 61
  61. Waller, A. S., R. Krajmalnik-Brown, F. E. Loffler, 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]
    62. 62
  62. Wang, Q., G. M. Garrity, J. M. Tiedje, and J. R. Cole. 2007. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 73:5261-5267.[Abstract/Free Full Text]
    63. 63
  63. Yang, Y. R., and J. Zeyer. 2003. Specific detection of Dehalococcoides species by fluorescence in situ hybridization with 16S rRNA-targeted oligonucleotide probes. Appl. Environ. Microbiol. 69:2879-2883.[Abstract/Free Full Text]
    64. 64
  64. Yang, Y. R., M. Pesaro, W. Sigler, and J. Zeyer. 2005. Identification of microorganisms involved in reductive dehalogenation of chlorinated ethenes in an anaerobic microbial community. Water Res. 39:3954-3966.[Medline]
    65. 65
  65. Yilmaz, L. S., and D. R. Noguera. 2004. Mechanistic approach to the problem of hybridization efficiency in fluorescent in situ hybridization. Appl. Environ. Microbiol. 70:7126-7139.[Abstract/Free Full Text]

Applied and Environmental Microbiology, November 2008, p. 6709-6719, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.00445-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rowe, A. R.
Right arrow Articles by Richardson, R. E.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rowe, A. R.
Right arrow Articles by Richardson, R. E.
Agricola
Right arrow Articles by Rowe, A. R.
Right arrow Articles by Richardson, R. E.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»