Bacterial community analysis of shallow groundwater undergoing
sequential anaerobic and aerobic chloroethene biotransformation
Todd R. Miller1, Mark P. Franklin1 & Rolf U. Halden1,2
1
Department of Environmental Health Sciences, Center for Water and Health, Bloomberg School of Public Health, Johns Hopkins University, Baltimore,
USA; and 2Department of Geography and Environmental Engineering, Whiting School of Engineering, Johns Hopkins University, Baltimore, USA
Correspondence: Rolf U. Halden, 615 N.
Wolfe Street, Rm E6618, Baltimore, MD
21205, USA. Tel.: 1410 955 2609; fax: 1410
955 9334; e-mail: [email protected]
Received 21 July 2006; revised 7 December
2006; accepted 20 December 2006.
First published online 26 March 2007.
DOI:10.1111/j.1574-6941.2007.00290.x
Editor: Riks Laanbroek
Keywords
trichloroethylene; dichloroethene;
tetraalkoxysilane; diesel.
Abstract
At Department of Energy Site 300, beneficial hydrocarbon cocontaminants and
favorable subsurface conditions facilitate sequential reductive dechlorination of
trichloroethene (TCE) and rapid oxidation of the resultant cis-dichloroethene (cisDCE) upon periodic oxygen influx. We assessed the geochemistry and microbial
community of groundwater from across the site. Removal of cis-DCE was shown to
coincide with oxygen influx in hydrocarbon-containing groundwater near the
source area. Principal component analysis of contaminants and inorganic compounds showed that monitoring wells could be differentiated based upon
concentrations of TCE, cis-DCE, and nitrate. Structurally similar communities
were detected in groundwater from wells containing cis-DCE, high TCE, and low
nitrate levels. Bacteria identified by sequencing 16S rRNA genes belonged to seven
phylogenetic groups, including Alpha-, Beta-, Gamma- and Deltaproteobacteria,
Nitrospira, Firmicutes and Cytophaga–Flexibacter–Bacteroidetes (CFB). Whereas
members of the Burkholderiales and CFB group were abundant in all wells (104–109
16S rRNA gene copies L1), quantitative PCR showed that Alphaproteobacteria
were elevated (4 106 L1) only in wells containing hydrocarbon cocontaminants.
The study shows that bacterial community structure is related to groundwater
geochemistry and that Alphaproteobacteria are enriched in locales where cis-DCE
removal occurs.
Introduction
Chloroethenes are ubiquitous pollutants found at many
priority (Superfund) cleanup sites owned and operated by
private, industrial, and Federal entities, including the US
Department of Energy (DOE). Among the various chloroethenes, trichloroethene (TCE) is of particular importance
due to this toxicant’s extensive usage as a degreasing agent
and as a heat exchange fluid either in pure form or in a
mixture with silicon-based lubricants of tetraalkoxysilane
structure (Vancheeswaran et al., 1999). Chloroethenes are
known or suspected carcinogens and their concentrations in
drinking water are regulated at the federal level (Chu &
Milman, 1981; Moore & Harrington-Brock, 2000; Rhomberg, 2000). Thus, removal of these contaminants from
groundwater is of great importance for public health.
In situ bioremediation is one strategy for the removal of
chloroethenes from groundwater. It frequently relies on the
degradative activity of microorganisms native to the site.
Under anaerobic conditions, dehalorespiring microorganFEMS Microbiol Ecol 60 (2007) 299–311
isms are known to utilize chlorinated ethenes as terminal
electron acceptors by replacing chlorines with hydrogen,
using naturally occurring, intentionally added, or cocontaminating compounds as electron donors (Vogel & McCarty,
1985; de Bruin et al., 1992; Holliger et al., 1993; Holliger &
Schumacher, 1994). In this process, tetrachloroethene (PCE)
is sequentially dechlorinated to TCE, cis-DCE, vinyl chloride
and finally to the nonhazardous compounds, ethene and
ethane which in turn can be oxidized to carbon dioxide
under aerobic conditions (de Bruin et al., 1992; Magnuson
et al., 1998). Incomplete reduction of PCE and TCE is
common, resulting in the accumulation of less chlorinated,
though toxic derivatives, including cis-DCE and vinyl chloride. This common scenario is usually caused by a lack of
suitable electron donors and/or the absence of microbial
species able to carry out complete dechlorination (Harkness
et al., 1999; Lendvay et al., 2003). While addition of alternate
electron donors and bioaugmentation with laboratory-cultivated PCE-to-ethene respiring populations has proven
successful in some instances, such an approach may be
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300
complicated by other factors including the periodic influx of
oxygen which can preclude growth of obligate anaerobes,
and the presence of indigenous dehalorespirers that may
outcompete bioaugmented species for reducing equivalents
(Becker, 2006). For this reason it is advantageous to fully
characterize geochemical parameters and microbial species
on a site-by-site basis.
At DOE Site 300 near Livermore, CA, spillage of TCE
together with tetrakis(2-ethylbutoxy)silane (TKEBS) has
created favorable subsurface conditions previously shown
to support the incomplete reductive dechlorination of TCE
to cis-DCE under anaerobic conditions by organisms closely
related to Dehalobacter restrictus (Lowe et al., 2002). This is
consistent with reports indicating that this bacterium converts PCE and TCE to cis-DCE during anaerobic growth in
the presence of molecular hydrogen, although it does not
further dechlorinate the resultant cis-DCE (Holliger et al.,
1993). Potential electron donors for dehalogenases are
dihydrogen and fumarate, both common end-products
produced by fermentative mixed microbial communities.
At Site 300, fermentation of 2-ethylbutanol and n-butanol,
released continuously during slow hydrolysis of silicon
lubricants, has been shown to support reductive dechlorination of TCE to cis-DCE (Vancheeswaran et al., 2003). The
accumulation of cis-DCE under anaerobic conditions is
problematic; however, it may biodegrade aerobically under
favorable conditions (Bradley & Chapelle, 1998; Azizian
et al., 2005).
In the presence of oxygen, cometabolism of chloroethenes
by nonspecific oxygenases may occur. A variety of species
have been shown to produce mono- and dioxygenases that
have relaxed substrate specificities, supporting the oxidation
of primary growth substrates and of a number of chlorinated ethenes serving as cosubstrates (Folsom et al., 1990;
Mcclay et al., 1995; Shim et al., 2001). For example,
methanotrophic species of the genera Methylocystis, Methylosinus, Methylomonas and Methylococcus possess soluble
methane monooxygenases that rapidly transform aliphatic
and aromatic substrates, including chlorinated ethenes to
less toxic compounds in the presence of suitable electron
donor, carbon and nitrogen sources (Koh et al., 1993; Grosse
et al., 1999). Methylococcus capsulatus also aerobically degrades methyl bromide (Oremland et al., 1994). Other TCEand DCE-cometabolizing species, including Burkholderia,
Ralstonia, Pseudomonas, Mycobacterium and Xanthomonas,
can utilize ammonia, toluene, ethylene, propylene, or other
hydrocarbons as growth substrates [reviewed in (Arp et al.,
2001)].
In this study, we observed removal of anaerobically
accumulated cis-DCE from shallow perched groundwater at
DOE Site 300 when oxygen was introduced due to soil vapor
extraction activites and/or rainfall events. To begin to
understand what members of the microbial community
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T.R. Miller et al.
might be involved in this removal process, we assessed the
geochemistry and microbial community composition in
groundwater from subsurface strata where cis-DCE removal
occurs in the presence of cocontaminating hydrocarbons.
Whereas previous microbial ecological studies of the site
employed 16S rRNA gene cloning and focused on two
monitoring wells in pristine and highly contaminated locations (monitoring wells T5 and D3, respectively) (Lowe
et al., 2002), the present study utilized denaturing gradient
gel electrophoresis (DGGE) and quantitative PCR (qPCR)
to explore the microbial community within a larger geochemical gradient intersected by a total of five sampling
locations containing multiple contaminants.
Materials and methods
Study site description and sample collection
Site 300 is an explosives test facility located in the Altamont
Hills of CA, 100 km southeast of San Francisco. It is
operated by the Department of Energy and listed as an EPA
Superfund site. The area under study is an Operable Unit
known as the Building 834 complex (B834) where TCE and
TKEBS were spilled from faulty piping systems over the
course of several decades. This chemical mixture is contaminating a shallow perched water-bearing zone in which
chloroethenes are migrating in a southwesterly direction.
Monitoring wells provide access to groundwater outside and
inside of the chloroethene plume, representing pristine to
highly contaminated conditions with varying amounts of
TCE, TKEBS and cis-DCE. In addition, diesel is present in
groundwater from well U1, situated in a location previously
occupied by a fuel tank.
For chemical analyses, water was pumped to the surface
using an in situ pump and stored in glass bottles certified for
volatile organic carbon analysis (I-Chem, Rockwood, TN).
Microorganisms were sampled by passing 4 L of groundwater through sterile 0.2 mm Mini capsule filters (Pall
Corporation, MI). Both water samples and filters collected
in May of 2002 were immediately placed on ice for transport
back to the laboratory, where they were stored at 20 1C.
DNA extraction
Genomic DNA was extracted directly from filters as previously described by Giovannoni et al. (1996), with minor
modifications. Each unit received 2 mL of lysis buffer
(20 mM EDTA, 400 mM NaCl, 0.75 sucrose and 50 mM Tris
HCl pH 9.0) and were incubated at 37 1C for 15 min.
Proteinase K (0.2 g mL1) and 1% SDS was added and the
units incubated at 55 1C for 60 min. The lysate was recovered
and extracted twice with equal volumes of phenol–chloroform–iso amyl alcohol (25 : 24 : 1; pH 8) followed by a
single extraction with an equal volume of chloroform. The
FEMS Microbiol Ecol 60 (2007) 299–311
301
Microbial ecology of chloroethene biotransformation
aqueous phase was removed and the DNA precipitated using
isopropanol and sodium acetate, washed with 70% ethanol
and resuspended in 300 mL of sterile distilled water.
PCR and DGGE
The 16S rRNA gene was amplified in a PCR using eubacterial primers 907R and GM5F-GC-clamp (Teske et al., 1996).
These primers amplify the variable V3 region of the 16S
rRNA gene producing 550 bp fragments suitable for phylogentic comparisons (Muyzer et al., 1995). Each 50 mL reaction mixture contained 1.5 mM MgCl2, 50 mM KCl, 10 mM
Tris HCl (pH 9.0), 2% bovine serum albumin (BSA),
100 pmol each dNTP, 50 pmol of each primer, 1 U of Redtaq
DNA polymerase (Sigma, Milwaukee, WI) and a total of
1 ng of extracted DNA. After a ‘hot start’ at 94 1C for 5 min
the reaction mix underwent a touchdown PCR whereby the
initial annealing temperature was set at 65 1C and decreased
by 1 1C every two cycles until a touchdown of 55 1C, at
which temperature 10 additional cycles were carried out.
Amplification was performed at 1 min of denaturation at
94 1C, 1 min of primer annealing, and 1 min of primer
extension at 72 1C, followed by 7 min of final primer
extension. We also attempted to amplify 16S rRNA genes
from known dehalorespiring bacteria using primer sets and
PCR conditions previously reported (Löffler et al., 2000;
Fagervold et al., 2005). PCR products were analyzed by
DGGE using a BioRad Dcode system (BioRad, Hercules,
CA). Replicate PCR products were loaded onto multiple 6%
(w/v) acrylamide gels [37.5 : 1 acrylamide: N,N 0 -methylenebis-acrylamide in 20 mM Tris buffer (pH 7.8), 10 mM
sodium acetate, and 0.5 mM disodium EDTA] containing
linear chemical gradients with a 5% change in denaturant
concentration ranging from 35–65% denaturant, where
100% denaturant is equal to 7 M urea and 40% deionized
formamide. The gels were run at 100 V for 18 h at 60 1C. Gels
were stained with 50 mg mL1 ethidium bromide and visualized using a UV transilluminator and gel documentation
system (Alpha Imager 2000 V5.5, Alpha Innotech Corporation, VA). All visible bands were excised from the gels using
sterile pipette tips and eluted from the gel matrix in 100 mL
of sterile distilled water overnight at 4 1C. A 1-mL aliquot of
the suspension was used for PCR amplification using the
same primers as described above and PCR products were
reanalyzed by DGGE to confirm the presence of a single
band. PCR products were purified with the QIAquick-spin
DNA purification system (Qiagen, Valencia, CA) as per
manufacturer’s instructions and subjected to cycle sequencing using an Applied Biosystems 3730 DNA Analyzer.
Analysis of DGGE gels
DGGE gel band patterns were subjected to computer
analysis to compare bacterial community structure. Band
FEMS Microbiol Ecol 60 (2007) 299–311
migration distances were measured from a common point at
the top of the gel to the middle of each band using Adobe
Photoshop (San Jose, CA). To facilitate band identification a
straight horizontal line was drawn across the middle of each
band, and three measurements were made per band. These
data were then used to construct a dendrogram according to
the unweighted pair group method with arithmatic mean
(UPGMA) using the SIMINT and SAHN modules of the
NTSYSPC Analysis software package (EXETER software, Setauket, NY).
Comparative sequence analysis
All sequences obtained from DGGE bands were analyzed for
chimeras in a single batch file using the Bellerophon server
(Huber et al., 2004). Putative chimeric sequences were then
reanalyzed individually using Pintail (Ashelford et al., 2005),
comparing percent base pair mismatches of the putative
chimeric sequence to its closest BLAST match using a window
size of 25 bp. Sequences were judged to be chimeric based
upon the deviation from an expectation statistic calculated
by the software. Nonchimeric sequences were then used in a
BLAST search (Altschul et al., 1990) to find the top five most
similar sequences in GenBank and all sequences were
manually aligned according to the 16S rRNA gene secondary
structure using Phydit (Chun, 1995) and E. coli 16S rRNA
gene secondary structure as the template. Additional sequences from reference strains obtained from GenBank were
added to delineate representative taxons. Evolutionary trees
were generated using the neighbor-joining, Fitch-Margoliash, and maximum parsimony algorithms in the PHYLIP
package (Felsenstein, 1989). Evolutionary distance matrices
for the neighbor-joining and Fitch-Margoliash methods
were generated as described by Jukes & Cantor (1969). The
confidence in tree topology was evaluated after 1000 bootstrap samplings of the neighbor-joining data, and only
values of 500 or higher were shown on the tree.
Chemical analyses
Concentrations of chloroethenes, TKEBS and inorganic
compounds in groundwater samples were determined by
GC/MS using standard methods as previously described
(Lowe et al., 2002). Contour maps showing contaminant
plumes were constructed from a database of regulatory
compliance monitoring data as previously described (Lowe
et al., 2002).
Principal component analysis
A principal component analysis of organic and inorganic
chemical concentrations was performed using NTSYSPC software. TKEBS and diesel contaminants were not included
due to infrequent sampling for these compounds. These
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302
T.R. Miller et al.
data were standardized using the STAND module and the
transformed data used to compute a correlation matrix
using the SIMINT module. Following calculation of Eigen
vectors, both observations and vectors were projected onto
the principal component axes using the MXPLOT module.
Real-time quantitative PCR (qPCR)
Group-specific primers corresponding to major bacterial
groups defined by a phylogenetic analysis were constructed
and used in qPCR to estimate group-level abundance in
groundwater. All sequences obtained from DGGE bands
were analyzed using PRIMROSE computer software (Ashelford
et al., 2002) to find unique oligonucleotides within sequences belonging to each phylogenetic group. Pairs of
oligonucleotides with similar melting temperatures
(55–65 1C) were chosen that could serve as group-specific
forward and reverse primers and that amplified a 75–150 bp
portion of the 16S rRNA gene. The OLIGOCHECK program
(Ashelford et al., 2002) was then used to determine group
specificity of each primer pair relative to 16S rRNA
gene sequences identified at the site and those within the
Ribosomal Database Project (release 8.1) combined (Cole
et al., 2003).
Real-time quantitative PCR (qPCR) was performed using
a Smart Cycler System (Cepheid, Sunnyvale, CA) as described elsewhere (Monis et al., 2005). Each 25 mL reaction
contained 1 qPCR buffer [50 mM KCl, 10 mM Tris HCl
(pH 8.0), 1.5 mM MgCl2, and 2 mM Syto 9 (Molecular
Probes, Eugene, OR)], 0.4 mM each deoxynucleotide triphosphate, 5 pmol each primer, 1 U Taq DNA polymerase (New
England Biolabs, Ipswich, MA) and 1–20 ng of template
DNA. The PCR conditions were an initial denaturation at
95 1C for 2 min followed by 45 cycles of denaturation at
95 1C for 30 s, annealing for 30 s and elongation at 72 1C for
30 s. Annealing temperatures were 55, 57, and 61 1C for
group I, II and V primer pairs, respectively (Table 1).
Fluorescence of Syto 9 was measured after each elongation
step and the cycle at which fluorescent intensities increased
above 30 amu was defined as the cycle threshold (Ct) value.
Background fluorescence, measured in cycles 1–5 was subtracted from all measurements. The Ct values produced
from a dilution series of known 16S rRNA gene standards
(described below) was used to estimate group level abundance in the unknown samples.
The 16S rRNA gene fragments from known bacteria were
excised from DGGE bands, reamplified and used as standards for qPCR reactions. Resulting PCR products were
purified using a Qiagen PCR cleanup kit according to the
manufacturer’s instructions and a portion analyzed by
DGGE to confirm a single band. The concentration of the
purified standard was determined by adding a 5 mL aliquot
to a 1 : 1000 dilution of Sybr Gold DNA stain in distilled
water and fluorescent intensity measured at excitation and
emission wavelengths of 490 and 530 nm, respectively, using
a SpectraMax Gemini EM fluorometer (Molecular Devices,
Sunnyvale, CA). Fluorescent intensity was compared to that
of a dilution series of bacteriophage lambda DNA (New
England Biolabs, Ipswich, MA), to determine the mass per
volume (ng mL1) of the purified 16S rRNA gene standards.
The 16S rRNA gene copy number of each standard was
calculated using the following formula: Copies 16S rRNA
gene = (Mass of standard ng mL1)/(length of standard
(bp) (1.09 1012) (Feris et al., 2003).
Nucleotide sequence accession numbers
The 550 bp partial 16S rRNA gene sequences from Site 300
groundwater bacteria were deposited in GenBank under
accession numbers DQ336570–DQ336606.
Results
Distribution of contaminants and inorganic
compounds
A contour map showing the distribution of TCE, cis-DCE
and TKEBS throughout the study site is shown in Fig. 1.
Well D3 is located in a source area where c. 3000 kg of TCE
Table 1. Melting temperature (Tm) and specificity of qPCR primers targeting three different phylogenetic groups in Site 300 groundwater
Target product
size (bp)
Group most closely
affiliated with
Group I (77)
CFB group
Group II (76)
Alphaproteobacteria
Group V (86)
Burkholderiales
Matches to target/nontarget
sequences in RDP (%)w
Primer name/sequence
Tm
CFB/forward (5 0 –3 0 ) YTYCGTGCCAGSAGCCGS
CFB/reverse (3 0 –5 0 ) TTTAAAGGGTSCGYAGGYGG
Alpha-forward (5 0 –3 0 ) ATAAGCYCCGGCTAACTCCG
Alpha-reverse (3 0 –5 0 ) GTTCGGAATTACTGGGCGTA
Burk-forward (5 0 –3 0 ) AAGCACCGGCTAACTACGTGCCA
Burk-reverse (3 0 –5 0 ) ACTGGGCGTAAAGCGTGCGCAG
62.8
59.3
60.4
57.3
64.2
65.8
91/7.4
86/2.1
93/8.4
16S rRNA gene sequences recovered from groundwater were aligned with each other and unique oligonucleotides within groups of phylogenetically
related sequences were identified as primer pairs for qPCR.
w
Indicates the percent of all target or nontarget sequences in RDP matched by the primer pairs allowing for no more than two mismatches.
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FEMS Microbiol Ecol 60 (2007) 299–311
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Microbial ecology of chloroethene biotransformation
Fig. 1. Contour map of the study location, DDE Site 300, showing the
extent of groundwater contamination with trichloroethene (TCE), cisdichloroethene (cis-DCE) and tetrakis(2-ethylbutoxy)silane (TKEBS), a
silicon-based lubricant. Five groundwater monitoring wells located inside
and outside of the contaminated area were sampled for bacterial
community analysis. Well D3 is located in an area where TCE and TKEBS
were spilled, whereas the diesel-contaminated well U1 is situated in a
location formerly occupied by a leaking fuel tank.
were spilled together with similarly substantial but unknown quantities of silicon oil (TKEBS). Longterm monitoring data was used to calculate the average yearly chemical
concentrations (n = 5–32). Groundwater in this location is
characterized by moderately high concentrations of TCE
(yearly average and SD of 40 25 mg L1) and relatively
high concentrations of the bacterial dechlorination product
cis-DCE (23 32 mg L1) (Fig. 2a). This is also the only well
containing insoluble quantities of the silicon oil TKEBS
(173 274 mg L1). Well U1 is closest to well D3 and
contains higher concentrations of TCE (123 28 mg L1)
but low concentrations of cis-DCE (2.0 1.8 mg L1). This
well is in an area where unknown amounts of diesel fuel
were spilled over the years. Marking the northern edge of the
TCE contour is well B3, which contains moderately high
concentrations of cis-DCE (8.6 4.9 mg L1), and comparatively little TCE (0.35 0.38 mg L1). At the southern end of
FEMS Microbiol Ecol 60 (2007) 299–311
Fig. 2. Groundwater chemistry at the study location represented as
annual average concentrations of volatile organic compounds (a) and
select anions and cations (b). Error bars indicate the SD of multiple
measurements from samples taken throughout the year (n = 5–32).
the TCE contour is well T2, located about 300 m downgradient of the source area. It contains elevated concentrations of TCE (21 6.2 mg L1), no detectable cis-DCE
(o 0.005 mg L1), and none of the hydrocarbon cocontaminants known to be associated with both anaerobic and
aerobic transformation of chloroethenes (Vancheeswaran
et al., 1999). Another well, T5, located further down
gradient, is considered a control well since no contaminants
are present.
Select electron acceptors and inorganic nutrients for
bacterial growth were also measured in site groundwater
(Fig. 2b). Nitrate (NO
3 ) levels varied considerably across the
sampling locations. Wells T2 (1.16 mM) and T5 (1.61 mM)
had average nitrate concentrations that were 1–2 orders of
magnitude higher than those found in wells B3, D3, and U1
(0.18, 0.17, and 0.006 mM, respectively) (Fig. 2b). Sulfate
levels were highest in wells B3 and T2, followed by wells
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U1, T5 and D3. Ammonia (NH1
4 ) was low in all wells
(0.003–0.02 mM) and nitrite (NO
2 ) did not vary considerably except for well U1, wherein concentrations were lower
(0.06 mM) than measured elsewhere (0.31–0.51 mM). Dissolved oxygen concentrations (data not shown) in wells U1
and D3 are low, and variable (o 0.4–5.1 mg L1; average and
SD of 1.6 0.42 and 1.7 1.59, respectively) while oxygen
levels in wells B3 and T2 are slightly higher (2.2 2.2 and
2.6 1.9 mg L1, respectively) and well T5 is oxygenated
year round (6.5 1.0 mg L1).
Yearly fluctuations in chloroethenes and
dissolved oxygen
At the source area near well D3, removal of the bacterial
dechlorination product cis-DCE occurs simultaneously with
oxygen influx. This is exemplified in Fig. 3 which shows
typical yearly trends in oxygen and chloroethene concentrations and is representative of a larger database of regulatory
compliance monitoring data. During August and September, dissolved oxygen in Well D3 groundwater dropped to
1 mg L1 while levels of cis-DCE increased from 8.3 to
42.7 mg L1. In October, a spike in oxygen levels occurred
and cis-DCE concentrations dropped to 1.7 mg L1. In
November, the well returned to a low-oxygen state and cisDCE levels increased from 1.2 to 16.4 mg L1. Continuous
dissolution of dense nonaqueous phase liquid containing
chloroethenes, silicon oil and other contaminants led to
Fig. 3. Yearly fluctuations in cis-DCE and TCE relative to dissolved
oxygen (DO) in groundwater at well D3 near the source area. cis-DCE
maxima occur during periods of low DO concentrations (o 1 mg L1) and
cis-DCE removal occurs simultaneously with oxygen influx. TCE remains
abundant except when DO falls below 1 mg L1 for 2 months or longer.
The concentrations presented are for one representative year selected
from a body of multi-year, long-term regulatory compliance monitoring
data.
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T.R. Miller et al.
concentrations remaining above 30 mg L1, except when
oxygen levels dropped below 1 mg L1 for 2 months or
longer, thereby promoting the reductive dehalogenation
process known to occur at the site (Lowe et al., 2002).
Principal component analysis
To determine overall geochemical relationships among the
wells, concentrations of TCE, cis-DCE, nitrate, nitrite,
ammonia and sulfate were subjected to a principal component (PC) analysis. Approximately, 50%, 26% and 15% of
the variability in the dataset could be explained by PC1,
PC2, and PC3, respectively. Figure 4 shows the first two axes
of the PC analysis along with the vectors (monitoring wells)
and observations (chemical measurements). The amount of
scattering of observations away from the origin indicates
chemical variability among all wells while the angle between
two vectors (dotted lines) indicates the level of correlation
between two wells. The first PC (PC1) clearly differentiates
wells on a horizontal plane, based upon TCE concentration,
whereas the second PC (PC2) separates the wells by cis-DCE
along the vertical axis. The third PC (not shown) separates
wells by nitrate levels along a z-axis. Accordingly, well B3
contains cis-DCE and no TCE, which places it in an almost
Fig. 4. Principal component (PC) analysis of chemical concentrations,
detected in groundwater from five monitoring locations at the site, and
resultant Eigen vectors (dotted lines). Observations of the same chemical
are circled. PC1, which accounted for 50.23% of the observed variations
in all measurements combined, differentiated wells based upon TCE
values, whereas PC2, accounting for 25.97% of the observed variability
differentiated wells based upon DCE values. A third PC, (not shown)
differentiated wells based upon NO
3 levels. Less sensitive parameters
1
included concentrations of NO
2 and NH4 .
FEMS Microbiol Ecol 60 (2007) 299–311
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Microbial ecology of chloroethene biotransformation
Fig. 5. This representative DGGE gel shows the microbial community structure in groundwater from five monitoring wells, as determined by separation
of fragments of 16S rRNA genes amplified from extracted total DNA (a). Following image annotation (b), the migration distances of individual bands
were interpreted by UPGMA cluster analysis to yield a dendrogram (c) indicating considerable species diversity at the site, with similar structure observed
in wells T5 and T2, as well as in D3 and U1. Several bands are shared among at least two wells (closed arrow) while one band is shared among four wells
(open arrow).
completely vertical orientation away from well T5, the
pristine control location. Wells T2 and U1 are separated by
a small angle. Both contain a small amount of cis-DCE and
comparatively significant levels of TCE, which places them
on a horizontal plane. Well D3 contains both DCE and TCE,
causing a trajectory falling between well B3 and the T2/U1
grouping. Sulfate, nitrite and ammonia are not as important
for variations in geochemistry among the wells, as evidenced
by the fact that all of these observations are grouped near
the origin.
Bacterial community profiles
We examined if variations in groundwater chemistry reflected differences in bacterial community structure using
DGGE. This technique separates heterogeneous samples of
16S rRNA gene fragments based upon their melting characteristics, and it represents a rapid tool for fingerprinting
analysis and preliminary identification of relatively abundant community members (Watts et al., 2001). RepresentaFEMS Microbiol Ecol 60 (2007) 299–311
tive gel patterns are shown in Fig. 5a. Bacterial 16S rRNA
gene fragments amplified from the study site produced 6–11
bands per well in the denaturing gel. The bacterial community from wells U1 and D3 produced the largest number of
bands (11 and 10, respectively) followed by well B3 (8
bands) then wells T2 and T5, which produced the least
number of bands (7 and 6, respectively). Bands were marked
with a horizontal line (Fig. 5b) and band migration distances measured, defining bands with equal distances as
operational taxonomic units (OTU). None of the OTUs
were shared among all wells. However, a total of seven were
shared among at least two wells (closed arrow) and one OTU
was shared among four wells (open arrow). The OTUs were
compared by cluster analysis in a dendrogram constructed
using the unweighted pair group method with arithmetic
mean (UPGMA) (Fig. 5c). Results indicated similar bacterial communities in wells T2 and T5, as well as in wells U1
and D3. Interestingly, well B3 was placed in an isolated
location, suggesting its bacterial community to be distinct
from all others.
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Identification of bacterial community members
To better understand these differences in bacterial species
composition, 16S rRNA gene fragments in DGGE bands
were sequenced and analyzed. A total of 37 nonchimeric,
nonredundant sequences were recovered and aligned with
GenBank entries (NCBI, Bethesda, MD). In neighbor-joining trees the bacteria clustered into seven well-supported
phylogenetic groups including Cytophaga–Flexibacter–Bacteroidetes (CFB), Alpha-, Beta-, Gamma- and Deltaproteobacteria, Nitrospira and Firmicutes (Fig. 6). Well D3, near the
source area, contained 16S rRNA gene sequences related to
that of Acidovorax sp. UFZ-B517 that grows aerobically on
chlorobenzene (clone number, expect value, percent sequence identity; 115-32, E = 0, 98%). This well also contained multiple sequences related to hydrogen oxidizing
bacteria within the genus Hydrogenophaga (112-23, E = 0,
99% and 115-29, E = 0, 99%) and sequences related to the
nitrogen fixing, root symbiont, Cupriavidus taiwanesis (11531, E = 0, 99%) (Wen et al., 1999; Alfreider et al., 2002;
Verma et al., 2004). Well U1, which is near well D3, also
contained 16S rRNA gene sequences similar to hydrogen
oxidizing bacteria within the genus Hydrogenophaga (11541, E = 0, 99%) (Wen et al., 1999). In addition, this well
contained sequences related to an aerobic benzene degrading bacterium (112-26, E = 0, 95%), an estradiol degrading
Novosphingomonas tardaugens ARI-1 (112-43, E = 0, 98%),
methylotrophic species (115-38, E = 0, 96%), a denitrifying
pivalic acid degrading bacterium (112-28, E = 0, 95%), a
thiosulfate oxidizing bacterium (115-39, E = 0, 98%) and
bacteria previously found in a uranium contaminated
aquifer (115-40, E = 0, 98%) (Rooney-Varga et al., 1999;
Fujii et al., 2002; Gihring et al., 2002; Graff & Stubner, 2003;
Probian et al., 2003; Lueders et al., 2004). Well B3 contained
sequences similar to a manganese oxidizing bacterium,
Leptothrix discophora strain SP-6 (115-18, E = 1e-133,
93%), a denitrifying pivalic acid degrading bacterium (11539, E = 0, 97%), hydrogen oxidizing bacteria (112-53, E = 0,
99%) and a type II methane oxidizing bacterium (115-14,
E = 0, 98%) (Siering & Ghiorse, 1996; Wen et al., 1999;
Heyer et al., 2002; Probian et al., 2003). Both wells T2 and
T5 contained many sequences related to uncultured and/or
uncharacterized bacteria or cloned sequences; however, well
T2 contained 16S rRNA gene sequences similar to that of the
filamentous bacterium Haliscomenobacter hydrosis (110-14,
E = 0, 98%), normally associated with wastewater treatment
systems, as well as 16S rRNA gene sequences related to a
hydrogen oxidizing bacterium (115-72, E = 0, 99%) and a
sulfate reducing bacterium (115-76, E = 0, 99%) (Wen et al.,
1999; Bade et al., 2000; Schauer & Hahn, 2005).
When grouped by monitoring well, bacterial community
structure varied by distance to the source area (well D3). For
example, in the proximate wells, D3, U1 and B3, the
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T.R. Miller et al.
majority of species identified (67%, 64%, and 80% of species
diversity, respectively) were related to the Burkholderiales,
whereas in well T2 most species were related to Firmicutes
(30%) or the CFB group (30%).
Abundance of bacterial groups
To estimate bacterial abundance at the group level, qPCR
primers were designed to amplify 16S rRNA genes from
bacterial clusters identified by the phylogenetic analysis.
Primer pairs for qPCR designed specifically for groups I, II
and V allowed for two mismatches between primer and
template (Table 1). All sequences within each group identified at Site 300 could be amplified by the appropriate primer
pair. When aligned with the Ribosomal Database Project,
91%, 86% and 93% of sequences amplified by the group I,
II, and V primer pairs were from the CFB group, Alphaproteobacteria and Burkholderiales, respectively. Within group
II, the majority of targets were Sphingomonas species (99%).
In qPCR reactions using 103 to 4 107 copies of known 16S
rRNA gene fragments as template, there was a reciprocal,
linear relationship between target template concentration
and cycle threshold (Ct) value. This linear relationship
occurred in both the presence and absence (Fig. 7a–c, open
and closed symbols, respectively) of 4 107 nontarget fragments, thereby affirming confidence in the group-level
specificity of the newly designed primer pairs.
Using these qPCR primers, bacterial abundances in
groundwater from different wells were assessed at the group
level. Overall, total copy numbers of all groups combined
were in the order B34 U14 T2 (8.94 8.24 2.8 108 copies L1), followed by well D3 (3.3 107 copies L1)
and then T5 (3.7 106 copies L1). Group II bacteria were
most abundant in wells near the source area, starting with
well U1 followed by wells D3 and B3; the more remote wells
T2 and T5 had the lowest counts (Fig. 7b). Bacteria of group
V were most abundant in well B3, followed by well T2,
then U1, D3 and T5. Bacteria belonging to group I were
most abundant in well T2 and U1, followed by well T5, B3
and D3. These data indicate that of the three groups
quantified, group II bacteria showed an increased abundance in wells containing higher levels of chloroethenes and
cocontaminants.
Discussion
The present study characterized geochemical parameters
within a contaminated aquifer and identified abundant
bacterial groups associated with cis-DCE removal. The data
indicate that (1) nitrate, TCE and DCE are principal drivers
of variability in groundwater geochemistry across the site,
(2) the bacterial community includes seven major phylogenetic groups, and some of these may be related to known
FEMS Microbiol Ecol 60 (2007) 299–311
307
Microbial ecology of chloroethene biotransformation
Fig. 6. Neighbor-joining tree showing the phylogenetic relationship of select GenBank entries to the 16S rRNA gene sequences (330 bp) obtained
from site groundwater (accession numbers indicated in parentheses). Bootstrap values (n = 1000) are reported for neighbor-joining analysis values
of 4 500. An ‘f’ or ‘p’ indicates agreement of the Fitch and parsimony analysis results with those of the neighbor-joining tree (bar represents 0.1 U of
evolutionary distance). Names of sequences obtained from Site 300 groundwater in this study are enclosed in boxes.
FEMS Microbiol Ecol 60 (2007) 299–311
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308
T.R. Miller et al.
Fig. 7. Three newly designed qPCR primer pairs exhibited a linear relationship between cycle-threshold values and total copies of target DNA in the
absence (, solid line) and presence (’, dotted line) of 107 copies of nontarget 16S rRNA gene fragments (a). Application of the primers demonstrated
an abundance of 16S rRNA genes from group-II bacteria in highly contaminated wells relative to other species quantified (b). Error bars represent the SD
of three independent measurements (see Materials and methods for additional detail).
chloroethene cooxidizing bacteria, and (3) group II bacteria
most similar to Alphaproteobacteria are abundant in
groundwater at monitoring wells featuring chloroethenes
and multiple hydrocarbon contaminants known to support
chloroethene degradation.
The qPCR data show that group-II bacteria are more
abundant in contaminated groundwater near the source
area. The combined concentration of cis-DCE and TCE is
highest in wells closest to the source area (wells D3, U1 and
B3) (Fig. 2a) and both wells D3 and U1 contain cocontaminants including TKEBS and diesel fuel, respectively. One
possible explanation for this finding is that group-II bacteria
are able to derive energy and biomass from the degradation
of cocontaminating hydrocarbons. The group-II primers
amplify 16S rRNA genes from Alphaproteobacteria, mainly
Sphingomonas species. Many Sphingomonas species are capable of degrading aromatic and aliphatic hydrocarbons, such
as those present in well U1 (diesel) and well D3 (TKEBS
breakdown products). While Sphingomonas spp. are generally not associated with TCE degradation they have been
linked to the cometabolism of TCE in mixed cultures (Lee
et al., 2002; Den et al., 2003). Other Alphaproteobacteria in
group II may also participate in TCE cometabolism, including
type II methanotrophs Methylocystis species (11514-B3) (Nakajima et al., 1992; McDonald et al., 1997; Grosse et al., 1999;
Heyer et al., 2002). Some Methylocystis species produce a
soluble methane monooxygenase capable of oxidizing aliphatic and aromatic substrates including TCE and other chlorinated ethenes (Grosse et al., 1999; Shigematsu et al., 1999).
The obvious question to answer is: which species cause
the rapid cis-DCE loss observed at Site 300 in response to
oxygen infiltration in wells near the source area (Fig. 3)? The
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phylogenetic analysis of rRNA gene sequences accomplished
in this study does not conclusively identify bacterial phenotypes. However, the physiology of phylogenetically related
organisms combined with group-specific qPCR data provides possible clues. The relative abundance of Alphaproteobacteria in source-area wells and the knowledge that some
of these organisms are phylogenetically related to known
hydrocarbon degraders (Sphingomonas species) and chloroethene-cometabolizing bacteria (Methylocystis species)
suggests these bacteria may be involved in the final step of
complete chloroethene degradation at the site. Experiments
targeting functional genes, such as soluble methane monooxygenase, butane monooxygenase and/or other mono and
dioxygenases whose primary substrate is associated with
diesel fuel or silicon oil may provide further evidence.
Accordingly, the hypothesized multi-step process of complete chloroethene degradation is initiated by the established
mechanism of 2-ethylbutanol and n-butanol release from
the hydrolysis of TKEBS (Vancheeswaran et al., 2003).
Excess amounts of these two carbon and energy sources
promote oxygen depletion and anaerobic conditions, as
shown previously (Vancheeswaran et al., 2003). Fermentation of the alcohols then can provide an ample supply of
dissolved molecular hydrogen, as demonstrated in microcosms prepared from site materials (Vancheeswaran et al.,
2003). Dehalorespiring organisms may use this energy
source to perform the reductive dechlorination of TCE to
DCE (Lowe et al., 2002; Vancheeswaran et al., 2003). DCE is
not dechlorinated further since organisms potentially capable of performing this task (Maymo-Gatell et al., 1997; He
et al., 2005) are absent from this site. This was demonstrated
previously (Lowe et al., 2002) and again in the present study,
FEMS Microbiol Ecol 60 (2007) 299–311
309
Microbial ecology of chloroethene biotransformation
in which primer pairs suitable for selective amplification of
Dehalococcoides related organisms (detection limit4 50 copies per reaction) (Fagervold et al., 2005) did not produce
an amplicon, whereas controls did (data not shown). The
documented periodic influx of oxygen into site groundwater
(Fig. 3) may help to explain the absence of Dehalococcoides
related organisms. Therefore, DCE likely is a dead-end
product of microbial metabolism at the site as long as
anaerobic conditions prevail. However, upon temporal
reaeration of the shallow groundwater, the compound can
be removed (Fig. 3). The present study identified group-II
bacteria as candidate organisms performing this final and
important function of DCE oxidation and succeeded in
providing an initial overview of species diversity and spatial
microbial distribution at the site. Information gained from
this study will guide future experiments focused on the
identification of individual species responsible for chloroethene cooxidation at Site 300.
Acknowledgements
This work was performed under the auspices of the US
Department of Energy by Lawrence Livermore National
Laboratory under Contract W-7405-Eng-48 and with funds
from DOE’s Accelerated Site Technology Deployment Program. The authors acknowledge Victor Madrid and Steven
Gregory for their help with groundwater monitoring, data
interpretation and mapping, as well as site selection and
sampling activities. We would also like to thank Rey de
Castro for his assistance in the interpretation of the principal component analysis results and Kristen Gibson for
assistance with quantitative PCR.
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