• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of aemPermissionsJournals.ASM.orgJournalAEM ArticleJournal InfoAuthorsReviewers
Appl Environ Microbiol. Nov 2007; 73(21): 6898–6904.
Published online Sep 7, 2007. doi:  10.1128/AEM.01218-07
PMCID: PMC2074934

Detection and Quantification of Geobacter lovleyi Strain SZ: Implications for Bioremediation at Tetrachloroethene- and Uranium-Impacted Sites[down-pointing small open triangle]

Abstract

Geobacter lovleyi strain SZ reduces hexavalent uranium, U(VI), to U(IV) and is the first member of the metal-reducing Geobacter group capable of using tetrachloroethene (PCE) as a growth-supporting electron acceptor. Direct and nested PCR with specific 16S rRNA gene-targeted primer pairs distinguished strain SZ from other known chlorinated ethene-dechlorinating bacteria and closely related Geobacter isolates, including its closest cultured relative, G. thiogenes. Detection limits for direct and nested PCR were approximately 1 × 106 and 1 × 104 16S rRNA gene copies per μl of template DNA, respectively. A quantitative real-time PCR (qPCR) approach increased the sensitivity to as few as 30 16S rRNA gene copies per μl of template DNA but was less specific. Melting curve analysis and comparison of the shapes of amplification plots identified false-positive signals and distinguished strain SZ from G. thiogenes when analyzed separately. These indicators were less reliable when target (strain SZ) DNA and nontarget (G. thiogenes) DNA with high sequence similarity were mixed, indicating that the development of qPCR protocols should not only evaluate specificity but also explore the effects of nontarget DNA on the accuracy of quantification. Application of specific tools detected strain SZ-like amplicons in PCE-dechlorinating consortia, including the bioaugmentation consortium KB-1, and two chlorinated ethene-impacted groundwater samples. Strain SZ-like amplicons were also detected in 13 of 22 groundwater samples following biostimulation at the uranium- and chlorinated solvent-contaminated Integrated Field-Scale Subsurface Research Challenge (IFC) site in Oak Ridge, TN. The numbers of strain SZ-like cells increased from below detection to 2.3 × 107 ± 0.1 × 107 per liter groundwater, suggesting that strain SZ-like organisms contribute to contaminant transformation. The G. lovleyi strain SZ-specific tools will be useful for monitoring bioremediation efforts at uranium- and/or chlorinated solvent-impacted sites such as the Oak Ridge IFC site.

Chlorinated ethenes such as tetrachloroethene (PCE) and trichloroethene (TCE) are common groundwater pollutants due to their extensive use for industrial and military applications (17). Hexavalent uranium, U(VI), has been released into the environment as a consequence of weapons-grade uranium production and milling (20, 52). Uranium processing typically requires chlorinated solvents, such as PCE and TCE; therefore, U(VI) and chlorinated ethene contamination often coexist (e.g., at the Integrated Field-Scale Subsurface Research Challenge [IFC] site [formerly the Field Research Center] in Oak Ridge, TN [http://www.esd.ornl.gov/nabirfrc]) (48, 50). Due to negative human health impacts, both chlorinated ethenes and uranium are regulated by the U.S. Environmental Protection Agency (EPA). Considerable effort has been devoted to elucidate the fate of these contaminants and to identify microbial populations involved in their transformation and detoxification (19, 43, 49). Although a number of phylogenetically diverse PCE- or U(VI)-reducing bacteria have been obtained from various environments (21, 22, 43, 49), none of the isolates were reported to mediate both activities. Only the recently described isolate Geobacter lovleyi strain SZ combines the ability to reductively dechlorinate PCE and reduce soluble U(VI) to sparingly soluble U(IV) (40, 45). Interestingly, strain SZ reduces PCE and U(VI) concomitantly (45), making this organism a promising candidate for bioremediation at mixed-waste sites such as the Oak Ridge IFC site. Geobacter spp. are commonly found in freshwater sediments and subsurface environments, but the distribution of members of the recently proposed dechlorinating Geobacter clade (45) is unclear. The goals of this study were to design specific and quantitative 16S rRNA gene-based approaches for assessing the environmental distribution and abundance of strain SZ-like Geobacter spp. in uranium- and chlorinated ethene-contaminated sites and to explore the value of the new tools for bioremediation-monitoring efforts.

MATERIALS AND METHODS

Sources of DNA.

DNA was isolated from pure cultures, PCE-dechlorinating enrichment cultures, bioaugmentation consortia, and groundwater and aquifer materials from contaminated sites. The pure cultures included G. lovleyi strain SZ (DSM 17278) (45), G. thiogenes (formerly Trichlorobacter thiogenes) (ATCC BAA-34, kindly provided by B. Griffin, Michigan State University) (8, 31), G. sulfurreducens (DSM 12127, kindly provided by K. Nevin, University of Massachusetts) (4), G. metallireducens (DSM 7210, kindly provided by K. Nevin) (26), Desulfuromonas michiganensis strain BB1 (DSM 15941) (47), Anaeromyxobacter dehalogenans strain 2CP-C (ATCC BAA-259) (39), Sulfurospirillum (formerly Dehalospirillum) multivorans (DSM 12446) (28, 41), Dehalococcoides sp. strain FL2 (14), Dehalococcoides sp. strain BAV1 (13), and Desulfitobacterium sp. strain Viet1 (23, 25). The Geobacter isolates were grown as described by Sung et al. (45). Strain BB1 and strain 2CP-C were grown as described by Sung et al. (47) and Sanford et al. (39), respectively, and all other pure cultures were grown as described by Amos et al. (1). R. Daprato (Georgia Institute of Technology) kindly provided genomic DNA and a plasmid containing a single copy of the 16S rRNA gene of Dehalobacter restrictus (DSM 9455) (15). Genomic DNA was obtained from three PCE-to-ethene dechlorinating enrichment cultures derived from chlorinated ethene-contaminated aquifer materials (the Hydrite site, Cottage Grove, WI; the TRW site, Minerva, OH; and the FMC site, San Jose, CA) (44); one PCE-to-cis-1,2-dichloroethene (cis-DCE) dechlorinating enrichment culture derived from TCE-contaminated aquifer materials collected at the East Gate Disposal Yard in Ft. Lewis, WA (34); and one PCE-to-ethene dechlorinating mixed culture derived from Kalamazoo River (MI) sediment (44). Bio-Dechlor INOCULUM, a PCE-to-ethene dechlorinating bioaugmentation inoculum (37), was grown as described previously (1) to obtain genomic DNA. Genomic DNA of the bioaugmentation inoculum KB-1 (10, 11, 30) was kindly provided by M. Duhamel, University of Toronto. DNA was also obtained from groundwater collected within a TCE plume originating from an industrial site near Milledgeville, GA, and from aquifer material collected at the East Gate Disposal Yard in Ft. Lewis, WA (34). Twenty-two groundwater samples were obtained from the uranium-contaminated Oak Ridge IFC site. Samples were collected from wells FW016 and FW029 of area 1 (16) and well TPB16 of area 2. Area 3 groundwater samples were from the unconditioned and nonstimulated control well FW106 (27 May 2004) (51) and from a successful pilot-scale uranium bioreduction demonstration plot (50, 51). The pilot-scale demonstration used a nested-cell approach comprising a biostimulated inner recirculation loop that periodically received an electron donor (i.e., ethanol) and an outer recirculation loop that minimized interactions between the low-pH and high-nitrate aquifer groundwater and the chemically and biologically conditioned groundwater in the biostimulated, inner loop (50, 51). The four-well recirculation system consisted of an outer loop injection well (FW024), an outer loop extraction well (FW103), an inner loop injection well (FW104), and an inner loop extraction well (FW026). The system also contained three multilevel sampling wells: FW100, FW101, and FW102. Specific multilevel sampling well depths are designated as outlined by Wu et al. (50). Periodic addition of ethanol to the inner recirculation loop began on 7 January 2004 (51). Groundwater samples were obtained on four occasions during the first 6 months of periodic biostimulation with ethanol: 2 February 2004 (wells FW103, FW100-2, FW100-3, FW101-3, and FW104); 5 February 2004 (well FW024); 27 May 2004 (wells FW026 and FW101-2); and 16 June 2004 (well FW102-3). Additional samples were obtained 1 to 1.5 years after the onset of periodic ethanol biostimulation as follows: 25 May 2005 (well FW104); 4 August 2005 (wells FW104 and FW102-2); and 5 August 2005 (wells FW026, FW101-1, FW102-1, FW102-4, FW100-1, and FW100-4). Eleven of the 22 samples came from wells FW029, FW104, FW026, FW101-3, FW101-1, FW101-2, FW102-2, and FW102-3, located in subsurface regions affected by biostimulation (i.e., ethanol addition), while the remaining samples came from wells FW016, TPB16, FW106, FW024, FW103, FW100-2, FW100-3, FW102-1, FW102-4, FW100-1, and FW100-4, located in regions not influenced or only marginally affected by biostimulation (16, 50, 51).

DNA isolation.

Genomic DNA was extracted from 5 to 10 ml of chlorinated ethene-dechlorinating pure and mixed cultures, as well as from cultures of G. thiogenes, G. metallireducens, and G. sulfurreducens, by use of a QIAamp DNA mini kit (QIAGEN, Valencia, CA) with modifications as described previously (1, 36). For quantitative real-time PCR (qPCR) standards, a 16S rRNA gene of strain SZ was cloned into plasmid Topo-TA pCR2.1 (Invitrogen, Carlsbad, CA) as described previously (18); plasmids containing strain SZ's 16S rRNA gene (pSZ16S) were extracted from Escherichia coli host cells by use of a QIAprep spin miniprep kit (QIAGEN) (36). Community genomic DNA was isolated from 1.8 to 1,715 liters of Oak Ridge IFC site groundwater and collected in final volumes of 10 to 450 μl as described previously (53); for qPCR, the DNA was diluted 1:10 or 1:20 before analysis. Genomic DNA was obtained from 1 liter of Milledgeville site groundwater and from 40 ml of a TCE-fed Ft. Lewis enrichment culture by use of a MoBio ultra clean water DNA kit (Jefferson City, MO) (36). DNA from 10 g of Ft. Lewis soil was extracted using a PowerMax soil DNA isolation kit (MoBio). Purified DNA was stored at −20°C until analysis.

PCR primers and conditions.

Specific PCR primer pairs were designed using Oligo design and analysis tools (Integrated DNA Technologies, Coralville, IA; http://www.idtdna.com/SciTools/SciTools.aspx?cat=DesignAnalyze) based on the nearly complete 16S rRNA gene sequence of strain SZ (GenBank accession number AY914177). The 16S rRNA gene sequences of strain SZ and related species, including G. thiogenes (GenBank accession number AF223382), G. sulfurreducens (GenBank accession number U13928), G. metallireducens (GenBank accession number L07834), and Desulfuromonas michiganensis strain BB1 (GenBank accession number AF357915), were aligned using the MegAlign program of the Lasergene software package (DNA Star, Inc., Madison, WI). The sequences of the selected primers, Geo196F and Geo999R, and mismatches between the primer sequences and the sequences of 16S rRNA gene fragments from closely related organisms are shown in Table Table1.1. The Geobacteraceae-targeted primer Geo564F described by Cummings et al. (5) was modified to produce primer Geo535R (Table (Table1).1). The expected sizes of the Geo196F/Geo999R and Geo196F/Geo535R amplicons are 820 and 357 bp, respectively.

TABLE 1.
Primers used to detect and quantify Geobacter lovleyi strain SZ and alignment of 16S rRNA gene sequence fragments of strain SZ with corresponding regions of closely related species

To experimentally verify primer specificity, purified genomic DNA of close Geobacter relatives and selected reductively dechlorinating species (D. michiganensis strain BB1, Anaeromyxobacter dehalogenans strain 2CP-C, Sulfurospirillum multivorans, Dehalobacter restrictus, Dehalococcoides sp. strain FL2, Dehalococcoides sp. strain BAV1, and Desulfitobacterium sp. strain Viet1) was subjected to PCR with both Geo196F/Geo999R and Geo196F/Geo535R primer pairs. The reaction mixture contained 2 μl of 10× PCR buffer (Applied Biosystems [ABI], Foster City, CA), 2 mM MgCl2, 0.13 mg of bovine serum albumin/ml (Promega, Madison, WI), 200 μM of each deoxynucleoside triphosphate (ABI), 2.5 U of AmpliTaq polymerase (Gibco BRL, Gaithersburg, MD), 100 nM of each primer, and 1 μl of template DNA (10 to 20 ng/μl) in a total reaction volume of 20 μl. Amplification with a gradient thermocycler (Eppendorf Mastercycler gradient 5331; Eppendorf, Hamburg, Germany) determined optimum annealing temperatures of 50 and 53°C for primer pairs Geo196F/Geo535R and Geo196F/Geo999R, respectively. The following temperature program was used for the designed primer pairs: 94°C for 2.2 min, 30 cycles of 94°C for 30 s, 50°C or 53°C for 45 s, and 72°C for 2.2 min, followed by 6 min at 72°C. For nested PCR, an initial amplification was performed with bacterial primer pair 8F and 1514R followed by a second round of PCR with the strain SZ-specific primer pairs and using 1:50 dilutions of the amplicons obtained in the first round of PCR as template DNA (24). PCR products were visualized in 1% agarose gels in Tris-acetate-EDTA buffer stained in an aqueous ethidium bromide solution (1 μg/ml). Select amplicons obtained with the Geo196F/Geo999R primer pair were purified (QIAquick PCR purification kit; QIAGEN) and partially sequenced using an ABI 3100 genetic analyzer (ABI).

The detection limits (i.e., sensitivities) of direct and nested PCR with the Geo196F/Geo999R primer pair were determined by performing PCR on a 10-fold dilution series of G. lovleyi strain SZ genomic DNA. The gene targets in the highest dilution were quantified via qPCR analysis, and the DNA concentrations in subsequent dilutions were calculated based on this estimate. The reported detection limits are the lowest dilutions that yielded a visible band in ethidium bromide-stained gels.

qPCR analysis.

qPCR analysis to quantify G. lovleyi strain SZ was performed using SYBR green-based detection chemistry and the Geo196F/Geo535R primer pair. The reaction mixture contained 15 μl of Power SYBR green PCR master mix (ABI), 300 nM of each primer, and 3 μl of template DNA in a total reaction volume of 30 μl. The PCR temperature program was as follows: 2 min at 50°C, 15 min at 95°C followed by 40 cycles of 30 s at 94°C, 30 s at 50°C, and 30 s at 72°C. qPCR was carried out with an ABI 7500 fast real-time PCR system, and melting curve analysis was performed with the default settings of the ABI software from 67°C to 95°C. Standard curves were generated following a procedure outlined by Ritalahti et al. (36), which used a 10-fold dilution series of quantified plasmid (concentrations determined spectrophotometrically at 260 nm) carrying a single copy of strain SZ's 16S rRNA gene (pSZ16S). To identify false positives, template DNA was replaced with sterile water (i.e., no-template controls). The genome of strain SZ contains two 16S rRNA gene copies (DOE Joint Genome Institute); therefore, dividing gene copy numbers by a factor of 2 yielded the cell numbers. The specificity of the Geo196F/Geo535R primer pair in the qPCR approach was evaluated with triplicate samples of purified genomic DNA of G. lovleyi strain SZ (12.5 ng/μl), G. thiogenes (16.1 ng/μl, 1.61 ng/μl, and 0.16 ng/μl), G. sulfurreducens (1.9 ng/μl), and G. metallireducens (6.9 ng/μl). To determine the influence of DNA from closely related species on the quantification of strain SZ, qPCR analysis was performed on DNA mixtures (see the supplemental material) with increasing amounts of G. thiogenes DNA. To determine if sample dilution affected strain SZ quantification, qPCR was performed on undiluted and 1:10-diluted strain SZ DNA.

RESULTS AND DISCUSSION

Specific, sensitive, and quantitative detection of strain SZ.

With strain SZ DNA as the template, the designed primer pairs Geo196F/Geo999R and Geo196F/Geo535R yielded amplicons of the expected sizes (820 bp and 357 bp, respectively) and sequences. In direct and nested PCR, both primer pairs distinguished strain SZ from related species, including G. thiogenes, G. sulfurreducens, G. metallireducens, and D. michiganensis strain BB1, and the other reductively dechlorinating bacteria tested. Direct PCR with the Geo196F/Geo999R primer pair required >106 16S rRNA gene copies per μl of template DNA to produce a visible band in ethidium bromide-stained agarose gels, whereas the nested-PCR approach was 2 orders of magnitude more sensitive. Although the primer pair Geo196F/Geo999R did not achieve the sensitivity reported for other 16S rRNA gene-targeted primers (e.g., see reference 24), the strain SZ-specific PCR approach proved valuable in evaluating the distribution of strain SZ-like organisms in environmental samples (see below). The qPCR approach was significantly more sensitive and had a quantification limit of ~30 16S rRNA gene copies per μl of template DNA. The qPCR approach produced linear standard curves ranging from 8.4 × 101 to 8.4 × 108 16S rRNA gene copies per PCR (R2 = 0.9991; amplification efficiency of 1.80 [35]). The qPCR quantification limit reported herein is consistent with the quantification limits reported for other qPCR target genes (e.g., Dehalococcoides 16S rRNA and reductive dehalogenase genes [36]) and slightly lower than the quantification limit observed with another Geobacter-targeted qPCR protocol (10). The latter method used primer pair Geo73f/Geo485r to target a Geobacter organism whose 16S rRNA gene sequence (GenBank accession number AY780563) shares 99% identity with that of G. lovleyi strain SZ (10). This primer pair does not contain the discriminating mismatches presented in Table Table11 and hence would not distinguish between G. lovleyi and G. thiogenes.

In qPCR analysis, the shapes of the amplification (i.e., fluorescent signal intensity) plots (not shown) and the melting curves (Fig. (Fig.1)1) were similar for triplicate samples of genomic DNA from strain SZ compared to the plasmid (pSZ16S) standards. Melting (i.e., disassociation) temperatures of PCR amplicons depend mainly on amplicon size and GC content and can be used as an additional diagnostic tool to verify target sequence amplification (2, 3, 38). As shown in Fig. Fig.1,1, the average melting temperatures (Tm) for the amplicons produced with genomic and plasmid (pSZ16S) DNA of strain SZ were 83.6 ± 0.0°C and 83.6 ± 0.4°C, respectively. These data agree well with the estimated Tm of 85°C calculated with an oligonucleotide properties calculator (version 3.13; http://www.basic.northwestern.edu/biotools/oligocalc.html) for the 357-bp amplicon. Minimal amplification (i.e., below the quantification limit) was observed with triplicate samples of G. sulfurreducens and G. metallireducens genomic DNA, and no amplification was observed without template DNA. Some amplification (i.e., fluorescence) was observed with G. thiogenes, although the shape of the amplification plot was markedly different than the shape of the plots for strain SZ. In fact, the software available with the ABI 7500 fast real-time PCR system failed to quantify samples of G. thiogenes due to a noisy baseline. The melting curves for samples of G. thiogenes, with an average Tm of 70.5 ± 0.0°C, were markedly different than the curves for strain SZ (Fig. (Fig.1),1), and the experimental Tm (70.5 ± 0.0°C) did not agree with the Tm of 84°C predicted for G. thiogenes. Similar melting curves and temperatures were obtained when 1:10 and 1:100 dilutions of the genomic DNA of G. thiogenes were used as templates in qPCR. When qPCR products were analyzed in agarose gels, amplicons of the expected size (357 bp) were observed for strain SZ, but no amplicons were produced with G. thiogenes template DNA. Fluorescence was not observed with strain SZ or G. thiogenes DNA when either the forward (Geo195F) or the reverse (Geo535R) primer was omitted from the PCR. Although some amplification of G. thiogenes DNA occurred with the described qPCR approach, multiple lines of evidence (e.g., melting curves, amplification plots, and noisy baselines) distinguished G. lovleyi strain SZ from G. thiogenes during qPCR analysis. Similar discrimination of closely related organisms by melting curve analysis has previously been described for distinguishing and identifying medically relevant microorganisms (summarized by Robinson et al. [38]). Hence, melting curve analysis may also be a useful tool to differentiate environmentally relevant bacteria with highly similar target sequences, potentially allowing for high-resolution discrimination (3).

FIG. 1.
Melting curve analysis following qPCR for samples of G. lovleyi strain SZ (genomic DNA or plasmid DNA with a 16S rRNA gene insert of strain SZ [pSZ16S]), samples of G. thiogenes (genomic DNA), and no-template controls. The melting curves represent averages ...

Although the qPCR approach clearly identified false-positive signals, G. thiogenes DNA influenced the quantification of strain SZ when DNA from both isolates was present in the same sample. In the absence of G. thiogenes DNA, the average estimate for the number of strain SZ cells per ml of a PCE-grown culture was 1.50 × 108 ± 0.11 × 108 (see the supplemental material). The estimated cell numbers increased linearly (R2 = 0.971) in the presence of increasing concentrations of G. thiogenes DNA by up to 1.78-fold (see the supplemental material). A 10-fold dilution of pure strain SZ template DNA did not have a significant effect on the cell number estimate, indicating that the observed effect of G. thiogenes DNA was not due to dilution of strain SZ DNA. The increased estimate for strain SZ in the presence of G. thiogenes is likely related to the nonspecific fluorescent signal produced from the G. thiogenes DNA. The melting curves following qPCR with mixtures of strain SZ and G. thiogenes DNA did not contain the characteristic peak for G. thiogenes (~70.5°C) (Fig. (Fig.1)1) but only the characteristic peak for strain SZ (average Tm of 83.9 ± 0.3°C). These results suggest that qPCR overestimates the abundance of strain SZ in the presence of G. thiogenes DNA and that melting curve analysis cannot identify when the overestimation occurs (i.e., when G. thiogenes and strain SZ DNA occur concomitantly in a sample). Hence, the development of qPCR protocols and other quantitative, nucleic acid-based tools should not only evaluate the specificity of the approach but also explore the effects of nontarget DNA, including DNA with high sequence similarity to the target DNA sequence, on the accuracy of quantification (27).

Application of the strain SZ-specific detection tools.

Direct PCR with the Geo196F/Geo999R primer pair detected strain SZ-like organisms in the PCE-to-ethene dechlorinating Hydrite enrichment culture (not shown). With acetate provided as the electron donor, the Hydrite culture accumulated cis-DCE, whereas complete reductive dechlorination to ethene occurred in cultures amended with hydrogen (44). This dechlorination pattern is consistent with dechlorinator physiology: strain SZ couples either acetate or hydrogen oxidation to the reduction of PCE and TCE to cis-DCE (45), and Dehalococcoides sp. strain GT, which was isolated from the Hydrite culture (46), requires hydrogen as the electron donor to drive the process to completion (i.e., ethene formation). No amplification with the Geo196F/Geo999R primer pair was observed with DNA samples from three other PCE-to-ethene dechlorinating mixed cultures in either direct or nested PCR (not shown). Direct PCR produced strain SZ-like amplicons with DNA from the KB-1 bioaugmentation consortium, but no amplicons were obtained with DNA from Bio-Dechlor INOCULUM, even with nested PCR. Recent studies have demonstrated growth of the SZ-like organism in the KB-1 consortium during PCE and TCE dechlorination to cis-DCE (9-11). Strain SZ-like organisms were detected in both groundwater samples from a TCE-impacted aquifer near Milledgeville, GA. Dechlorination of TCE to stoichiometric amounts of cis-DCE was observed in microcosms and occurred in situ after biostimulation (42), suggesting that strain SZ-like organisms were stimulated and played a role in the observed dechlorination. No SZ-specific signals were obtained with DNA extracted from TCE-contaminated Ft. Lewis soil. However, PCR amplification with the Geo196F/Geo999R primer pair occurred with DNA obtained from a TCE- and lactate-amended enrichment culture derived from the Ft. Lewis soil, and qPCR estimated 4.34 × 106 ± 0.96 × 106 strain SZ-like cells per ml of culture. Additionally, three 16S rRNA gene sequences with 96 to 99% similarity to that of strain SZ were detected in a clone library constructed with DNA extracted from the Ft. Lewis enrichment culture (34).

Detection of strain SZ-like organisms in chlorinated ethene-dechlorinating enrichment cultures, in the KB-1 bioaugmentation consortium, and in environmental samples from chlorinated ethene-impacted aquifers, coupled with the fact that previous studies detected SZ-like 16S rRNA gene sequences in PCE-to-ethene dechlorinating mixed cultures and contaminated aquifers, suggests that dechlorinating Geobacter spp. may be relevant contributors to chlorinated ethene detoxification (6, 7, 9-11, 29). Apparently, chlorinated ethene-dechlorinating Geobacter species are not rare in the environment, thus expanding our understanding of the physiological versatility of the Geobacteraceae. Additional dechlorinating Geobacter isolates are needed to refine the parameters delineating the putative dechlorinating clade within the Geobacteraceae (45).

Detection of strain SZ at the Oak Ridge IFC site.

Strain SZ-like organisms were detected in community genomic DNA extracted from 13 of 22 Oak Ridge IFC site groundwater samples. Specific amplicons were obtained with direct PCR (not shown) and nested PCR (Fig. (Fig.2A)2A) from samples of wells FW026, FW101-2, and FW102-3, located within the area 3 biostimulated zone (50, 51). Nested PCR yielded an additional positive signal with DNA collected from well FW029 (area 1), which was also influenced by biostimulation (Fig. (Fig.2A)2A) (16). Sequence analysis of at least 350 bp of all amplicons confirmed sequence identity with the 16S rRNA gene sequence of strain SZ. No amplification occurred (Fig. (Fig.2A)2A) with samples from wells FW016 (area 1), TPB16 (area 2), and FW106 (area 3), which are located in areas that received no biostimulation treatment.

FIG. 2.
(A) Detection of strain SZ-like organisms via nested PCR in chlorinated ethene- and uranium-contaminated Oak Ridge IFC site groundwater taken from regions impacted by biostimulation (wells FW026 [lane 3], FW029 [lane 4], FW101-2 [lane 5], and FW102-3 ...

As depicted in Fig. Fig.2B,2B, strain SZ-like organisms could also be enumerated in various groundwater samples collected within the uranium bioreduction plot at the Oak Ridge IFC site (50, 51). In February 2004, approximately 1 month after the initial amendment of ethanol to the inner recirculation loop, the number of strain SZ cells was less than 5 × 104 per liter groundwater (Fig. (Fig.2B).2B). Minimal amplification (i.e., fluorescence) was observed in most of the samples from February 2004 (Fig. (Fig.2B),2B), and cell number estimates for these samples were extrapolated from the linear qPCR standard curve. These estimates must be interpreted cautiously since the fluorescent signals were below the limit for accurate quantification (~30 16S rRNA gene copies per μl of template DNA; see above). The melting curves for these samples did not have the characteristic peak for strain SZ, but instead several smaller peaks were visible (Fig. (Fig.2C).2C). All samples that could be quantified accurately had melting curves with the characteristic peak for strain SZ (Fig. (Fig.2C).2C). These results demonstrate that melting curve analysis was useful to distinguish nonspecific amplification and identify false-positive signals with environmental samples. Samples taken following biostimulation indicated substantial growth of strain SZ-like organisms (Fig. (Fig.2B).2B). For example, cell numbers in samples from the inner recirculation loop injection well (FW104) increased by over 3 orders of magnitude from February 2004 to May and August 2005 (Fig. (Fig.2B).2B). The samples from May and August 2005 were taken 1 to 1.5 years after the onset of periodic ethanol addition to the inner recirculation loop, indicating that strain SZ persisted at high levels in the biostimulated zone throughout the biostimulation period. In August 2005, strain SZ cell numbers ranged from 105 to 106 per liter of groundwater in regions impacted by biostimulation (Fig. (Fig.2B).2B). In contrast, the number of SZ cells was significantly lower in samples taken at the same time from areas not impacted by biostimulation (Fig. (Fig.2B).2B). These observations indicate that strain SZ-like organisms were present at the Oak Ridge IFC site and, importantly, responded to biostimulation. The addition of ethanol and/or adjustment of other geochemical parameters (e.g., pH, nitrate removal [50, 51]) stimulated growth of strain SZ-like organisms. Although strain SZ cannot utilize ethanol as a direct electron donor, ethanol fermentation yields hydrogen and acetate, which are electron donors used by strain SZ (45). There is ample evidence that microbes indigenous to the Oak Ridge IFC site produce acetate when stimulated by ethanol: acetate accumulated (i) in ethanol-amended microcosms containing sediment from well FW104 (33), (ii) in an ethanol-amended soil column containing sediment from FW104 (12), and (iii) in portions of the area 3 biostimulated zone following ethanol addition (51). In a sulfate-reducing and U(VI)-reducing enrichment culture derived from the soil column and dominated by Desulfovibrio-like species, ethanol addition led to the accumulation of acetate in molar excess of the added amount, indicating homoacetogenenic activity (32). Hydrogen was produced intermittently. When the enrichment was transferred without sulfate, Clostridium-like bacteria became dominant while acetate and hydrogen accumulated. These observations indicate that several microbial processes produce acetate (and hydrogen) from ethanol in the Oak Ridge IFC site subsurface and that strain SZ could potentially partner with different types of native bacteria to reduce U(VI) and PCE. Previous work has demonstrated that G. lovleyi strain SZ reduces U(VI) and PCE simultaneously (45), and hence strain SZ is a likely contributor to in situ bioremediation at the mixed-waste Oak Ridge IFC site (48, 50).

The identification of key organisms contributing to contaminant detoxification [e.g., U(VI) reduction and reductive dechlorination] represents a targeted approach for bioremediation monitoring. Although comprehensive information about the microbial community composition and dynamics at bioremediation sites is desirable, the application of targeted monitoring approaches to delineate and quantify the contributions of select organisms may produce tangible results in the short term. Monitoring key contributors, such as strain SZ-like organisms and other U(VI)-reducing bacteria (e.g., Anaeromyxobacter spp. and other metal reducers), to U(VI) bioreduction and reductive dechlorination at the Oak Ridge IFC site and other contaminated sites may provide sufficient information for productive implementation of bioremediation technologies.

Supplementary Material

[Supplemental material]

Acknowledgments

This research was supported by the Environmental Remediation Science Division (ERSD), Biological and Environmental Research (BER), U.S. Department of Energy (grant no. DE-FG02-04ER63718), by the Strategic Environmental Research and Development Program (projects ER-1586 and ER-1419), and in part by the National Science Foundation under grant no. 0090496 (CAREER award to F.E.L.). B.K.A. was supported by an EPA STAR fellowship (no. U916150) and an NSF graduate research fellowship. K.E.F. acknowledges support from an NSF graduate research fellowship and partial support through an NSF IGERT fellowship.

Oak Ridge National Laboratory is managed by University of Tennessee UT-Battelle LLC for the Department of Energy under contract DE-AC05-00OR22725.

We thank B. Lynch for operating the ABI 3100 sequencer and the Seattle District Army Corps of Engineers personnel for providing Ft. Lewis soil samples.

Footnotes

[down-pointing small open triangle]Published ahead of print on 7 September 2007.

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

REFERENCES

1. Amos, B. K., R. C. Daprato, J. B. Hughes, K. D. Pennell, and F. E. Löffler. 2007. Effects of the nonionic surfactant Tween 80 on microbial reductive dechlorination of chlorinated ethenes. Environ. Sci. Technol. 41:1710-1716. [PubMed]
2. Applied Biosystems. 2005. Power SYBR Green PCR master mix and RT-PCR protocol (P/N 4367218 Rev. B). Applied Biosystems, Foster City, CA.
3. Bælum, J., T. Henriksen, H. C. B. Hansen, and C. S. Jacobsen. 2006. Degradation of 4-chloro-2-methylphenoxyacetic acid in top- and subsoil is quantitatively linked to the class III tfdA gene. Appl. Environ. Microbiol. 72:1476-1486. [PMC free article] [PubMed]
4. Caccavo, F., D. J. Lonergan, D. R. Lovley, M. Davis, J. F. Stolz, and M. J. Mcinerney. 1994. Geobacter sulfurreducens sp. nov., a hydrogen- and acetate-oxidizing dissimilatory metal-reducing microorganism. Appl. Environ. Microbiol. 60:3752-3759. [PMC free article] [PubMed]
5. Cummings, D. E., O. L. Snoeyenbos-West, D. T. Newby, A. M. Niggemyer, D. R. Lovley, L. A. Achenbach, and R. F. Rosenzweig. 2003. Diversity of Geobacteraceae species inhabiting metal-polluted freshwater lake sediments ascertained by 16S rDNA analyses. Microb. Ecol. 46:257-269. [PubMed]
6. Daprato, R. C., F. E. Löffler, and J. B. Hughes. 2007. Comparative analysis of three tetrachloroethene to ethene halorespiring consortia suggests functional redundancy. Environ. Sci. Technol. 41:2261-2269. [PubMed]
7. Dennis, P. C., B. E. Sleep, R. R. Fulthorpe, and S. N. Liss. 2003. Phylogenetic analysis of bacterial populations in an anaerobic microbial consortium capable of degrading saturation concentrations of tetrachloroethylene. Can. J. Microbiol. 49:15-27. [PubMed]
8. De Wever, H., J. R. Cole, M. R. Fettig, D. A. Hogan, and J. R. Tiedje. 2000. Reductive dehalogenation of trichloroacetic acid by Trichlorobacter thiogenes gen. nov., sp. nov. Appl. Environ. Microbiol. 66:2297-2301. [PMC free article] [PubMed]
9. Duhamel, M., and E. A. Edwards. 2007. Growth and yields of dechlorinators, acetogens, and methanogens during reductive dechlorination of chlorinated ethenes and dihaloelimination of 1,2-dichloroethane. Environ. Sci. Technol. 41:2303-2310. [PubMed]
10. Duhamel, M., and E. A. Edwards. 2006. Microbial composition of chlorinated ethene-degrading cultures dominated by Dehalococcoides. FEMS Microbiol. Ecol. 58:538-549. [PubMed]
11. Duhamel, M., K. Mo, and E. A. Edwards. 2004. Characterization of a highly enriched Dehalococcoides-containing culture that grows on vinyl chloride and trichloroethene. Appl. Environ. Microbiol. 70:5538-5545. [PMC free article] [PubMed]
12. Gu, B., W.-M. Wu, M. A. Ginder-Vogel, H. Yan, M. W. Fields, J. Zhou, S. Fendorf, C. S. Criddle, and P. M. Jardine. 2005. Bioreduction of uranium in a contaminated soil column. Environ. Sci. Technol. 39:4841-4847. [PubMed]
13. He, J., K. M. Ritalahti, K.-L. Yang, S. S. Koenigsberg, and F. E. Löffler. 2003. Detoxification of vinyl chloride to ethene coupled to growth of an anaerobic bacterium. Nature 424:62-65. [PubMed]
14. He, J., Y. Sung, R. Krajmalnik-Brown, K. M. Ritalahti, and F. E. Löffler. 2005. Isolation and characterization of Dehalococcoides sp. strain FL2, a trichloroethene (TCE)- and 1,2-dichloroethene-respiring anaerobe. Environ. Microbiol. 7:1442-1450. [PubMed]
15. Holliger, C., D. Hahn, H. Harmsen, W. Ludwig, W. Schumacher, B. Tindall, F. Vazquez, N. Weiss, and A. J. B. Zehnder. 1998. Dehalobacter restrictus gen. nov. and sp. nov., a strictly anaerobic bacterium that reductively dechlorinates tetra- and trichloroethene in an anaerobic respiration. Arch. Microbiol. 169:313-321. [PubMed]
16. Istok, J. D., J. M. Senko, L. R. Krumholz, D. Watson, M. A. Bogle, A. Peacock, Y.-J. Chang, and D. C. White. 2004. In situ bioreduction of technetium and uranium in a nitrate-contaminated aquifer. Environ. Sci. Technol. 38:468-475. [PubMed]
17. Johnson, B. L., and C. T. DeRosa. 1997. The toxicologic hazard of superfund hazardous waste sites. Rev. Environ. Health 12:235-251. [PubMed]
18. Krajmalnik-Brown, R., T. Hölscher, I. N. Thomson, F. M. Saunders, K. M. Ritalahti, and F. E. Löffler. 2004. Genetic identification of a putative vinyl chloride reductase in Dehalococcoides sp. strain BAV1. Appl. Environ. Microbiol. 70:6347-6351. [PMC free article] [PubMed]
19. Lloyd, J. R. 2003. Microbial reduction of metals and radionuclides. FEMS Microbiol. Rev. 27:411-425. [PubMed]
20. Lloyd, J. R., and J. C. Renshaw. 2005. Bioremediation of radioactive waste: radionuclide-microbe interactions in laboratory and field-scale studies. Curr. Opin. Biotechnol. 16:254-260. [PubMed]
21. Löffler, F. E., J. R. Cole, K. M. Ritalahti, and J. M. Tiedje. 2003. Diversity of dechlorinating bacteria, p. 53-87. In M. M. Häggblom and I. D. Bossert (ed.), Dehalogenation: microbial processes and environmental applications. Kluwer Academic Press, New York, NY.
22. Löffler, F. E., and E. A. Edwards. 2006. Harnessing microbial activities for environmental cleanup. Curr. Opin. Biotechnol. 17:274-284. [PubMed]
23. Löffler, F. E., K. M. Ritalahti, and J. M. Tiedje. 1997. Dechlorination of chloroethenes is inhibited by 2-bromoethanesulfonate in the absence of methanogens. Appl. Environ. Microbiol. 63:4982-4985. [PMC free article] [PubMed]
24. Löffler, F. E., Q. Sun, J. Li, and J. M. Tiedje. 2000. 16S rRNA gene-based detection of tetrachloroethene-dechlorinating Desulfuromonas and Dehalococcoides species. Appl. Environ. Microbiol. 66:1369-1374. [PMC free article] [PubMed]
25. Löffler, F. E., J. M. Tiedje, and R. A. Sanford. 1999. Fraction of electrons consumed in electron acceptor reduction and hydrogen thresholds as indicators of halorespiratory physiology. Appl. Environ. Microbiol. 65:4049-4056. [PMC free article] [PubMed]
26. Lovley, D. R., S. J. Giovannoni, D. C. White, J. E. Champine, E. J. P. Phillips, Y. A. Gorby, and S. Goodwin. 1993. Geobacter metallireducens gen. nov. sp. nov., a microorganism capable of coupling the complete oxidation of organic compounds to the reduction of iron and other metals. Arch. Microbiol. 159:336-344. [PubMed]
27. Ludwig, W., and K.-H. Schleifer. 2000. How quantitative is quantitative PCR with respect to cell counts? Syst. Appl. Microbiol. 23:556-562. [PubMed]
28. Luijten, M. L. G. C., J. de Weert, H. Smidt, H. T. S. Boschker, W. M. De Vos, G. Schraa, and A. J. M. Stams. 2003. Description of Sulfurospirillum halorespirans sp. nov., an anaerobic, tetrachloroethene-respiring bacterium, and transfer of Dehalospirillum multivorans to the genus Sulfurospirillum as Sulfurospirillum multivorans comb. nov. Int. J. Syst. Evol. Microbiol. 53:787-793. [PubMed]
29. Macbeth, T. W., D. E. Cummings, S. Spring, L. M. Petzke, and K. S. Sorenson. 2004. Molecular characterization of a dechlorinating community resulting from in situ biostimulation in a trichloroethene-contaminated deep, fractured basalt aquifer and comparison to a derivative laboratory culture. Appl. Environ. Microbiol. 70:7329-7341. [PMC free article] [PubMed]
30. Major, D. W., M. L. McMaster, E. E. Cox, E. A. Edwards, S. M. Dworatzek, E. R. Hendrickson, M. G. Starr, J. A. Payne, and L. W. Buonamici. 2002. Field demonstration of successful bioaugmentation to achieve dechlorination of tetrachloroethene to ethene. Environ. Sci. Technol. 36:5106-5116. [PubMed]
31. Nevin, K. P., D. E. Holmes, T. L. Woodward, S. F. Covalla, and D. R. Lovley. 2007. Reclassification of Trichlorobacter thiogenes as Geobacter thiogenes comb. nov. Int. J. Syst. Evol. Microbiol. 57:463-466. [PubMed]
32. Nyman, J., M. Gentile, and C. S. Criddle. 2007. Sulfate requirement for growth of U(VI)-reducing bacteria in an ethanol-fed enrichment. Bioremediat. J. 11:21-32.
33. Nyman, J. L., T. L. Marsh, M. A. Ginder-Vogel, M. Gentile, S. Fendorf, and C. S. Criddle. 2006. Heterogeneous response to biostimulation for U(VI) reduction in replicated sediment microcosms. Biodegradation 17:303-316. [PubMed]
34. Pennell, K. D., J. Costanza, F. E. Löffler, and K. E. Fletcher. 2006. Fate of TCE in Fort Lewis soils under thermal remediation conditions: sorption and biotic degradation, poster 132, p. F-111. Abstr. SERDP/ESTCP Partners Environ. Technol. Tech. Symp. Workshop, 28 to 30 November 2006, Washington, DC.
35. Pfaffl, M. W. 2001. A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29:e45. [PMC free article] [PubMed]
36. Ritalahti, K. M., B. K. Amos, Y. Sung, Q. Wu, S. S. Koenigsberg, and F. E. Löffler. 2006. Quantitative PCR targeting 16S rRNA and reductive dehalogenase genes simultaneously monitors multiple Dehalococcoides strains. Appl. Environ. Microbiol. 72:2765-2774. [PMC free article] [PubMed]
37. Ritalahti, K. M., F. E. Löffler, E. E. Rasch, and S. S. Koenigsberg. 2005. Bioaugmentation for chlorinated ethene detoxification: bioaugmentation and molecular diagnostics in the bioremediation of chlorinated ethene-contaminated sites. Ind. Biotechnol. 1:114-118.
38. Robinson, B. S., P. T. Monis, and P. J. Dobson. 2006. Rapid, sensitive, and discriminating identification of Naegleria spp. by real-time PCR and melting-curve analysis. Appl. Environ. Microbiol. 72:5857-5863. [PMC free article] [PubMed]
39. Sanford, R. A., J. R. Cole, and J. M. Tiedje. 2002. Characterization and description of Anaeromyxobacter dehalogenans gen. nov., sp. nov., an aryl-halorespiring facultative anaerobic myxobacterium. Appl. Environ. Microbiol. 68:893-900. [PMC free article] [PubMed]
40. Sanford, R. A., Q. Wu, Y. Sung, S. H. Thomas, B. K. Amos, E. K. Prince, and F. E. Löffler. 2007. Hexavalent uranium supports growth of Anaeromyxobacter dehalogenans and Geobacter spp. with lower than predicted biomass yields. Environ. Microbiol. 9:2885-2893. [PubMed]
41. Scholz-Muramatsu, H., A. Neumann, M. Meβmer, E. Moore, and G. Diekert. 1995. Isolation and characterization of Dehalospirillum multivorans gen. nov., sp. nov., a tetrachloroethene-utilizing, strictly anaerobic bacterium. Arch. Microbiol. 163:48-56.
42. Seguiti, F., C. Drummond, E. Petrovskis, F. E. Löffler, and K. M. Ritalahti. 2006. Management/site closure strategies for a trichloroethene (TCE) impacted aquifer following bioaugmentation, abstr. J-21. In B. M. Sass (ed.), Remediation of chlorinated and recalcitrant compounds. Proceedings of the Fifth International Conference on Remediation of Chlorinated and Recalcitrant Compounds, 21 to 25 May 2006, Monterey, CA.
43. Smidt, H., and W. M. de Vos. 2004. Anaerobic microbial dehalogenation. Annu. Rev. Microbiol. 58:43-73. [PubMed]
44. Sung, Y. 2005. Isolation and ecology of bacterial populations involved in reductive dechlorination of chlorinated solvents. Ph.D. dissertation. Georgia Institute of Technology, Atlanta.
45. Sung, Y., K. E. Fletcher, K. M. Ritalahti, R. P. Apkarian, N. Ramos-Hernández, R. A. Sanford, N. M. Mesbah, and F. E. Löffler. 2006. Geobacter lovleyi sp. nov. strain SZ, a novel metal-reducing and tetrachloroethene-dechlorinating bacterium. Appl. Environ. Microbiol. 72:2775-2782. [PMC free article] [PubMed]
46. Sung, Y., K. M. Ritalahti, R. P. Apkarian, and F. E. Löffler. 2006. Quantitative PCR confirms purity of strain GT, a novel trichloroethene-to-ethene-respiring Dehalococcoides isolate. Appl. Environ. Microbiol. 72:1980-1987. [PMC free article] [PubMed]
47. Sung, Y., K. M. Ritalahti, R. A. Sanford, J. W. Urbance, S. J. Flynn, J. M. Tiedje, and F. E. Löffler. 2003. Characterization of two tetrachloroethene-reducing, acetate-oxidizing anaerobic bacteria and their description as Desulfuromonas michiganensis sp. nov. Appl. Environ. Microbiol. 69:2964-2974. [PMC free article] [PubMed]
48. U.S. Department of Energy. 1999. From cleanup to stewardship (DOE EM-0466). Office of Environmental Management, U.S. Department of Energy, Washington, DC.
49. Wall, J. D., and L. R. Krumholz. 2006. Uranium reduction. Annu. Rev. Microbiol. 60:149-166. [PubMed]
50. Wu, W.-M., J. Carley, M. Fienen, T. Mehlhorn, K. Lowe, J. Nyman, J. Luo, M. E. Gentile, R. Rajan, D. Wagner, R. F. Hickey, B. Gu, D. Watson, O. A. Cirpka, P. K. Kitanidis, P. M. Jardine, and C. S. Criddle. 2006. Pilot-scale in situ bioremediation of uranium in a highly contaminated aquifer. 1. Conditioning of a treatment zone. Environ. Sci. Technol. 40:3978-3985. [PubMed]
51. Wu, W.-M., J. Carley, T. Gentry, M. A. Ginder-Vogel, M. Fienen, T. Mehlhorn, H. Yan, S. Caroll, M. N. Pace, J. Nyman, J. Luo, M. E. Gentile, M. W. Fields, R. F. Hickey, B. Gu, D. Watson, O. A. Cirpka, J. Zhou, S. Fendorf, P. K. Kitanidis, P. M. Jardine, and C. S. Criddle. 2006. Pilot-scale in situ bioremedation of uranium in a highly contaminated aquifer. 2. Reduction of U(VI) and geochemical control of U(VI) bioavailability. Environ. Sci. Technol. 40:3986-3995. [PubMed]
52. Young, A. L., and W. B. Andrews. 2000. The status and challenges of managing risks in the United States Department of Energy's Environmental Management Program. Environ. Sci. Pollut. Res. 2:7-12.
53. Zhou, J., M. A. Bruns, and J. M. Tiedje. 1996. DNA recovery from soils of diverse composition. Appl. Environ. Microbiol. 62:316-322. [PMC free article] [PubMed]

Articles from Applied and Environmental Microbiology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...