Spatial heterogeneity of dechlorinating bacteria and limiting factors for in situ trichloroethene dechlorination revealed by analyses of sediment cores from a polluted field site


  • Editor: Max Häggblom

Correspondence: Christoph C. Tebbe, Institut für Biodiversität, vTI – Bundesforschungsinstitut für Ländliche Räume, Wald und Fischerei, Braunschweig, Bundesallee 50, 38116 Braunschweig, Germany. Tel.: +49 531 596 2553; fax: +49 531 596 2599; e-mail:


Microbiological analyses of sediment samples were conducted to explore potentials and limitations for bioremediation of field sites polluted with chlorinated ethenes. Intact sediment cores, collected by direct push probing from a 35-ha contaminated area, were analyzed in horizontal layers. Cultivation-independent PCR revealed Dehalococcoides to be the most abundant 16S rRNA gene phylotype with a suspected potential for reductive dechlorination of the major contaminant trichloroethene (TCE). In declining abundances, Desulfitobacterium, Desulfuromonas and Dehalobacter were also detected. In TCE-amended sediment slurry incubations, 66% of 121 sediment samples were dechlorinating, among them one-third completely and the rest incompletely (end product cis-1,2-dichloroethene; cDCE). Both PCR and slurry analyses revealed highly heterogeneous horizontal and vertical distributions of the dechlorination potentials in the sediments. Complete reductive TCE dechlorination correlated with the presence of Dehalococcoides, accompanied by Acetobacterium and a relative of Trichococcus pasteurii. Sediment incubations under close to in situ conditions showed that a low TCE dechlorination activity could be stimulated by 7 mg L−1 dissolved carbon for cDCE formation and by an additional 36 mg carbon (lactate) L−1 for further dechlorination. The study demonstrates that the highly heterogeneous distribution of TCE degraders and their specific requirements for carbon and electrons are key issues for TCE degradation in contaminated sites.


Chlorinated ethenes are organic solvents hazardous to human health, but have been used extensively by industrialized countries during the last century. Because of improper handling, soils and sediments at many former industrial or military sites have been contaminated with these solvents. Such contaminations typically persist for many years as dense non-aqueous-phase liquids under anaerobic conditions (McCarty, 1995). Contaminated sites pose a risk because the main pollutants, such as perchloroethene (PCE) and trichloroethene (TCE), or their dechlorination products, for example cis-1,2-dichloroethene (cDCE) and vinyl chloride (VC), may dissolve in groundwater and thereby endanger drinking water supplies. Ideally, all contaminated sites should be cleaned of the chlorinated ethenes, but for most this is not feasible due to the enormous engineering efforts required (Christ et al., 2005).

Research, especially during the last decade, has demonstrated that bacteria have the potential to degrade chlorinated ethenes. Thus, one solution for managing and eventually remediating contaminated sites could be to prove that natural attenuation processes mediated by site-indigenous bacteria are ongoing (Lee et al., 1998; Wiedermeier et al., 1999; Clement et al., 2002). The compensation of factors limiting the indigenous bacteria in degrading chlorinated ethenes or the possibility of increasing the size of metabolically active bacterial populations by bioaugmentation could be a means to enhance such natural purification processes (Ellis et al., 2000; Scow & Hicks, 2005; Hood et al., 2008). Several lines of evidence based on different hydrogeological, chemical, isotopic and microbiological analyses are normally advisable to assess natural attenuation or its potential at a polluted site (McKelvie et al., 2007).

Typically, microbiological assessments of natural attenuation at polluted sites are based on groundwater sample analyses (Davis et al., 2002; Lowe et al., 2002; Connon et al., 2005; Nijenhuis et al., 2007). However, the information gained is limited because it only provides an integrative view of the distribution of pollutants and their degraders and not the spatial patterns that must exist in the sediments (Alfreider et al., 1997). It can be suspected, although still only supported with little analytical evidence, that in fact, natural attenuation processes in a contaminated aquifer depend on the spatial location of the pollutants, electron donors and acceptors, as well as the patchiness of bacterial populations with a potential for reductive dechlorination (Skubal et al., 2001; Davis et al., 2002).

The objective of the present study was, therefore, to explore the diversity and activity of bacterial populations involved in reductive dechlorination of TCE at a polluted site, considering their possible heterogeneous distribution in the sediment. Instead of groundwater analyses, microbiological studies were performed in this study with sediment cores, collected by direct push probing rather than commonly applied drilling. Direct push has the advantage that the spatial structures of the sediment layers are much better preserved (Cho et al., 2000; Costanza & Davis, 2000; Rossabi et al., 2003).

Relevant sampling points were selected at a 35-ha site contaminated with chlorinated ethenes on the basis of depth-dependent measurements of hydrochemical indicators of reductive dechlorination. Indicators include, i.e. the presence of TCE metabolites (cDCE, VC, ethene) and potential alternative electron acceptors (nitrate, sulfate). Additionally, hydrogeochemical analyses of the groundwater were used to complement and understand the microbiological results. The microbiological methods in this study were (1) cultivation-independent PCR detections of 16S rRNA genes of known TCE-degrading bacteria amplified from sediment DNA and enrichment cultures, (2) sediment (slurry) incubations after TCE amendment in the laboratory and detection of metabolites and the dominant 16S rRNA genes and, finally, (3) monitoring TCE dechlorination as it occurs in intact sediment cores maintained under close to in situ conditions and determining the concentration-dependent effect of an added electron donor on that activity.

Materials and methods

Properties of the field site

The field site of this study is located in northeastern Germany in the transition region between the West-Prignitz plateau in the north and the glacial valley of the Elbe River in the south. It was used as a military garrison and airfield by the former Soviet army until 1992. The contaminated near-surface geological structure is characterized by a shift between glacial sandy and clayey layers. The contaminated upper aquifer in the northern part is shallow (10 m), consisting of clay to coarse clay and characterized by a slower groundwater flow (hydraulic conductivity, filtration coefficient kf=2.8 × 10−4 m s−1) than the southern part, where the aquifer is sandy to pebbly up to a depth of 24 m, and the groundwater flow is higher (kf=7 × 10−4 m s−1).

Groundwater sampling and collection of sediment cores

Groundwater was sampled in this study with specially devised double-packer pumps that allowed separation of water samples at distances of 1 m depth.

All microbiological analyses of this study were conducted with sediment material taken from intact cores. The samples were collected by polyvinyl chloride (PVC) liners (4 cm diameter × 50–100 cm length), taking advantage of the direct push technique (Geoprobe®) under an argon atmosphere (Rossabi et al., 2003). The top 10 cm of the sediment cores, selected for molecular analysis, were cut off directly after sampling and immediately stored at −20 °C. The remaining liner length was transported to the laboratory in plastic tubes under argon gas while maintaining it below a maximum of 20 °C. Within 24 h after sampling, sediment samples from the top 3–10 cm of the cores were inoculated under a stream of sterilized reducing gas (90% v/v N2, 10% v/v CO2) into mineral medium to determine the dechlorination potential. The top 2 cm of these sediment cores were discarded and the remaining 40–80 cm sediment material was used to determine the dechlorination rate.

Chemical analyses

Groundwater samples were analyzed for the volatile chlorinated hydrocarbons TCE, cDCE, VC as well as for ethene and methane by GC (GC 8500, Perkin Elmer, Waltham, MA). The instruments were equipped with a headspace sampler (HS 40) with flame ionization detection (FID) and a capillary column (J&W GS-Gaspro, 30 m × 0.32 mm, Agilent, Santa Clara, CA). The following temperature program was used: 5 min at 40 °C, 40 °C min−1 to 180 °C and hold for 15 min. The sediment slurries were tested for TCE, cDCE, VC (Eisenbeis et al., 1997) and ethene by GC, and for pyruvate and lactate by HPLC (Beckman Coulter LC127, UV-Vis detector, 210 nm, guard column Microguard Cation H, 50 mm × 4.6 mm, analytical column Aminex HPX-87 H, 300 mm × 7.8 mm, Biorad, München, Germany; solvent 0.005 mol L−1 H2SO4, flow rate 0.6 mL min−1, 35 °C). Ethene was quantified by GC (GC; Auto System XL, Perkin Elmer) using FID (molecular sieve, 60–80 mesh; 1 m, carrier gas N2, flow rate 40 mL min−1; oven temperature 220 °C).

Incubation of sediment slurries and their TCE dechlorination activity

The dechlorination potential of the sediment samples was measured under batch conditions in sediment slurries (serum bottles, 116 mL, with 70-mL deoxygenated mineral medium of the following composition, in g L−1: KH2PO4 0.1, NH4Cl 0.125, NaCl 0.5, MgCl2·6H2O 0.2, KCl 0.25, CaCl2·2H2O 0.075; trace elements: 0.01 mL L−1 HCl (25%), in mg L−1: FeSO4·7H2O 1.0, ZnCl2 0.07, MnCl2·4H2O 0.1, H3BO3 0.006, CoCl2·6H2O 0.13, CuCl2·2H2O 0.002, NiCl2·6H2O 0.024, NaMoO4·2H2O 0.036, Na2SeO3·5H2O 0.0052, Na2WO4·2H2O 0.0033; vitamins in mg L−1: 4-aminobenzoic acid 0.04, d(+) biotin 0.01, nicotinic acid 0.1, Ca-d(+) pantothenate 0.05, pyridoxamine·2 HCl 0.15, thiamine dichloride 0.1, cyanocobalamine 0.05; carbonate buffer 30 mmol L−1; TCE 50–100 μmol L−1). Because of the low concentration of electron donor in the sandy sediment, sodium pyruvate (5 mmol L−1) was added to the sediment slurries to stimulate dechlorination. Pyruvate was chosen because preceding experiments revealed that under the given conditions, it was more efficient to stimulate complete dechlorination than other more traditional electron donors, i.e. lactate, butyrate, ethanol or methanol (unpublished data). No further organic supplements were added in order to minimize artificial influences.

Sediment samples (5 cm3) were transferred into the culture medium in three replicates. A control (two replicates) consisting of the same amount of sterilized sediment was included to consider sorption and abiotic dechlorination of TCE. The samples and controls were closed with Teflon®-coated butyl rubber septa and aluminum crimp caps and incubated at 20 °C in the dark. After 2 weeks, the cultures were additionally fed with 5 mmol pyruvate L−1. The sediment slurries were tested for biotic and abiotic dechlorination and electron donor oxidation every 2–3 weeks for up to 12 weeks. Sediment slurries with complete dechlorination of TCE to ethene were fed with 5 mmol pyruvate L−1 and doses of 50 μmol TCE L−1 until a total of 300 μmol TCE L−1 was reduced to ethene to increase the yield of dechlorinating bacteria for further investigation.

Incubation of intact sediment cores under close to in situ conditions

The in situ dechlorination rate at sampling point S29 was measured in six intact sediment cores under continuous flow conditions. The sediment cores (4 × 80 cm) were incubated under N2 gas at 15 °C inside a PVC shell (flushed daily with N2) within 24 h after sampling. The native groundwater was deoxygenated (Arcangeli & Arvin, 1995) and pumped peristaltically (enclosed in a PVC box under N2 gas) out of stainless-steel barrels via stainless-steel tubes into the columns at the in situ flow rate [0.5 L day−1, hydraulic residence time (HRT) 24–48 h]. After 92 days of operation, 10 mmol lactate L−1 was added as a potential electron donor and carbon source to the groundwater inflow of two of the sediment cores in order to determine the influence of the electron donor on the dechlorination process.

Water samples were taken using gas-tight syringes (Hamilton, Reno, NV) from the columns and analyzed for the chlorinated ethenes, lactate (inflow and effluent) and for ethene (effluent). The dechlorination rates, TCE to cDCE and cDCE to VC, were calculated from differences in their concentrations between in- and outflow of the sediment cores using mass balances (zero-order rate constant).

DNA extraction from sediment and PCR

DNA was extracted from 10 g (wet weight) of sediment material using the MoBio Soil DNA Kit (Mega Prep, MoBio, Dianova, Hamburg) as suggested by the manufacturer, including an ethanol precipitation step. DNA was purified using the Wizard DNA Clean Up System (Promega, Mannheim, Germany) and eluted in 50 μL of TE (10 mM Tris, 1 mM EDTA, pH 8). Sediment slurries (70 mL), obtained from the incubated sediment samples for determination of anaerobic TCE dechlorination, were centrifuged (Eppendorf Centrifuge 5403, 4100 g, 15 min), yielding approximately 0.5 g of pellet material (wet weight). DNA from approximately 0.5 g wet weight of these pellets was then extracted using the protocol described by Lueders et al. (2004). The extracted DNA was resuspended in 50 μL 10 mM Tris-HCl (pH 8.5). All DNA extracts were stored at −20 °C until further analyses.

The presence of Dehalococcoides, Desulfitobacterium, Dehalobacter and suspected TCE-dechlorinating Desulfuromonas populations was evaluated using specific 16S rRNA gene-targeted primer pairs. Dehalococcoides- and Desulfitobacterium-specific primers were designed in this study using the arb software environment based on the ssujun02 database updated for 16S rRNA gene sequences of Desulfitobacterium and Dehalococcoides from public databases (NCBI). The specificity of the selected primers was examined in silico using the probe-check function of the arb software (Ludwig et al., 2004), the probe-match tool of the Ribosomal Database Project (RDP) (Cole et al., 2003) and blast searches (NCBI; of the primer sequences. Dehalococcoides-specific PCR was performed using the primers dhc193f (5′-GGTTCAYTAAAGCCGYAAGG-3′) and dhc1048r (5′-CCTGTGCAARYTCCTGACT-3′) and PCR cycling conditions as follows: initial denaturation at 95 °C for 15 min; 30 cycles of 95 °C for 30 s, annealing at 53 °C for 30 s and extension at 72 °C for 60 s; and a final extension at 72 °C for 10 min. For Desulfitobacterium-specific PCR, the primers dsb434f (5′-TACTGTCTTCAGGGACGAAC-3′) and dsb1299r (5′-TGAGACCAGCTTTCTCGGAT-3′) were chosen. Thermocycling conditions were identical to those of dhc193f and dhc1048r, except for annealing, which was performed at 60 °C. The Desulfitobacterium- and Dehalococcoides-targeted primers yielded amplicons of 865 and 855 bp, respectively.

PCR specific for Dehalobacter 16S rRNA genes was performed using the primers deb179f (5′-TGTATTGTCCGAGAGGCA-3′) and deb1007r (5′-ACTCCCATATCTCTACGG-3′) (Schlotelburg et al., 2002). For the detection of Desulfuromonas, the forward primer 5′-AACCTTCGGGTCCTACTGTC-3′ (Escherichia coli 16S rRNA gene position 205–222) and reverse primer 5′-GCCGAACTGACCCCTATGTT-3′ (1033–1015) were used (Löffler et al., 2000). PCR cycling conditions were performed as described by the authors, with the exception that the initial denaturation step at 95 °C was extended to 15 min. All PCR reactions were carried out in a total volume of 25 μL containing 1 μL of extracted DNA (1 : 10 dilution of original DNA solution), 0.2 mM of each deoxynucleoside triphosphate (Qbiogene, Heidelberg, Germany), 0.5 μM of each primer (MWG Biotech, Ebersberg, Germany) and 1.25 U Taq polymerase (Hotstar Taq, Qiagen) with the corresponding 1 × PCR buffer containing 1.5 mM MgCl2. Reaction mixtures for Desulfuromonas-specific PCR contained a final concentration of 2.5 mM MgCl2. For the amplification of DNA extracted from sediments, the T4 gene 32 protein (T4gp32; Roche Applied Science, Mannheim, Germany) was added at a final concentration of 25 ng μL−1 to each reaction to overcome the inhibitory effects of coextracted substances (Tebbe & Vahjen, 1993; Vahjen & Tebbe, 1994). Positive controls used in this study were genomic DNA of Dehalococcoides ethenogenes strain 195 (kindly provided by Ivonne Nijenhuis, Helmholtz-Zentrum für Umweltforschung UFZ, Leipzig, Germany), Desulfitobacterium strain PCE-S, Dehalobacter restrictus (DSMZ 9455) and Desulfuromonas chloroethenica (DSMZ 12431), the latter two purchased from DSMZ (Braunschweig, Germany). The specificity of the primers was also evaluated and confirmed with negative DNA controls originating from bacterial pure cultures and rhizosphere soils, respectively. Rhizosphere soils were included because the presence of the targeted bacteria in detectable amounts was unlikely, and was, in fact, never detected (data not shown). The specificity of the PCR detections was confirmed for Dehalococcoides by amplicon sequencing and for Desulfitobacterium by single-strand conformation polymorphism (SSCP) gel electrophoresis (conditions see below) and comparison with positive controls from a pure culture. Primer specificity of the other two dechlorinating bacteria had already been established in other studies (Löffler et al., 2000; Schlotelburg et al., 2002). All PCR amplifications were conducted in a Primus 96 thermocycler (MWG Biotech). Nested PCR was applied to increase the sensitivity of detection for the targeted bacteria in sediment DNA. In these nested PCR, a 1 : 25 dilution for sediment DNA or a 1 : 250 diluted aliquot for DNA from enrichment cultures from the first PCR was included as a template in the second PCR. PCR conditions were as described above, except that T4gp32 was not added. For the second PCR, the universal bacterial 16S rRNA gene primers Com1 and Com2-Ph (Schwieger & Tebbe, 1998) were used, and the following thermocycling conditions were applied: 15 min at 95 °C, followed by 25 cycles of 60 s at 95 °C, 60 s at 50 °C, 70 s at 72 °C and, for the last cycle, 5 min at 72 °C. All PCR products were analyzed for size and yield by electrophoresis in a 1% agarose gel stained with ethidium bromide (Sambrook & Russell, 2001).

The application of universal primers for the second PCR required additional controls to exclude false-positive detection due to the transfer of template DNA (with no specific PCR product) from the first PCR. Therefore, for the analyses of sediment DNA, a 1 : 25 dilution of the original template, corresponding to the dilution caused by the first PCR, was further diluted 1 : 25, as carried out for all samples after PCR amplification with ‘dechlorinator’-specific primers, and then directly added to the second PCR as a template. Consequently, for the examination of DNA from enrichment cultures, a 1 : 25 diluted community DNA was further diluted 1 : 250 and then added as a control. This allowed detection of false-positive amplification caused by contamination of template DNA from the first DNA. Absence of target DNA from the putative dechlorinating bacteria was indicated by negative PCR results.

To exclude false-negative detection of the targeted bacterial groups from sediment DNA, each extracted DNA was tested separately for PCR inhibition by coextracted humic compounds (Tebbe & Vahjen, 1993) using universal primers Com1 and Com2-Ph under the conditions described above. Only DNA with no inhibitory effect (which was the rule) was considered for further analyses in search of potentially TCE-dechlorinating bacteria.

SSCP and DNA sequencing

Bacterial communities enriched from TCE-amended sediment slurries were analyzed by genetic profiles with SSCP, using the single-strand removal method (Schwieger & Tebbe, 1998), as described in detail elsewhere (Dohrmann & Tebbe, 2004). Briefly, partial 16S rRNA genes of the bacterial communities were amplified directly from total nucleic acid by PCR using primers Com1 (nonphosphorylated) and Com2-Ph (phosphorylated) and subsequently, the phosphorylated DNA strand was removed by λ-exonuclease digestions. The concentrations of the PCR mixture (50 μL; 2 μL diluted template DNA) and the cycling conditions were used as described above for the specific nested PCR approach, except that 30 cycles were used instead of 25 cycles. SSCP markers were generated from bacterial pure cultures by PCR using Com1 and Com2-Ph and subsequent λ-exonuclease digestions. The single-strand PCR products were separated in polyacrylamide gels (MDE, FMS Bioproducts) using a PerfectBlue dual gel system Twin L (Peqlab Biotechnologie, Erlangen, Germany) set at 400 V and a running temperature of 20 °C for 6800 V h. After the electrophoretic run, the gels were silver-stained for visualizing the DNA (Bassam et al., 1991).

DNA sequencing from bands of SSCP gels was performed as described previously (Dohrmann & Tebbe, 2004; Brinkmann et al., 2008). Bands of interest were cut and eluted from the silver-stained gels. The single-strand DNA was amplified by PCR using primers Com1 and Com2-Ph. The resulting PCR products were cloned and DNA of clones containing vector inserts was extracted. The inserted DNA was PCR-amplified with primers Com1 and Com2-Ph, and PCR products were again subjected to SSCP analysis to compare the band position in the original community profile. Only clones carrying the expected inserts were analyzed further by DNA sequencing. DNA sequences were analyzed by cycle sequencing (Schmalenberger et al., 2001). Sequences were aligned using the arb software package, checked for chimera by pintail ( (Ashelford et al., 2005) and compared with sequences from databases using the blastn search analysis (

Nucleotide sequence accession numbers

All sequences obtained in this study have been deposited in the database of the European Bioinformatics Institute ( The respective accession numbers are listed in Results.


Site exploration for the presence of chlorinated ethenes, nitrate, sulfate and dissolved organic carbon (DOC) in groundwater

A total of 101 monitoring wells covering the suspected polluted site of approximately 35 ha were established at the beginning of the study (Fig. 1a). After the monitoring wells were constructed, 3 months were allowed to restore the groundwater flow (main direction: north to south), before groundwater sampling was carried out for chemical analyses. The samples were analyzed for the chlorinated ethenes TCE, cDCE and VC, as well as for ethene and other chemical parameters (Supporting Information, Table S1). The maximum values of TCE were 40 mg L−1, 12 mg L−1 for cDCE, 0.06 mg L−1 for VC and 0.28 mg L−1 for ethene (Fig. 1b–d; Table S1).

Figure 1.

 Map of the TCE-contaminated field site with sampling points of this study indicated as numbers. (a) Gray shaded areas show the two main regions of contamination. (b) Maximum concentrations of TCE, (c) cDCE and (d) VC explored by groundwater analyses from filtered wells (maximum concentrations per well indicated). The main direction of the groundwater flow was from north to south.

Two contaminated areas could be distinguished: one in the north, encompassing the prospective sampling points S1–S28, with relatively low concentrations of TCE, but higher levels of cDCE and VC, and one in the south, including S43–S67, with higher concentrations of TCE and cDCE, but lower concentrations of VC (Fig. 1). While nitrate (not detectable) and sulfate concentrations (<100 mg L−1) were relatively low in the northern part, they were elevated in the southern part, with 1–20 mg nitrate L−1 groundwater and 50–700 mg sulfate L−1, respectively (Table S1). The concentration of DOC ranged between 5 and 25 mg L−1, with no specific regions of higher or lower concentrations.

Detection of suspected TCE-dechlorinating bacteria from sediment samples by PCR

A total of 23 sampling points were selected to obtain sediment cores. Total DNA was directly extracted from sediment samples of different depths of each core in order to characterize the presence of bacterial phylotypes with a known potential to reductively dechlorinate TCE, either partially (Dehalobacter, Desulfuromonas, Desulfitobacterium) to cDCE or completely to ethene (Dehalococcoides). To increase the sensitivity of detection, nested PCR of partial 16S rRNA genes, first amplifying the targeted group with specific primers, and, then enhancing the PCR products, with a subsequent PCR using universal bacterial primers were applied (for specificity and controls, see Materials and methods). The lower detection limit, as established with sediment-amended dilutions of D. ethenogenes as a positive control, was 15 cells g−1 sediment (data not shown).

Out of a total of 108 sediment samples analyzed, only 19% were completely negative for any of the targeted bacteria (Table 1). Dehalococcoides was detected in 73% of all samples, Desulfitobacterium in 29%, Desulfuromonas in 19% and Dehalobacter in 17%, respectively. One single core (S49) was completely negative for any of the targeted bacteria, but only two depths were analyzed for this core. The PCR results for the other cores varied with depths, indicating a heterogeneous distribution of the suspected dechlorinating bacteria within vertical distances of only 10–20 cm within the sediment. None of the sediment samples indicated the simultaneous presence of all four targeted bacteria. Interestingly, almost all sediments (97%) with Desulfitobacterium were also positive for Dehalococcoides. This was not the case with Desulfuromonas and Dehalobacter, which only coincided with Dehalococcoides in 37% and 17% of the cases, respectively.

Table 1.   Analysis of the spatial heterogeneity of putative dechlorinating bacteria and anaerobic dechlorination potentials for TCE in sediment cores sampled across a polluted field site
PCR detection of putative TCE dechlorinating bacteria*Anaerobic dechlorination of TCE in slurry incubations
Depth (m)DebDsmDsbDhcDepth (m)cDCEEthene§
  • Detection of putative dechlorinating bacteria by nested PCR from directly extracted sediment DNA using primers specific for Dehalobacter (Deb), Desulfuromonas (Dsm), Desulfitobacterium (Dsb) and Dehalococcoides (Dhc). Anaerobic dechlorination was determined in sediment slurries amended with 50–100 μM TCE and 5 mM pyruvate as an electron donor and carbon source. Incomplete dechlorination was indicated by the metabolite cDCE and complete dechlorination by detection of ethene.

  • *

    +, strong PCR product after nested PCR; (+), weak product; −, no product.

  • +, formation of the respective metabolite; − the metabolite was not detected within 90 days of incubation.

  • Depths below soil surface, sediment samples represented by mixed material from up to 10 cm above the indicated depths.

  • §

    § Ethene detection was always accompanied by detection of VC.

Sampling point 1
Sampling point 2
Sampling point 5
Sampling point 9
Sampling point 13
Sampling point 15
Sampling point 20
Sampling point 21
Sampling point 22
Sampling point 23
Sampling point 24
Sampling point 28
Sampling point 30
Sampling point 31
Sampling point 32
Sampling point 33
Sampling point 43
Sampling point 44
Sampling point 46
Sampling point 48
Sampling point 49
Sampling point 52
Sampling point 62

In addition to S49, sediment samples from S31 and S32 were also characterized by mainly negative PCR results, suggesting that populations of the suspected dechlorinating bacteria were very low at these points. Interestingly, all three of these sampling points were outside of or at the borders of the areas where dechlorination products had been detected.

TCE dechlorination potential of sediment slurries in the presence of pyruvate

The potential of site-indigenous bacteria to degrade TCE was tested with a total of 121 sediment samples from the same 22 sediment cores that were analyzed by PCR, plus five additional cores. The lower detection limit of the slurry method was 100 cells g−1 sediment dry weight as established with slurry-inoculated dilutions of Desulfitobacterium strain PCE-S cells (data not shown).

As judged by the decline of TCE in these sediment slurries, and the concomitant formation of dechlorination products after an incubation period of up to 90 days, 66% of the samples were positive, and of these, 32% indicated complete dechlorination (formation of ethene), while the other samples formed cDCE from TCE (incomplete dechlorination) (Table 1; results from five sampling points, encompassing 13 samples, which were not analyzed by PCR or included in the table). None of the sediment samples showed ethene formation without transient formation of cDCE and VC. Within most sediment cores, the potential to dechlorinate TCE varied considerably with depth. Exceptions, however, were found with cores from sampling points S1, S2 and S23, in which sediment samples from (almost) all depths indicated the potential for complete reductive dechlorination. The elevated concentrations of VC (Fig. 1d) and ethene at the field site reported above were in accordance with the dechlorination potentials found here.

The sediment slurries from sediment cores in the southern part of the contaminated site (e.g. S32, S44 and S46) consistently indicated incomplete dechlorination (only cDCE formation from TCE), which was in agreement with increased concentrations of cDCE and TCE and very low VC concentrations found in the groundwater at these points (Fig. 1b–d).

Bacterial diversity enriched in sediment slurries with TCE and pyruvate

The sediment slurries that were used to characterize dechlorination potentials in the presence of TCE and pyruvate were also analyzed for their bacterial composition using cultivation-independent genetic profiles (SSCP technique) of PCR-amplified partial 16S rRNA genes. Compared with SSCP profiles obtained from directly extracted sediment DNA (data not shown), the SSCP profiles from these enrichments were composed of fewer, but more pronounced bands, indicating selection of specific members of the bacterial community during the slurry incubations. High consistency was seen in the dominant bands in SSCP profiles from the enrichment cultures with complete dechlorination (Fig. 2a), while profiles from incubated sediments with incomplete dechlorination were more variable (Fig. 2b).

Figure 2.

 SSCP profiles of PCR-amplified partial bacterial 16S rRNA genes from incubated sediment slurries (enrichment cultures) with TCE as an electron acceptor and pyruvate as an electron donor and carbon source. Bands selected for DNA sequencing are indicated by arrows. Sequence information is given in Table 2. Lane R shows reference strains Desulfitobacterium strain PCE-S (A), Desulfuromonas chloroethenica (B), Dehalobacter restrictus (C) and Dehalococcoides ethenogenes 195 (D). M, lane with SSCP markers; NC, negative control. Lanes indicated with E (a) show bacterial community profiles of enrichment cultures with complete dechlorination of TCE (formation of ethene), and those indicated with D (b) of enrichment cultures with an incomplete dechlorination (formation of cDCE; D2, D4–D6, D8–D10) or no dechlorination (D1, D3, D7).

DNA sequencing of the three consistently dominant bands of the SSCP profiles from completely dechlorinating enrichment cultures revealed the presence of Dehalococcoides, an Acetobacterium and a close relative of Trichococcus pasteurii, with 99–100% identity to sequences in the GenBank database (Table 2). Other sequences retrieved from inconsistently occurring bands from completely dechlorinating cultures were related to Clostridia and Proteobacteria (classes Beta-, Gamma- and Deltaproteobacteria). Most sequences, i.e. six out of 10, retrieved from the cultures with only incomplete dechlorination, were related to members of the Enterobacteriales (Gammaproteobacteria). In addition, sequences related to Lactobacilli, Clostridia and a Bacteroidetes were detected. No band in these SSCP profiles corresponded to the three dominant bands seen in the completely dechlorinating cultures.

Table 2.   Characterization of bacteria from incompletely and completely dechlorinating enrichment cultures inoculated with sediment samples, based on PCR-amplified bacterial SSU rRNA genes and database comparison of their DNA sequences from dominant SSCP bands (see Fig. 2)
Band numberClosest relative in GenBank databasePhylum, class and orderSimilarity
DNA sequences from completely dechlorinating cultures (formation of ethene)
1AAcetobacterium carbinolicum VNs25Firmicutes, Clostridia, Clostridiales100E6FM994628
1BAcetobacterium carbinolicum VNs25Firmicutes, Clostridia, Clostridiales99.7E11FM994629
2A,B,CTrichococcus pasteurii R-31594Firmicutes, Bacilli, Lactobacillales100E6, E11, E14FM994630
3A,BDehalococcoides sp. BAV1Chloroflexi, Dehalococcoidetes100E6, E14FM994633
4APseudomonas stutzeri DN7Proteobacteria, Gammaproteobacteria, Pseudomonadales99.5E11FM994635
4BPseudomonas stutzeri DN7Proteobacteria, Gammaproteobacteria, Pseudomonadales99.2E13FM994636
5AAeromonadales bacterium TP662 (unclassified)Proteobacteria, Gammaproteobacteria, Aeromonadales98.9E11FM994637
5BAeromonadales bacterium TP662 (unclassified)Proteobacteria, Gammaproteobacteria, Aeromonadales99.7E11FM994638
6Geobacter sp. TMJ1Proteobacteria, Deltaproteobacteria, Desulfuromonadales99.2E14FM994639
7ASporotalea colonicaFirmicutes, Clostridia, Clostridiales100E11FM994640
7BUncultured Sporotalea sp. clone 175Firmicutes, Clostridia, Clostridiales100E14FM994641
8Rahnella aquatilisProteobacteria, Gammaproteobacteria, Enterobacteriales100E14FM994642
9Variovorax sp. 1-O-1Proteobacteria, Betaproteobacteria, Burkholderiales100E14FM994643
DNA sequences from incompletely dechlorinating cultures (formation of cDCE with no ethene)
10Lactobacillales bacterium HY-36-1 (unclassified)Firmicutes, Bacilli, Lactobacillales95.9D4FM994645
15Serratia sp. AC-CS-1BProteobacteria, Gammaproteobacteria, Enterobacteriales100D4FM994651
16Rahnella aquatilisProteobacteria, Gammaproteobacteria, Enterobacteriales100D4FM994652
17Raoultella terrigena KNUC166Proteobacteria, Gammaproteobacteria, Enterobacteriales100D4FM994653
18Enterobacter asburiae XJUHX-5Proteobacteria, Gammaproteobacteria, Enterobacteriales100D4FM994654
11Desulfosporosinus sp. 44A-T3aFirmicutes, Clostridia, Clostridiales100D6FM994646
12Pseudomonas stutzeri DN7Proteobacteria, Gammaproteobacteria, Pseudomonadales99.4D6FM994647
13Uncultured Bacteroidetes bacterium clone HI2Bacteroidetes99.4D6FM994648
14AKlebsiella sp. PSB-WS1-8Proteobacteria, Gammaproteobacteria, Enterobacteriales98.1D6FM994649
14BKlebsiella sp. PSB-WS1-8Proteobacteria, Gammaproteobacteria, Enterobacteriales100D6FM994650
19Enterobacter sp. S6BBProteobacteria, Gammaproteobacteria, Enterobacteriales99.7D6FM994655

Enrichment cultures were also analyzed for the presence of putative dechlorinating bacteria as indicated by detection of their specific 16S rRNA genes. While all completely dechlorinating cultures produced a strong PCR product for Dehalococcoides, this organism was never detected in cultures where only cDCE was formed (data not shown), which supported the observation of the missing Dehalococcoides band in those SSCP profiles. In contrast, the bacterial species with suspected potential for incomplete dechlorination were infrequently detected in both completely and incompletely dechlorinating cultures. Desulfitobacterium was more often detected than Desulfuromonas. Dehalobacter was only detected in one single sample. Several enrichment cultures (D1, D8, D9 and D10) were negative for all targeted bacteria, suggesting that other bacteria were involved in the formation of cDCE from TCE, for example Geobacter sp. (Duhamel & Edwards, 2006), or other not yet characterized dechlorinating bacteria.

Dechlorination activities of intact sediment cores under close to in situ conditions

To estimate dechlorination activities in situ and to study the effect of an added electron donor and carbon source on the TCE dechlorination rates, intact sediment cores were taken from a TCE-contaminated area adjacent to S29, where only incomplete dechlorination to cDCE had been detected (see Table 1). The cores were operated with groundwater (DOC 7 mg L−1) at a flow rate of 20 mL h−1 (HRT 6.5 h) containing TCE at concentrations between 13 and 32 μmol L−1 (Fig. 3a). After 25 days, the initially low in situ dechlorination activity (TCE to cDCE 0.02–0.3 μmol L−1 h−1, no formation of ethene) rapidly increased to a maximum rate that was limited by the TCE concentration of the inflowing groundwater (e.g. 1.8 μmol L−1 h−1 at 106 days).

Figure 3.

 Reductive dechlorination of TCE in an intact sediment core transferred to the laboratory and incubated with TCE-contaminated groundwater from the sampling site (for details, see Materials and methods). (a) TCE concentrations of the inflowing groundwater. (b) Conversion rates of TCE to cDCE and VC and responses to the addition of lactate to the groundwater (0.5–4.8 mM effluent concentration) after 92 days of incubation. The day of addition is indicated by an arrow.

Lactate addition to the groundwater (effluent concentration 1–5 mmol L−1) after 92 days of incubation resulted in an immediate VC formation from TCE up to a maximum rate of 0.48 μmol L−1 h−1 in both cores analyzed (Fig. 3b). With decreasing lactate concentrations, the VC-formation rate also decreased and no VC was detected at 0.6 mmol lactate L−1 (124 days). Complete reductive dechlorination to ethene started at a low rate (0.1 nmol mL−1 day−1) after approximately 100 days (data not shown). These low rates did not respond to the addition of lactate.


Concentrations of TCE and its metabolites cDCE and VC found in the groundwater suggested that TCE dechlorination was more efficient in the northern area than in the southern area of the polluted site of this study. Possibly, the lower groundwater flow rate with 2.8 × 10−4 m s−1 in the northern area allowed a more efficient degradation of DOC sources under reducing conditions. While electron acceptors such as nitrate and sulfate that compete with reductive dechlorination for hydrogen were absent or in low concentrations in the northern area, they were present in fairly high amounts in the southern part, which correlates with higher TCE concentrations. Reductive dechlorination of TCE to cDCE should be precluded in the presence of nitrate (Lovley, 1985; Cord-Ruwisch et al., 1998), whereas sulfate reduction and dechlorination of cDCE and VC can coexist because of their demand for similar hydrogen concentrations (Hoelen & Reinhard, 2004). Despite the presence of nitrate, however, cDCE and traces of VC were detected in the southern area, which could indicate that incomplete dechlorination took place, even though it cannot completely be ruled out that metabolites may have also been transported by groundwater from another active zone. The coexistence of competing metabolic processes within centimeter distances can be explained by suspected spatial heterogeneities in sediments, which may account for different microhabitats, redox conditions and metabolic processes.

To explore such heterogeneities in more detail, the microbiological analyses of this study were based on sediment samples rather than groundwater. Sediment cores were collected from different points of the 35-ha area and divided into 10-cm-thick horizontal layers. These sediment layers were then used for both cultivation-independent detection of bacteria suspected to reductively dechlorinate TCE and to determine the potential of sediment material to dechlorinate TCE in liquid media containing a carbon source and an electron donor. Both approaches revealed high heterogeneity in both dimensions, horizontally within the area on a meter scale and, vertically, with depths on a centimeter scale.

PCR detection of 16S rRNA genes directly amplified with specific primers from sediment DNA revealed a wide abundance of bacteria with a potential to reductively dechlorinate TCE within the area of study. The relative abundance of these phylotypes within the sediment-inhabiting bacterial community, however, was low, as indicated by the fact that nested PCR was necessary to amplify their specific 16S rRNA genes. Among the 16S rRNA gene types, Dehalococcoides was most frequently detected in the sediment samples. The others, Dehalobacter, Desulfuromonas and Desulfitobacterium, were also detected, but they occurred less frequently. In another study, a groundwater-based exploration of a TCE-contaminated site, encompassing three of the four phylotypes targeted here, also revealed the presence of Dehalococcoides in nearly all the samples examined. Dehalobacter and Desulfitobacterium were found to be less abundant (Nijenhuis et al., 2007). Interestingly, in this study, the detection of Desulfitobacterium was highly correlated with the detection of Dehalococcoides, while this was not the case with either Desulfuromonas or Dehalobacter. This indicates that Desulfitobacterium and Dehalococcoides have similar in situ preferences and that they may even collaborate with each other metabolically. Correlations between two species at a spatial scale, as observed here, would have probably been overlooked with common groundwater analyses.

Only three sediment cores were characterized by negative PCR results for all dechlorinating bacteria targeted in this study, and those were all located at the border of the contaminated areas. This suggests that selection processes for dechlorinating bacteria took place in the presence of TCE or its metabolites. However, once the bacterial populations have been selected they may, due to slow decay rates, persist much longer in the sediment than the pollutants for which they had been selected. This may explain, in addition to the sampling error caused by the enormous field heterogeneity, why the detection of Dehalobacter, Desulfuromonas, Desulfitobacterium or Dehalococcoides could not directly be linked to a specific dechlorination pattern (complete or incomplete) or to in situ dechlorination activities. Therefore, rRNA or mRNA rather than DNA-based techniques could be more promising to develop molecular tools for the detection of natural attenuation due to their lower half-life value and better link to metabolic activity (Lee et al., 2008).

Anaerobic incubations of sediment slurries amended with TCE and pyruvate also demonstrated wide abundance and spatial heterogeneity of reductively dechlorinating bacteria in the sediments. While incomplete dechlorination with cDCE as an end product was indicated in 70% of the 27 sediment cores analyzed, complete dechlorination with the formation of ethene was found in only 30%. It should be noted that a total of 121 sediment samples were analyzed from these 27 cores. The potential for complete dechlorination was mainly found with samples from sediment cores of the northern part, which was in accordance with the conclusions drawn from the hydrogeochemical analyses described above.

To further link both methods, PCR detection and activity measurements from sediment slurry incubations with TCE, the bacterial community structures found in slurry incubations with pyruvate after 6 months of incubation were analyzed by PCR–SSCP. Completely dechlorinating enrichments were, in fact, always colonized by Dehalococcoides, accompanied by two bacterial phylotypes, i.e. a suspected homoacetogenic Acetobacterium and Trichococcus pasteurii. The same phylotypes were detected in anaerobic enrichments of a trichlorodibenzo-p-dioxin-dechlorinating mixed culture with a Dehalococcoides as putative dechlorinating species, underlining their importance as syntrophic partners (Ballerstedt et al., 2004). The striking absence of Dehalococcoides in all incompletely dechlorinating cultures and 100% presence in the completely dechlorinating ones confirms that the organism is a good indicator for complete dechlorination. This despite the fact that also cometabolically VC to ethene dechlorinating strains such as D. ethenogenes strain 195 have been described in the literature (Maymo-Gatell et al., 2001). Possibly, strains such as D. ethenogenes strain 195 may be less competitive compared with those Dehalococcoides strains able to obtain energy out of the complete dechlorination at the site investigated in this study and other sites that demonstrated incomplete dechlorination in the absence of Dehalococcoides (Fennell et al., 2001; He et al., 2003; Yoshida et al., 2007).

Despite the high genetic and laboratory-proven potential for complete dechlorination found here, the presence of TCE in the sediments demonstrated that its dechlorination was severely hampered under field conditions. Limiting factors related to the site-specific physicochemical and hydrological conditions can be held responsible (Song et al., 2002; Macbeth et al., 2004). In this study, the close to in situ conditions used on intact sediment cores run with site-collected groundwater showed, even without carbon source amendments, TCE to cDCE dechlorination (1–2 μmol L−1 h−1). This activity could transiently be stimulated by addition of lactate. Dechlorination of cDCE to VC was initiated immediately after addition of 0.5 mmol L−1 lactate. Interestingly, the low ethene formation (approximately 0.1 nmol L−1 day−1) observed before did not respond equally. In other studies, dechlorination of perchloroethene to cDCE occurred at lactate concentrations above 0.7 mmol L−1 in sediment columns inoculated with Dehalococcoides, whereas conversion of VC to ethene required more than twofold higher concentrations (Behrens et al., 2008). Thus, increasing concentrations of DOC should stimulate complete dechlorination of ethenes at the contaminated site of this study.

In conclusion, the microbiological analyses of sediment samples collected from intact cores revealed a high spatial variability of the presence of potentially dechlorinating bacteria and TCE-dechlorination activity within a centimeter scale of depths. Compared with groundwater analyses, the investigation of sediment material has the advantage that functional units of a microbial community are better preserved and major limiting factors can be explored under more realistic conditions. A further understanding of the diversity of TCE-metabolizing bacteria, their specific activities and interactions and their spatial distribution in contaminated aquifers, as it can be obtained from intact sediment core analyses, should provide better knowledge for devising and optimizing management practices of contaminated field sites in the future.


We thank Evelin Schummer and Suse Gaiser for their excellent technical assistance. We also thank Sybille Grandel and Andreas Dahmke (University of Kiel) for their supporting interest in this project and many stimulating discussions. The study was part of the KORA initiative (Retention and degradation processes to reduce contaminants in groundwater and soil;, supported by the German Ministry for Education and Research, Projektträger Forschungszentrum Karlsruhe, Bereich für Wassertechnologie und Entsorgung (PKTA-WTE; Project numbers 02WN0370–02WN0372).