Geochemistry and microbial diversity of a trichloroethene-contaminated Superfund site undergoing intrinsic in situ reductive dechlorination

FEMS Microbiology Ecology 40 (2002) 123^134

www.fems-microbiology.org

Geochemistry and microbial diversity of a trichloroethene-contaminated Superfund site undergoing intrinsic in situ reductive dechlorination Mary Lowe a , Eugene L. Madsen b , Karen Schindler a , Courtney Smith a , Scott Emrich a , Frank Robb c , Rolf U. Halden d; a Physics Department, Loyola College, Baltimore, MD 21210, USA Department of Microbiology, Cornell University, Ithaca, NY 14853, USA Center of Marine Biotechnology, University of Maryland Biotechnology Institute, Baltimore, MD 21202, USA Environmental Protection Department, Lawrence Livermore National Laboratory, Livermore, CA 94551, USA b

c d

Received 11 October 2001; received in revised form 1 February 2002; accepted 5 February 2002 First published online 22 April 2002

Abstract This study explored the geochemistry and microbial diversity of a Superfund site containing trichloroethene (TCE) and an unusual copollutant, tetrakis(2-ethylbutoxy)silane. Geochemical analysis of contaminated groundwater indicated subsurface anaerobiosis, reductive dechlorination of TCE to predominantly cis-1,2-dichloroethene, and (transient) accumulation of 2-ethylbutanol and 2-ethylbutyrate as a result of tetrakis(2-ethylbutoxy)silane breakdown. Comparative analysis of 106 16S rDNA and 61 16S^23S rDNA intergenic spacer region sequences ^ obtained from pristine and contaminated groundwater via DNA extraction, PCR amplification, cloning and sequencing ^ revealed that the contaminated groundwater featured (i) a distinct microbial community, (ii) reduced species diversity, (iii) various anaerobes, and (iv) bacteria closely related to the TCE-dechlorinating, dichloroethene-accumulating genus Dehalobacter, whereas (v) the TCE-dechlorinating, ethene-producing species Dehalococcoides ethenogenes was not detectable. Thus, geochemical and molecular biological results were in excellent agreement in this first ecological field study linking in situ reductive dechlorination of TCE to metabolism of tetraalkoxysilanes. 7 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. Keywords : 16S rDNA; 16S^23S intergenic spacer region ; Tetraalkoxysilane; Bioremediation ; Reductive dechlorination

1. Introduction Bioremediation of contaminated subsurface environments by microorganisms is an emerging technology [1]. It is becoming recognized that the lack of su⁄cient microbial diversity at contaminated sites may explain why biodegradable pollutants fail to be biotransformed in situ [2]. While new ¢ndings on the metabolic routes and bottlenecks of degradation are still accumulating [3], it is already clear that the capacity of indigenous microbial populations to adapt to the presence of toxic pollutants and to biode-

* Corresponding author. Present address: Department of Environmental Health Sciences, Bloomberg School of Public Health, Johns Hopkins University, 615 N. Wolfe Street, Suite W6001, Baltimore, MD 212052103, USA. Tel.: +1 (410) 955 2609 ; Fax: +1 (410) 955 9334. E-mail address : [email protected] (R.U. Halden).

grade these compounds may be the most important factor in determining the fate of subsurface contaminants (for a review, see [4]). In situ reductive dechlorination is considered to be the most promising mechanism for bioremediation of chloroethene spill sites [5]. When trichloroethene (TCE) represents the primary contaminant, sequential reductive dechlorination may yield cis-1,2-dichloroethene (cis-DCE), minor quantities of trans-1,2-dichloroethene, vinyl chloride, and ultimately non-toxic, chlorine-free end products in the form of ethene and ethane [6,7]. Many of these transformations can be performed cometabolically by methanogenic, acetogenic and sulfate-reducing microorganisms that produce enzymes containing transition metal cofactors (see [8,9] and references therein). The reactions proceed at orders of magnitude faster rates, however, when carried out by respiratory organochlorine-reducing

0168-6496 / 02 / $22.00 7 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII : S 0 1 6 8 - 6 4 9 6 ( 0 2 ) 0 0 2 2 9 - 5

FEMSEC 1354 23-5-02

124

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

bacteria that gain energy during the process. The latter are obligate anaerobes of phylogenetically diverse origin and include strains of Dehalobacter restrictus, Dehalospirillum multivorans, Desulfuromonas sp. and Dehalococcoides ethenogenes [7,10^12]. Detection of any or all of these microorganisms at a chloroethene spill site would represent one important line of evidence indicating the potential applicability of in situ bioremediation to the site. Certainly, many other factors need to be considered by regulatory agencies before committing to biological cleanup : geochemical site data, the availability of electron donor compounds suitable for driving microbial dechlorination reactions, and types and abundances of chloroethene daughter products [1,13]. The chloroethene spill site investigated in this report ^ Lawrence Livermore National Laboratory’s (LLNL) Building 834 Operable Unit at Site 300 ^ is interesting from a microbiological standpoint for at least two reasons. First, maximum groundwater concentrations of the toxic contaminant TCE historically have been close to the point of saturation (V1084 mg l31 ) [14], thereby creating an unusually challenging environment for indigenous microorganisms. Second, TCE was spilled together with tetrakis(2-ethylbutoxy)silane (TKEBS), a silicon-based lubri-

cant that potentially may support both the anaerobic [15,16] and aerobic [17] metabolism of chloroethenes. Previous laboratory experiments had shown that the four branched alkane side chains of the water-insoluble TKEBS can be released under ambient conditions via slow hydrolysis [17]. Additional evidence from groundwater microcosm studies suggests that the liberated 2-ethylbutanol can be fermented to 2-ethylbutyrate, acetate and hydrogen [15,18]. If these anaerobic fermentation reactions occur in the ¢eld at a relevant scale, TKEBS may support reductive dechlorination of chlorinated ethenes by serving as a longterm electron donor source. In order to gain insight into the microbial diversity, physiology, and geochemistry of the study site, two monitoring wells were selected for sampling and analysis. The selected contaminated well is located in the source area of the site where mixtures of TCE and TKEBS were spilled. The second well, which served as a background well, is situated outside of the chloroethene-impacted area. Specific objectives of the present study were to (i) search for evidence of ongoing in situ reductive dechlorination, (ii) analyze the microbial community composition and identify population shifts that may have occurred in response to the presence of TCE and TKEBS, and (iii) determine the relative usefulness of two culture-independent microbial pro¢ling tools that are based on the 16S rRNA gene and the 16S^23S rDNA intergenic spacer region (ISR).

2. Materials and methods 2.1. Site description

Fig. 1. Plan view of the study location containing the primary contaminants TCE and TKEBS as well as the TCE metabolite cis-DCE.

Site 300 is an active explosives test site situated in the remote Altamont Hills approximately 100 hwy km southeast of San Francisco. It is operated by the Department of Energy and listed as an EPA Superfund site. The area under study, the Building 834 Complex, is located in the southeastern corner of Site 300. It was constructed in the 1950s for conducting thermal shock and humidity testing of weapons components. The building complex is located on an isolated hilltop at an elevation of 311 m. In the vicinity of Well W-834-D3 (Fig. 1), periodic leakage of a mixture of TCE (100^30%) and TKEBS (0^70%) to the subsurface from a piping system resulted in the environmental release of approximately 3000 kg of TCE and an unknown quantity of the unregulated TKEBS compound. The above mixture of two non-aqueous phase liquids is heavier than water, which allowed both contaminants to jointly penetrate the water table and migrate deep into the subsurface. Following discovery of soil and groundwater contamination in the early 1980s, the piping system was dismantled and the facility abandoned. The hydrogeology of the Building 834 area is complex. The source area ^ located on a hilltop ^ is underlain by a shallow (V15 m) perched water-bearing zone of variable

FEMSEC 1354 23-5-02

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

thickness ( 6 0.1^4 m). A contour plot, generated from groundwater compliance monitoring data using Earthvision software, indicates that the TCE plume measures about 500 m by 200 m (Fig. 1). It occurs in a semi-consolidated conglomerate of sand and gravel that is underlain by a bedrock sequence containing several layers of low permeability. The most shallow of these aquitards (Tps clay), is located 10^20 m below the surface and prevents the downward migration of contaminants through the various layers of unsaturated sandstone to the regional aquifer, located some 85 m below. Lateral migration of dissolved contaminants is restricted by the limited extent of saturation and the topography. Aqueous concentrations of TCE typically are in the mg l31 range whenever groundwater is encountered. Two groundwater monitoring wells, W-834-D3 and W834-T5 (hereafter referred to as D3 and T5, respectively), were selected for this study (Fig. 1). D3 is located in an area where both TCE and TKEBS were spilled to the ground; no known potential carbon and energy sources other than TKEBS exist in this location. The well has a depth of 10 m and is screened in the conglomerate overlaying the initial clay layer (perching horizon). T5 is located 430 m to the south, outside of the TCE plume. This well has a depth of 24 m and is also screened in the conglomerate layer. 2.2. Chemical and geochemical site characterization Groundwater samples, collected at the wellhead in certi¢ed glass vials, were shipped on ice to B.C. Laboratories, Inc. (Bakers¢eld, CA, USA) and analyzed for volatile organic compounds by purge and trap gas chromatography/ mass spectrometry (GC/MS) using U.S. EPA method 601. Concentrations of dissolved gases were determined by Microseeps, Inc. (Pittsburgh, PA, USA) via GC/MS analysis of headspace gas samples that were generated in the ¢eld using the bubble strip method [19]. Concentrations of TKEBS and its transformation products were determined by liquid^liquid extraction and GC/MS analysis as described previously [17]. Values of pH, Eh, and dissolved oxygen were measured in the ¢eld with parameter-speci¢c electrodes as described elsewhere [20]. Concentrations of inorganic salts and sul¢de were determined spectrophotometrically in the ¢eld using CHEMetrics (Calverton, VA, USA) test kits per manufacturer’s directions. A database of regulatory compliance monitoring data was used for generating plume contour maps and for determining historical maximum concentrations. These analytical data were produced by certi¢ed commercial laboratories using standard methods. 2.3. Collection of groundwater samples for microbial pro¢ling Groundwater samples were collected on August 6, 1998

125

from wells T5 and D3. Four liters of groundwater, obtained from the wellhead, were passed through SterivexGP ¢lters (0.22 Wm pore size; Millipore Corp., Bedford, MA) and immediately placed on ice. The ¢lters were shipped on dry ice and stored at 380‡C. To remove the microorganisms, the ¢lters were agitated with a vortex overnight at 4‡C, and then back£ushed twice with diluted phosphate bu¡er (1 part M9 bu¡er [21] plus 2 parts water). 2.4. Extraction of community DNA A direct lysis procedure was adapted from [22]. Samples ( 9 300 Wl) were suspended in 500 Wl of bu¡er A (500 mM Tris^HCl, pH 8.0, 100 mM NaCl, 1 mM sodium citrate). Next, 100 Wl of 10 mg ml31 lysozyme were added and samples were incubated for 1 h at 37‡C. Following initial incubation, 100 Wl of 10 mg ml31 of activated proteinase K were added and incubated for an additional 30 min. Next, 500 Wl of lysis bu¡er (200 mM Tris^HCl, pH 8.0/100 mM NaCl, 4% (w/v) SDS/10% (w/v) aminosalicylic acid) were added. Three freeze/thaw cycles were performed using a dry ice^ethanol bath and a 65‡C water bath. Finally, the community DNA was puri¢ed using saturated phenol, pH 7.9 (Ambion, Austin, TX, USA), followed by addition of a mixture of phenol, chloroform and isoamyl alcohol (24:24:1). Organic and DNA phases were separated by Phase Lock Gel (Eppendorf, Westbury, NY, USA). The genomic DNA was precipitated with 3 M sodium acetate (0.1 of total volume) and 1 vol. of isopropanol. Pellet paint (Novagen, Madison, WI, USA) was added to the precipitate for easy pellet identi¢cation. The pellet was then washed in 100% ethanol, vacuum-dried, and resuspended in 25 Wl sterile water. The concentration of puri¢ed DNA was estimated from its optical density at 260 nm and 255 nm (pellet paint maximum) using a Beckman DU640 spectrophotometer. 2.5. PCR ampli¢cations To amplify the 16S rDNA gene, 1.0 Wl of extracted community DNA 6 200 ng was added in a total reaction volume of 100 Wl containing 2.5 U Amplitaq DNA polymerase (Perkin-Elmer, Woburn, MA, USA), 1UGeneAmp PCR bu¡er ; 200 WM each dNTP, 0.6 WM each bacterial primer 8F (AGAGTTTGATCCTGGCTCAG) [23] and 1492R (GGTTACCTTGTTACGACTT) [24]. The tubes were placed in a thermal cycler equilibrated at 94‡C, followed by a 2-min denaturation step at 94‡C ; 25 cycles of denaturation (30 s at 94‡C), annealing (45 s at 55‡C), and extension (60 s at 72‡C); and a ¢nal extension for 8 min at 72‡C. A 1% agarose gel stained with ethidium bromide showed a single band at V1500 bp. No band was visible for the negative control (reagents only; no DNA template).

FEMSEC 1354 23-5-02

126

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

To amplify the ISR, 1 Wl of community DNA was added to a total reaction volume of 30 Wl containing 2.5 U Taq (Fisher Biotech), 1UPCR bu¡er, 55 WM each dNTP, 20 pmol primer R2 (5P-GGGWGAAGTCGTAACAAG-3P) and 20 pmol primer R5 (5P-TTAGCACGTCCTTCATCGCC-3P). R2 overlaps 1492R by 13 bases. R5 is located within the ¢rst 100 bases of the 23S rDNA coding region. The thermal cycling protocol consisted of a 5-min denaturation at 94‡C, followed by addition of Taq (hotstart); 35 cycles of denaturation (30 s at 94‡C), annealing (30 s at 50‡C), and extension (1.5 min at 72‡C); followed by a ¢nal extension for 5 min at 72‡C. Separation of the products on a 1% agarose gel showed a continuous range of product lengths with prominent bands at 930, 590, and 520 bp for D3, and weak bands at 500 and 450 bp for T5. No band was visible for the negative control containing no DNA template. Prior to cloning, all amplicons were puri¢ed using the Geneclean III Kit (Bio 101) according to the manufacturer’s protocol. 2.6. Cloning Cloning was accomplished with the TOPO-TA cloning kit (Invitrogen version J and earlier) according to the manufacturer’s instructions using 4 Wl of puri¢ed PCR product, 1 Wl of pCR2.1-TOPO plasmid vector, and chemically competent TOP10FP One Shot Escherichia coli. Clones were picked at random and grown overnight in Luria^Bertani (LB) broth containing 150 Wg ml31 ampicillin. For long-term storage, 0.05 ml dimethylsulfonate was added to 0.45 ml of the liquid culture, and the mixture was incubated at room temperature for 30 min prior to storage at 380‡C. Alternatively, the cells were stored in an LB broth/glycerol mixture (85:15) at 380‡C. 2.7. Isolation of plasmid DNA Plasmid DNA was extracted and puri¢ed using a QIAprep Spin Miniprep kit (Qiagen, Valencia, CA, USA) according to the manufacturer’s protocol. The optional column wash was performed. Plasmid DNA was eluted with 50 Wl of sterile water. The presence of an insert was determined by running a restriction digest using EcoRI (Promega, Madison, WI, USA) and 1 Wl of puri¢ed pDNA on a 1% agarose gel. 2.8. Sequencing All plasmid DNA samples (0.5^1.0 Wg) were sequenced with a Perkin-Elmer ABI 373 using primer T7 (5P-TAATACGACTCACTTAGGG-3P). A subset was also sequenced with universal M13 Reverse Primer (5P-CAGGAAACAGCTATGAC) or 338F (5P-TCCTACGGGAGGCAGC) [23].

2.9. Sequence analysis A set of C++ software utilities was developed to process the sequences rapidly. All sequences were placed on the positive strand and contained at least one of the PCR primers. Vector sequences were removed. After editing, each sequence was typically 700^800 bp in length, spanning approximately half of the 16S rDNA and often the entire ISR. Full-length 16S sequences were constructed either by hand or by use of the CAP2 contig assembly software (http://hercules.tigem.it/ASSEMBLY/assemble. html). The software was set to optimize reads s 450 bp with a threshold for sequence assembly of 80% identity and the feature to remove ‘bad’ 5P and 3P ends. All sequences for each well were placed in a FASTA-format ¢le and analyzed by BLAST in September 2000 using the Win32 version of the NIH Blastcl3 network client (ftp :// ncbi.nlm.nih.gov/blast/network/netblast/). The Blastn program utilized the nr database and reported output in hypertext format. The quality of the microbial identi¢cation was determined from the bit score, percent identity, and segment length. Sequences of 16S rDNA which did not meet the identi¢cation criteria described in Section 3 were tested with CHIMERA_CHECK Version 2.7 (http:// www.cme.msu.edu/RDP/html/analyses.html) ; no chimeric sequences were found. In addition, 16S and ISR sequences from Well T5 were compared with the corresponding sequences from Well D3 using Standalone Blast and the ‘formatdb’ and ‘blastall’ commands. Alignments and dendrograms were generated using the program PileUp from the Wisconsin Package (Genetics Computer Group, Madison, WI, USA). Input ¢les consisted of all sequences containing the ¢rst half of the 16S rRNA gene. Complete 16S rDNA sequences were truncated in the middle. For the ISR sequences, the input ¢les consisted of all complete and incomplete sequences. The gap creation and extension penalties were 4 and 0, respectively. The branch lengths in the dendrograms (Figs. 2 and 3) are linearly proportional to the distances between sequences with the maximum normalized distance of 1 at the root (left) and 0 at the right. Reference sequences for the 16S and ISR regions were obtained from the Institute of Genomic Research (TIGR, http://www.tigr.org) or GenBank. From TIGR, only E. coli K-12 was ¢nished. The un¢nished sequences, as of July/August 2000, were Geobacter sulfurreducens, Desulfovibrio vulgaris, and D. ethenogenes. All other reference strains were obtained from GenBank. All sequences were trimmed to have the closest approximation to the primer pairs 8F/1492R and R2/R5 at the terminal ends. If a primer could not be found, then no trimming occurred. 2.10. Nucleotide sequence accession numbers The ISR and 16S rRNA gene sequences appear in Gen-

FEMSEC 1354 23-5-02

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

127

Fig. 2. Dendrogram of 16S rDNA sequences from pristine groundwater (Well T5; boldface type and pre¢x ‘t’) and from TCE-contaminated groundwater (Well D3; plain type and pre¢x ‘d’). Reference strains (ref) provide a network for examining the lineage of isolated clones. Reference sequences obtained from GenBank have accession numbers, whereas those obtained from TIGR do not. Tentative taxonomic a⁄liation was assigned to cloned sequences matching GenBank entries (see text for criteria).

Bank under accession Nos. AF422492 to AF422528 (ISR, T5), AF422529 to AF422538 and AF422540 to AF422575 (ISR, D3), AF422576 to AF422620 (16S, T5), and AF422621 to AF422689 (16S, D3).

the number of individuals (i.e., clones). Other indices included : Shannon^Weaver (H = (C/N)(N log10 N34 ni log10 ni )) and Evenness (e = H/log S), where C = 2.3, and ni is the number of individuals in the ith species [26^28].

2.11. Diversity measures 3. Results The clone coverage [23,25] is given by c = [13(S/ N)]U100%, where S is the number of species and N is

Geochemical analyses conducted on well waters gath-

FEMSEC 1354 23-5-02

128

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

ered from inside and outside the TCE-contaminated area of the study site (Table 1) provided clear evidence for intrinsic in situ reductive dechlorination of chloroethenes. TCE and the co-contaminating silicon-based oil, TKEBS, were present in Well D3, but not in Well T5. Also, within the contaminated zone, undetectable levels of oxygen and a negative Eh value indicated highly reducing conditions. In contrast, oxygen and Eh readings in the pristine Well T5 indicated oxidizing conditions. Depletion of nitrate and sulfate in D3 relative to T5 also was consistent with alkoxysilane-induced anaerobiosis. Furthermore, the presence of 2-ethylbutanol and 2-ethylbutyrate ^ both representing transformation products of TKEBS previously observed in controlled laboratory experiments [15,17] ^ and the accumulation of acetate in D3 suggested that the microorganisms native to this subsurface habitat had adapted to the presence of TCE and were performing fermentative reactions including anaerobic transformation of the tetraalkoxysilane compound. The key observation indicating in situ reductive dechlorination within the contaminated zone was the detection of TCE daughter products that serve as signature compounds for anaerobic bioattenuation, most importantly cis-DCE, ethene and ethane (Table 1). The latter two were at very low concentrations. None of these three compounds were released during site operations; rather, their likely origin is via in situ microbial metabolism, driven indirectly by TKEBS hydrolysis [17] as observed earlier in laboratory

studies [15]. Fig. 1 depicts the spatial distribution of contaminants at the site. The TCE plume has extended in a southwesterly direction approximately 400 m beyond the initial point of release. The plume of cis-DCE ^ the ¢rst daughter product in the reductive dechlorination sequence ^ has not migrated as extensively as the TCE, presumably due to its relatively greater susceptibility to both anaerobic and aerobic [17] biological breakdown processes. Increases in chloride concentrations can serve as an additional indicator of in situ reductive dechlorination. However, the chloride concentration in D3 was not elevated relative to T5 (data not shown), probably because of site heterogeneity and patchy releases of chloride salts throughout the LLNL complex. Given the clear geochemical evidence for in situ reductive dechlorination, we undertook a non-culture-based survey of nucleic acid sequences re£ecting the microbial community composition in the two wells. Particular attention was directed toward: (i) contrasts between communities residing in contaminated versus non-contaminated zones; (ii) links between phylotypes previously shown to be active in reductive dechlorination and related anaerobic microbial processes, and (iii) insights that might develop by comparing data sets based on 16S rDNA and ISR. Sequencing was accomplished for a total of 45 and 61 16S rDNA inserts from T5 and D3, respectively, and for 26 and 35 ISR inserts from the two respective locations. All of these were used in diversity calculations (see below).

Table 1 Chemistry and geochemistry of perched groundwater from two monitoring wells located inside and outside of the TCE-contaminated zonea Analyte Primary contaminants and metabolites TCE (historical maximum, February 9, 1993) cis-1,2-DCE (historical maximum, September 25, 1997) Vinyl chloride Ethene (historical maximum, May 9, 2000) Ethane (historical maximum, May 9, 2000) Secondary contaminants and metabolites TKEBS (historical maximum, December 21, 1995) 2-Ethylbutanol 2-Ethylbutyrate Acetate Hydrogen (historical maximum, May 9, 2000) Geochemical indicators pH Eh Dissolved oxygen Nitrate Nitrite Sulfate Sul¢de

Unit

Contaminated Well W-834-D3

Control Well W-834-T5

mg l31

56 X 13 800 16 X 11 390 BD ND 0.0013 ND 0.0004

BD

BD

nM

11 X 8 7300 (LNAPL) 7X9 94 X 141 21 X 32 ND 7.4

mV mg l31 mg l31 mg l31 mg l31 mg l31

7.2 X 0.1 3121 X 87 62 65 6 0.5 13 X 5 61

mg l31 mg l31 mg l31 mg l31 mg l31 mg l31 mg l31 mg l31

a

BD BD ND BD

BD BD BD ND

7.7 X 0.2 178 X 46 6.3 X 0.4 137 X 6 6 0.5 30 X 2 61

Unless otherwise noted, reported values represent average concentrations ( X 1 S.D.) of three measurements made between July 28 and September 23, 1998. BD, below detection ; ND, not determined ; LNAPL, light non-aqueous phase liquid.

FEMSEC 1354 23-5-02

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

The dendrograms shown in Figs. 2 and 3 depict the relationships among 16S rDNA and ISR sequences from uncontaminated and contaminated well waters. Sixteen reference strains (designated by ‘ref’) were selected to (i) represent common bacteria, (ii) correspond to clones with high BLAST scores, and (iii) represent cultured bacteria known to carry out reductive dechlorination, i.e., D. multivorans, D. restrictus, Desul¢tobacterium sp., D. ethenogenes, Dehalococcoides sp. strain CBDB1, Desulfuromonas chloroethenica, and D. vulgaris [10^12]. ISR sequences were available for only 10 of the 16 selected reference strains. In the 16S rDNA tree (Fig. 2), a tentative taxonomic a⁄liation was made if the BLAST raw alignment score was v 400 and the percent similarity was v 90%. In prac-

Fig. 3. Dendrogram of 16S^23S rDNA ISR sequences from pristine groundwater (Well T5; boldface type and pre¢x ‘t’) and from TCE-contaminated groundwater (Well D3; plain type and pre¢x ‘d’). Reference strains (ref) provide a network for examining the lineage of isolated clones.

129

tice, this resulted in segment lengths v 346 bases. For the ISR tree (Fig. 3), a percentage of v 90% and a segment length of v 150 bp were used. For cases where several organisms could meet the identi¢cation criteria, only the organism with the highest score is indicated. Also shown in Figs. 2 and 3 is an additional BLAST statistic of the form n/m = p%, where m is the segment length (i.e., length of the aligned region), n is the number of exact matches within the segment, and p is the percentage of identical matches. Fig. 2 portrays 16S rDNA sequences of 19 clones from the uncontaminated well (T5 ; sequences in boldface type) and 47 clones from the TCE-contaminated well (D3; plain type) along with 16 reference strains. Isolated sequences were distributed across three divisions of the Bacteria: Low G+C Gram-positive bacteria, Proteobacteria, and candidate phylum OP11. Key characteristics of the 16S rDNA tree include: (i) a ¢rst clade containing two green non-sulfur reference strains but no isolated clones, thereby indicating the absence in both sampling locations of sequences related to the dehalorespiring organism Dehalococcoides spp.; (ii) a second clade (bracketed by reference strain Clostridium botulinum and d162) indicating the presence ^ in contaminated groundwater only ^ of three clones (d025, d154, and d011) that resembled (by BLAST) the uncultured clone SJA-19 and its close relative, PCE-dechlorinating bacterium D. restrictus, all clustered with four other Low G+C Gram-positive reference strains and an unidenti¢ed clone (d162); (iii) a third clade composed of 13 clones (bracketed by d021 and d040) that were obtained exclusively from contaminated groundwater ; these had no close relatives, but resembled (by BLASTsimilarity score) endosymbiont sequences reported by Ravenschlag et al. [23]; (iv) a fourth clade containing two K proteobacterial clones (t008 and t009), both originating in the uncontaminated location, and one being related to Agrobacterium sanguineum (t008); (v) a ¢fth clade composed of eight clones obtained exclusively from contaminated groundwater (bracketed by d028 and d064), several of which had high similarity to Zoogloea and Agrobacterium (aerobic K and L Proteobacteria); (vi) a sixth clade containing three unidenti¢ed clones obtained from the uncontaminated location (t022, t041, t051) plus two Q proteobacterial reference strains (E. coli K-12 and Pseudomonas stutzeri); (vii) a seventh clade (bracketed by d085 and Thiobacillus cuprinus) consisting of two reference strains and 16 clones that were obtained from either uncontaminated or contaminated groundwater ^ 12 of these clones originated in well D3 and grouped loosely with the MTBE-degrading reference strain; (viii) a single clone (t043) of uncertain lineage obtained from uncontaminated groundwater ; (ix) a single clone (d076) of uncertain lineage obtained from contaminated groundwater; (x) a deeply branched, heterogeneous clade of 11 sequences (bracketed by t019 and t015) consisting of ¢ve clones from uncontaminated groundwater, three clones from con-

FEMSEC 1354 23-5-02

130

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

taminated groundwater, and three N proteobacterial reference strains (D. vulgaris, G. sulfurreducens and D. chloroethenica); (xi) a lack of sequences related to the PCE/TCEdechlorinating reference strain D. multivorans; (xii) a heterogeneous clade (bracketed by t030 and t046) consisting of four clones of uncertain lineage, three originating in uncontaminated groundwater and one in contaminated well water (d071; related to Bacteroides sp.) ; (xii) another heterogeneous clade (bracketed by d010 and d073) representing the candidate phylum OP11, consisting of ¢ve clones obtained from both uncontaminated and contaminated groundwater; and lastly (xiii) an absence of sequences belonging to the Archaea group represented by the Methanosarcina reference strain. Overall, the 16S rDNA dendrogram reveals the presence of two distinct microbial communities in the two sampling locations. BLAST comparison of sequences also showed no overlap in the form of identical matches. Species diversity indices for the two sampling locations, summarized in the top half of Table 2, indicate reduced species diversity and reduced uniformity in the contaminated location relative to the background. While use of the 16S rDNA gene has become widespread in ecological investigations (e.g. [29,30]), few studies have explored the value of ISR sequence analysis. Therefore, the DNA extracts used for the 16S rDNA analysis were also used to clone ISRs. The resulting dendrogram (Fig. 3) contains 26 clones from T5 and 35 clones from D3 along with 10 reference sequences. Few database entries matched our cloned sequences using BLAST, because of (i) the speci¢city of ISR sequences, (ii) the currently relatively small size of the database, and (iii) the limited availability of ISR sequences for the selected reference strains. Dendrogram associations probably have limited taxonomic signi¢cance although high BLAST scores should provide taxonomic insights. A striking feature of the ISR tree is the clustering of T5 and D3 clones in separate areas of the dendrogram. The top three small clades (bracketed by t1018 and t1019; t1017 and t1031; t1011 and d4008) are composed almost exclusively of T5 clones, with the one exception (d4008) appearing on a separate, remote branch. The ¢rst large clade contains 19 clones from the contaminated well (bracketed by

d0413 and d4048) plus the reference strain D. ethenogenes, that occurs on a separate branch indicating limited similarity. The second large clade (bracketed by d4033 and d4042) contains 14 clones from the contaminated location. Nine of these have 90+% similarity to the MTBE-degrading reference strain. The third large clade (bracketed by t1026 and E. coli K-12) contains nine clones from the uncontaminated site, six of them being similar to the P. stutzeri reference strain. On the bottom of the ISR tree, a ¢nal clade (bracketed by t1003 and reference strain Methanosarcina frisius) combines four clones obtained from uncontaminated groundwater and, on a separate distant branch, the Archaea reference strain. Thus, the strongest clue to the identity of bacteria, as determined by ISR sequence analysis, is the dominance of Pseudomonads in Well T5 (six of 26 clones or 23% total) whereas the identity of frequent clones in Well D3 (bracketed by d4013 and d4048; 19 of 35 clones ; 54% total) is unknown. The separate clustering of T5 and D3 clones in the ISR dendrogram strongly suggests the presence of two distinct microbial communities. Species diversity indices (Table 2, bottom half) support this conclusion, indicating reduced diversity and diminished uniformity in the contaminated site relative to the background.

4. Discussion Geochemical and molecular biological analyses of groundwater from LLNL Site 300 revealed the presence of a microbial community that is tolerant of high levels of toxic chloroethenes (as illustrated by TCE and cis-DCE concentrations of 800 and 250 mg l31 , respectively; both detected on February 9, 1993) and that ferments an unusual co-contaminant linked to in situ reductive dechlorination reactions. The presence of cis-DCE ^ with historical concentrations as high as 390 mg l31 ^ provides irrefutable evidence that intrinsic in situ reductive dechlorination is a major degradative pathway governing the fate of TCE at this national priority site. The occurrence and relevance of further reductive dechlorination of cis-DCE to vinyl chloride and ethene/ethane is of lesser certainty, however. Concentrations of vinyl chloride never exceeded

Table 2 Microbial species diversity indices for pristine groundwater (Well W-834-T5) and TCE-contaminated groundwater (Well W-834-D3), calculated from 16S rDNA and 16S^23S rDNA ISR sequence data Monitoring well 16S rDNA T5 D3 ISR T5 D3

Clone coverage (c)

Shannon^Weaver index (H)

Evenness (e)

18% 61%

3.59 2.40

2.29 1.70

35% 83%

2.68 1.20

2.18 1.54

FEMSEC 1354 23-5-02

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

the detection limit of 5 Wg l31 and levels of ethene and ethane, although detectable in the contaminated area, historically have remained four to six orders of magnitude below those of the primary contaminant TCE (Table 1). Given these concentration di¡erences, we conclude that microbial reductive dechlorination of TCE at the site is largely limited to a single dechlorination step yielding cis-DCE. As discussed below, this conclusion was supported by the results of DNA pro¢ling of contaminated groundwater. 4.1. Sequences potentially linked to bioremediation processes A comprehensive census of the microbial populations in the two study locations was not attempted. Rather, contrasts between the communities, as re£ected by retrieved nucleic acid sequences, were sought. The signi¢cance of DNA sequence-pro¢ling data strongly depends on procedural and analytical approaches taken. Potential sources of bias in molecular biological studies include sample matrix (groundwater vs. sediment), sample collection, DNA extraction protocols, primer selection, PCR, and data analysis tools [25,29,31]. In an attempt to minimize procedural bias, we standardized both the 16S rDNA and the ISR assay as much as possible by processing identical samples, using the same computational procedures, and the same data analysis techniques (BLAST and PileUp). A major advantage of the multiple alignment program PileUp is that, by default, terminal gaps are not penalized; therefore, sequences of very di¡erent lengths (e.g., ISRs and incomplete sequences) can be aligned. This approach di¡ers from the Clustal alignment algorithm adopted in a recently updated version of the ARB software package ([32]; http://ubik.microbiol.washington.edu/ClustalW/ clustalv.html ; Genetics Computer Group, personal communication). Important limitations of PileUp are that it does not provide a rigorous phylogenetic analysis, and that the lack of scale bars complicates direct comparison among multiple dendrograms. However, in this study PileUp was a valuable tool allowing for (i) rapid visualization of microbial diversity, (ii) comparison of sequences that varied in length, and (iii) comparison of two di¡erent pro¢ling techniques. Certain 16S rDNA sequences retrieved from D3 are particularly noteworthy with respect to anaerobiosis, reductive dechlorination, and dehalorespiration. Several 16S rDNA clones showed strong sequence similarity to known representatives of anaerobic bacteria (e.g., G. sulfurreducens, Desulfovibrio). Three out of 61 clones (d025, d154, and d011; 5%) matched closely to GenBank sequence SJA-19, which has been hypothesized to represent yet uncultivated members of the genus Dehalobacter [33]. Providing a de¢nitive mechanistic link between microbial community structure and metabolic functions is very challenging and has been achieved only in a few exceptional

131

studies [34,35]. However, the close relationship of the three SJA-19-related clones to the reference strain D. restrictus is of particular interest, because the occurrence and relative abundance of these sequences may be interpreted as a link between geochemical ¢eld data and the physiology of a known microorganism. D. restrictus is a strict anaerobe that exclusively uses hydrogen as an electron donor when dehalorespiring PCE and TCE to the dead-end product cis-DCE [36,37]. The responsible enzyme is a PCE reductive dehalogenase that recently has been puri¢ed [11]. Thus, predictions based on this type of metabolism match the groundwater chemistry observed in the contaminated sampling location. Three additional sequences (d010, d153, d073) fell into the candidate division OP11, which has been hypothesized to play a signi¢cant role in the bioremediation of an aquifer contaminated with hydrocarbons and chlorinated solvents [24]. However, the pristine well also contained two sequences (t010 and t037) related to OP11. Given the accumulation of cis-DCE in anaerobic site groundwater, it is also interesting to note that the nucleic acid survey failed to detect D. ethenogenes and related sequences. This dehalorespiring, ethene-producing bacterium ([7] and references cited therein) either is not present at the site, or it may occur at levels that are insu⁄cient to a¡ect site chemistry; or it may not be detectable with the non-speci¢c cloning approach taken in this study. In a recent survey that used a more selective nested-PCR approach for detection of D. ethenogenes, one chloroethenecontaminated location and three pristine freshwater sediments tested positive for the target organism [38]. Physiological traits associated with additional 16S rDNA sequences depict a community composition consistent with site biogeochemical reactions. The large clade in Fig. 2 (bracketed by d022 and reference strain PM1), consisting of 11 unidenti¢ed clones loosely a⁄liated with an MTBE-degrading bacterium, showed close similarity to Z93960, a sequence found in activated sludge from a large municipal wastewater treatment plant [39]. The authors noted that the most dominant group in this sludge sample was the L1 group of Proteobacteria. This group encompasses Acidovorax and other genera well known for the utilization of a wide range of carbon sources. Similarly, in the ISR data (Fig. 3), at least nine sequences, obtained from the contaminated well, also were closely related to the MTBE-degrading bacterium that recently was reported to occur at leaking underground storage tank (LUST) sites with MTBE bioattenuation potential [40]. Consistent detection of sequences related to an MTBE-metabolizing bacterium was unexpected because MTBE was not among the contaminants. Extensive extrapolation from 16S rDNA similarity to potential metabolic function of uncultured microorganisms is unwise because signi¢cant disparities may exist between predicted microbial functions and the actual role uncultured microorganisms play in natural environments [30,41].

FEMSEC 1354 23-5-02

132

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

4.2. Interpretation of 16S rDNA and ISR dendrograms One goal of this study’s sequencing e¡orts was to compare the utility of 16S rDNA and ISR sequences in assessing the impact of organic contaminants on the microbial community of the site. Given the moderate number of 16S rDNA and ISR sequences obtained in this study (106 and 61, respectively), it was not surprising to see values for clone coverage ranging from a mere 18% (16S rDNA assay for T5) to 83% (ISR for D3; Table 2). In addition, most of the ISR sequences were inexact matches to GenBank accessions and thus may represent novel species. However, such interpretations must be tentative, due to the small number of ISR sequences in the database. An expanded ISR database containing environmentally relevant sequences is currently being assembled [42], and should make ISR analysis increasingly useful in the future. Sequences obtained in this study may contribute to achieving this goal. In support of the geochemical information compiled in Table 1, the microbial data suggest that distinct selective pressures were present in the pristine and contaminated areas. No overlap was evident between microbial communities in D3 and T5. This is consistent with a similar study in which electrophoresis was used to analyze ISR sequences ampli¢ed from microbial communities native to two adjacent soils from the Amazon region [43]. Despite their close proximity, selective pressure resulting from agricultural use at one of the locations appeared to have triggered the development of two distinct populations [43]. Studies using directly ampli¢ed ISR sequences have also proven to be e¡ective in di¡erentiating closely related pure cultures of Archaea and Bacteria recently isolated from environmental samples [44,45]. In this study, results of 16S rDNA and ISR analyses were in good agreement, as both indicated greater diversity and more evenness in the pristine well relative to the contaminated one. This alteration of the microbial composition in D3 must be attributed to the selective e¡ect of both solvent-induced toxicity and improved availability of carbon/energy sources and electron acceptors. Consistency between 16S rDNA and ISR data underscores the potential usefulness of ISR analysis for purposes of microbial community pro¢ling. In addition, the short length of ISRs provides more e⁄cient PCR ampli¢cation and typically allows for accurate sequencing of most ISRs in two reads. Thus, ISR analysis may o¡er several advantages that can make it a valuable addition to conventional 16S rDNA techniques. 4.3. Signi¢cance of tetrakis(2-ethylbutoxy)silane Hydrolysis of the tetraalkoxysilane co-contaminant can occur either abiotically or biotically, and may result in the release of up to four 2-ethylbutanol moieties per transformed molecule [17]. Laboratory experiments with micro-

cosms constructed from anaerobic Site-300 groundwater showed that fermentation of the released 2-ethylbutanol can result in the formation of 2-ethylbutyrate, acetate and hydrogen [15,18]. From data collected in the present ¢eld study, it is evident that these reactions also occur in situ and that they are of environmental relevance. Concentrations of 2-ethylbutyrate and acetate £uctuated considerably but, overall, had high average values of 94 and 21 mg l31 , respectively, for three sampling events that occurred within a 2-month period (Table 1). Hydrogen concentrations, ¢rst determined in the spring of 2000, were in the low nM range indicating conditions favorable for sulfate reduction and methanogenesis [19]. These observations strongly suggest that the tetraalkoxysilane compound is a key factor determining the fate of TCE at the site; a forthcoming study containing data from seasonal ¢eld samples and anaerobic groundwater microcosms provides additional evidence strengthening this claim [18]. It is remarkable that some 15^45 years after the accidental release of the silicon-based lubricant, fermentation of this compound continues in the subsurface and apparently provides a source of hydrogen for a microbial community whose intrinsic bioremediation activity previously has been observed only in a laboratory setting [15]. In summary, this study provided insights into the microbial activity and bacterial diversity at LLNL Site 300. The documentation of intrinsic in situ bioremediation at the site, achieved by using a combination of geochemical and molecular biological analyses, lends credibility to the proposed application [15,46] of TKEBS and similar tetraalkoxysilanes as long-term slow-release compounds [15] facilitating in situ reductive dechlorination of chloroethenes over periods of years or even decades. An additional bene¢t of these compounds ^ that could not be explored in this work ^ is their potential for triggering aerobic cometabolism of chloroethenes [17]. Tetraalkoxysilanes and their respective hydrolysis products have been shown to serve as the primary substrates of aerobic enrichment cultures capable of co-oxidizing TCE and cis-DCE [17].

Acknowledgements This work was supported by funds from DOE EM-40 under contract W-7405-Eng-48, DOE’s Accelerated Site Technology Deployment Program, Loyola matching funds for students pertaining to DOE NABIR grant DE-FG0299ER62868, DOE grant DE-FG07-96ER62320, and NSF grant CTS-9253633. We would like to thank L. Semprini, S. Vancheeswaran, S. Yu, and M-Y. Chu for quantifying tetraalkoxysilanes and their transformation products. We also would like to thank S. Gregory and V. Madrid for assisting in groundwater sampling and plume contouring, and D. Brown for assisting in DNA extraction and processing. We acknowledge the assistance of L. Petersen, K.

FEMSEC 1354 23-5-02

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

Sorensen and R. Starr in converting the Building 834 Study Area at LLNL Site 300 into an ASTD deployment site for monitored natural attenuation and in situ bioremediation. Finally, we would like to thank the Institute of Genomic Research (TIGR) for sequence data.

[18]

[19]

References [1] National Research Council (2000) Natural Attenuation for Groundwater Remediation. National Academy Press, Washington, DC. [2] Ellis, D.E., Lutz, D.J., Odom, J.M., Buchanan, R.J., Bartlett, C.L., Lee, M.D., Harkness, M. and DeWeerd, K.A. (2000) Bioaugmentation for accelerated in situ anaerobic bioremediation. Environ. Sci. Technol. 34, 2254^2260. [3] Wackett, L.P. and Hershberger, D.C. (2001) Biocatalysis and Biodegradation. ASM Press, Washington, DC. [4] Pieper, D.H. and Reineke, W. (2000) Engineering bacteria for bioremediation. Curr. Opin. Biotechnol. 11, 2626. [5] Morse, J.J., Alleman, B.C., Gossett, J.M., Zinder, S.H., Fennell, D.E., Sewell, G.W. and Vogel, C.M. (1998) A treatability test for evaluating the potential applicability of the reductive anaerobic biological in situ treatment technology (RABITT) to remediate chloroethenes. Draft Technical Protocol. Environmental Security Technology Certi¢cation Program, Department of Defense. http://www.estcp. org/documents/techdocs/Rabbitt_Protocol.pdf. [6] DeBruin, W.P., Kotterman, M.J.J., Posthumus, M.A., Schraa, G. and Zehnder, A.J.B. (1992) Complete biological reductive transformation of tetrachloroethene to ethane. Appl. Environ. Microbiol. 58, 1996^2000. [7] Maymo-Gatell, X., Anguish, T. and Zinder, S.H. (1999) Reductive dechlorination of chlorinated ethenes and 1,2-dichloroethane by Dehalococcoides ethenogenes 195. Appl. Environ. Microbiol. 65, 3108^ 3113. [8] van Eekert, M.H.A. and Schraa, G. (2001) The potential of anaerobic bacteria to degrade chlorinated compounds. Water Sci. Technol. 44, 49^56. [9] El Fantroussi, S., Naveau, H. and Agathos, S.N. (1998) Anaerobic dechlorinating bacteria. Biotechnol. Prog. 14, 167^188. [10] Adrian, L., Szewzyk, U., Wecke, J. and Gorisch, H. (2000) Bacterial dehalorespiration with chlorinated benzenes. Nature 408, 580^583. [11] Holliger, C., Wohlfarth, G. and Diekert, G. (1999) Reductive dechlorination in the energy metabolism of anaerobic bacteria. FEMS Microbiol. Rev. 22, 383^398. [12] Zinder, S.H. (2002) in : Encyclopedia of Environmental Microbiology, Vol. 1:507^516 (Bitton, G., Ed.). John Wiley, New York. [13] Hohnstock-Ashe, A.M., Plummer, S.M., Yager, R.M., Baveye, P. and Madsen, E.L. (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. [14] Boving, T.B. and Brusseau, M.L. (2000) Solubilization and removal of residual trichloroethene from porous media: comparison of several solubilization agents. J. Contam. Hydrol. 42, 51^67. [15] Semprini, L., Vancheeswaran, S., Yu, S.H., Chu, M.Y. and Halden, R.U. (2000) Tetraalkoxysilanes as slow release substrates to promote aerobic and anaerobic dehalogenation reactions in the subsurface. Abstr. Pap. Am. Chem. Soc. 220, 125-ENVR Part 1. [16] Vancheeswaran, S. (1998) Abiotic and biological transformation of TBOS and TKEBS, and their role in the biological transformation of TCE and c-DCE. Master’s Thesis. Department of Civil Engineering, Construction, and Environmental Engineering, Oregon State University, Corvallis, OR. [17] Vancheeswaran, S., Halden, R.U., Williamson, K.J., Jr., J.D.I. and Semprini, L. (1999) Abiotic and biological transformation of tetra-

[20]

[21]

[22]

[23]

[24]

[25]

[26] [27]

[28]

[29]

[30] [31]

[32]

[33]

[34]

[35] [36]

133

alkoxysilanes and tetrachloroethene/cis-dichloroethene cometabolism driven by tetrabutoxysilane-degrading microorganisms. Environ. Sci. Technol. 33, 1077^1085. Vancheeswaran, S., Yu, S., Daley, P., Halden, R.U., Williamson, K.J., Ingle, J.D., Jr. and Semprini, L. (2002) Anaerobic biotransformation of trichloroethene driven by tetraalkoxysilanes at Site 300, Lawrence Livermore National Laboratory, CA, submitted for publication. Chapelle, F.H., Vroblesky, D.A., Woodward, J.C. and Lovley, D.R. (1997) Practical considerations for measuring hydrogen concentrations in groundwater. Environ. Sci. Technol. 31, 2873^2877. Lawrence Livermore National Library. (2000) LLNL Livermore Site and Site 300 Environmental Restoration Project Standard Operating Procedures (SOPs). Lawrence Livermore National Laboratory, Livermore, CA. Sambrook, J., Fritsch, E.F. and Maniatis, T. (1989) Molecular Cloning: a Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Barns, S.M., Fundyga, R.E., Je¡ries, M.W. and Pace, N.R. (1994) Remarkable archaeal diversity detected in a Yellowstone National Park hot spring environment. Proc. Natl. Acad. Sci. USA 91, 1609^1613. Ravenschlag, K., Sahm, K., Pernthaler, J. and Amann, R. (1999) High bacterial diversity in permanently cold marine sediments. Appl. Environ. Microbiol. 65, 3982^3989. Dojka, M.A., Hugenholtz, P., Haack, S.K. and Pace, N.R. (1998) Microbial diversity in a hydrocarbon- and chlorinated-solvent-contaminated aquifer undergoing intrinsic bioremediation. Appl. Environ. Microbiol. 64, 3869^3877. Giovannoni, S.J., Mullins, T.D. and Field, K.G. (1995) in: Molecular Ecology of Aquatic Microbes (Joint, I., Ed.), pp. 217^248. SpringerVerlag, Berlin. Atlas, R. and Bartha, R. (1993) Microbial Ecology. Benjamin/Cummings Publishing Company, Redwood City, CA. Marilley, L., Vogt, G., Blanc, M. and Aragno, M. (1998) Bacterial diversity in the bulk soil and rhizosphere fractions of Lolium perenne and Trifolium repens as revealed by PCR restriction analysis of 16S rDNA. Plant Soil 198, 219^224. Nubel, U., Garcia-Pichel, F., Kuhl, M. and Muyzer, G. (1999) Quantifying microbial diversity: morphotypes, 16S rRNA genes, and carotenoids of oxygenic phototrophs in microbial mats. Appl. Environ. Microbiol. 65, 422^430. Amann, R., Ludwig, W. and Schleifer, K.-H. (1995) Phylogenetic identi¢cation and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59, 143^169. Pace, N.R. (1997) A molecular view of microbial diversity and the biosphere. Science 276, 734^740. Suzuki, M.T. and Giovannoni, S.J. (1996) Bias caused by template annealing in the ampli¢cation of mixtures of 16S rRNA genes by PCR. Appl. Environ. Microbiol. 62, 625^630. Thompson, J.D., Plewnlak, F. and Poch, O. (1999) A comprehensive comparison of multiple sequence alignment programs. Nucleic Acids Res. 27, 2682^2690. Wintzingerode, F.v., Selent, B., Hegemann, W. and Gobel, U.B. (1999) Phylogenetic analysis of an anaerobic, trichlorobenzene-transforming microbial consortium. Appl. Environ. Microbiol. 65, 283^ 286. Orphan, V.J., House, C.H., Hinrichs, K.U., McKeegan, K.D. and DeLong, E.F. (2001) Methane-consuming archaea revealed by directly coupled isotopic and phylogenetic analysis. Science 293, 484^ 487. Boetius, A. et al. (2000) A marine microbial consortium apparently mediating anaerobic oxidation of methane. Nature 407, 623^626. Holliger, C., Schraa, G., Stams, A.J.M. and Zehnder, A.J.B. (1993) A highly puri¢ed enrichment culture couples the reductive dechlorination of tetrachloroethene to growth. Appl. Environ. Microbiol. 59, 2991^2997.

FEMSEC 1354 23-5-02

134

M. Lowe et al. / FEMS Microbiology Ecology 40 (2002) 123^134

[37] Holliger, C. et al. (1998) Dehalobacter restrictus gen. Nov. and sp. Nov., a strictly anaerobic bacterium that reductively dechlorinates tetra- and trichloroethene in an anaerobic respiration. Arch. Microbiol. 169, 313^321. [38] Lo«¥er, F.E., Sun, Q., Li, J. and Tiedje, J.M. (2000) 16S rRNA genebased detection of tetrachloroethene-dechlorinating Desulfuromonas and Dehalococcoides species. Appl. Environ. Microbiol. 66, 1369^ 1374. [39] Snaidr, J., Amann, R., Huber, I., Ludwig, W. and Schleifer, K.-H. (1997) Phylogenetic analysis and in situ identi¢cation of bacteria in activated sludge. Appl. Environ. Microbiol. 63, 2884^2896. [40] Kane, S.R., Beller, H.R., Legler, T.C., Koester, C.J., Pinkart, H.C., Halden, R.U. and Happel, A.M. (2001) Aerobic biodegradation of methyl tert-butyl ether by aquifer bacteria from leaking underground storage tank sites. Appl. Environ. Microbiol. 67, 5824^5829. [41] Achenbach, L.A. and Coates, J.D. (2000) Disparity between bacterial phylogeny and physiology. ASM News 66, 714^715. [42] Garcia-Martinez, J., Bescos, I., Rodriguez-Sala, J. and Rodriguez-

[43]

[44]

[45]

[46]

Valera, F. (2001) RISSC: a novel database for ribosomal 16S-23S RNA gene spacer regions. Nucleic Acids Res. 29, 178^180. Borneman, J. and Triplett, E.W. (1997) Molecular microbial diversity in soils from Eastern Amazonia: evidence for unusual microorganisms and microbial population shifts associated with deforestation. Appl. Environ. Microbiol. 63, 2647^2653. Baudart, J., Lemarchand, K., Brisabois, A. and Lebaron, P. (2000) Diversity of Salmonella strains isolated from the aquatic environment as determined by serotyping and ampli¢cation of the ribosomal DNA spacer regions. Appl. Environ. Microbiol. 66, 1544^1552. DiRuggiero, J., Tuttle, J.H. and Robb, F.T. (1995) Rapid di¡erentiation of hyperthermophilic Archaea by restriction mapping of the intergenic spacer regions of the ribosomal-RNA operons. Mol. Mar. Biol. Biotechnol. 4, 123^127. Yang, Y. and McCarty, P.L. (2000) Biomass, oleate, and other possible substrates for chloroethene reductive dechlorination. Bioremed. J. 4, 125^133.

FEMSEC 1354 23-5-02