Occurrence of Dehalococcoides and ReductiveDehalogenase Genes in Microcosms, a ConstructedWetland and Groundwater from a Chlorinated EtheneContaminated Field Site as Indicators for In Situ ReductiveDehalogenation

Éva Mészáros & Gwenaël Imfeld &

Marcell Nikolausz & Ivonne Nijenhuis

Received: 17 May 2013 /Accepted: 30 September 2013 /Published online: 18 October 2013# Springer Science+Business Media Dordrecht 2013

Abstract Thus far, members of the genus Dehalo-coccoides are the only microorganisms known todehalogenate chlorinated ethenes to ethene and therebydetoxify these common groundwater pollutants. Therefore,it is important to characterize the taxonomic and functional

diversity of these key microorganisms and their reductivedehalogenase (RDase) genes in contaminated aquifers forassessing the natural attenuation potential. Little is knownabout the diversity of RDase genes under field conditions orin laboratory systems under selective pressure during de-chlorination activities. Here, we evaluate the diversity ofDehalococcoides sp. and threeRDase genes in groundwateras well as in water from a constructed wetland and micro-cosms setupwith contaminated groundwater from the samefield site in Bitterfeld (Saxony-Anhalt, Germany). The pres-ence and relative abundance of Pinellas and Cornell sub-groups ofDehalococcoideswas evaluated by a novel directsequencing method, which revealed that all sequences wereidentical and affiliated to the Pinellas subgroup. Contrarily,our results showed remarkable differences at the functionalgene level between the systems. Of the vinyl chloridereductase genes, vcrA was detected in samples from thegroundwater, wetland, and microcosms, whereas bvcAwasonly found in wetland and microcosm samples. Thetrichloroethene dehalogenase gene, tceA could not be de-tected at all, although complete dehalogenation activity ofhigher chlorinated ethenes was observed. Our study dem-onstrates that although the Dehalococcoides 16S rRNAgene sequences retrieved from the investigated systemswere identical, the RDase gene diversity varied among thesystems, according to the spectrum of the chlorinated eth-enes present.

Keywords RDase genes . Reductive dehalogenation .

Chlorinated ethenes . Bitterfeld . Direct sequencing

Water Air Soil Pollut (2013) 224:1768DOI 10.1007/s11270-013-1768-x

Electronic supplementary material The online version of thisarticle (doi:10.1007/s11270-013-1768-x) containssupplementary material, which is available to authorized users.

É. MészárosDepartment of Microbiology, Eötvös LorándUniversity of Science,Budapest, Hungary

É. Mészáros :G. Imfeld : I. Nijenhuis (*)Department of Isotope Biogeochemistry,Helmholtz Centre for Environmental Research – UFZ,Leipzig, Germanye-mail: ivonne.nijenhuis@ufz.de

G. ImfeldLaboratory of Surface Hydrology and Geochemistry(LHyGeS – UMR 7517), University of Strasbourg/ENGEES, CNRS,Strasbourg, France

M. NikolauszDepartment of Environmental Biotechnology,Helmholtz Centre for Environmental Research – UFZ,Leipzig, Germany

M. NikolauszDepartment of Bioenergy, Helmholtz Centre forEnvironmental Research – UFZ,Leipzig, Germany

1 Introduction

Chlorinated ethenes are among the most common con-taminants observed in groundwater systems worldwide(Bradley 2003; Christ et al. 2005; Löffler & Edwards2006; Moran et al. 2007), which tend to persist inanaerobic subsurface environments (Smidt & de Vos2004; Löffler & Edwards 2006). Under anaerobic con-ditions, which are common in aquifers contaminatedwith chlorinated aliphatic hydrocarbons, the mainmech-anism resulting in chlorinated ethene transformation tonontoxic ethene is reductive dechlorination (Maymo-Gatell et al. 1997). Thus far, Dehalococcoides sp. hasbeen identified as the only group of microorganismscapable of the complete dehalogenation of the chlori-nated ethenes to nontoxic ethene. Avariety of anaerobicdechlorinating Dehalococcoides sp. has been isolatedand their reductive dehalogenases (RDase) have beenidentified. In field applications, identification and char-acterization of key organisms and their functional genesthat catalyze reductive dechlorination (i.e., RDase) arecrucial to assess their relevance and implement naturalattenuation strategies. Various studies targeting the 16SrRNA genes have shown that Dehalococcoides sp. isindigenous to many sites contaminated with chlorinatedethenes worldwide (Major et al. 2002; Lendvay et al.2003; Löffler & Edwards 2006; Tas et al. 2009), and thattheir occurrence often correlates with in situ productionof ethene from dechlorination of higher chlorinatedethenes (He et al. 2003, 2005; Hendrickson et al.2002). However, the presence and diversity of 16SrRNA gene of Dehalococcoides do not specificallyreflect the dechlorinating potential due to the highlydiverse content of RDase genes in 16S rRNA gene-identical strains (Duhamel et al. 2004; Lovley 2003).Little is known so far about the diversity of RDase genesunder field conditions or the development ofDehalococcoides and its RDase genes under selectivepressure which can be present during enrichment forexample.

The RDase enzymes catalyze the cleavage of chlorineatoms from chlorinated ethene molecules during the re-ductive dechlorination (Smidt & de Vos 2004; Habashet al. 2004). The tceA gene encoding a trichloroethene(TCE) RDase enzyme (Magnuson et al. 2000) is present inthe genomes of Dehalococcoides strains 195 and FL2.These strains can metabolically reduce TCE to vinyl chlo-ride (VC), and cometabolically reduce VC to ethene (Heet al. 2005; Tas et al. 2009). Dehalococcoides strains VS

and GT are also capable of chlorinated ethene reduction,which involves the vcrA gene encoding a VC RDase(Müller et al. 2004; Sung et al. 2006). In strain BAV1,the bvcA gene also encodes a VC RDase (Krajmalnik-Brown et al. 2004), which is involved in the dechlorinationof VC (He et al. 2005). These strains have been recentlyaffiliated with the newly described species D. mccartyi(Löffler et al. 2012).

Previous studies have demonstrated that dichloroethenes(DCE), VC, and ethene were present as microbial dechlo-rination products of tetrachloroethene (PCE) andTCE in theBitterfeld/Wolfen contaminated aquifer (Saxony-Anhalt,Germany; Nijenhuis et al. 2007; Imfeld et al. 2008,2011). Further, it was suggested that reductive dechlori-nation occurs in situ and correlates with the presence ofdechlorinating bacteria (i.e., members of the generaDehalococcoides, Desulfitobacterium, Dehalobacter,Desulfuromonas, and Geobacter) in different parts ofthe aquifer (Nijenhuis et al. 2007; Imfeld et al. 2008,2011).

Microcosms were set up using Bitterfeld groundwater,and Dehalococcoides-dominated cultures that metabolizechlorinated ethenes were enriched. Though capable ofPCE and TCE dechlorination to ethene, (Nijenhuis et al.2007; Cichocka et al. 2010), no tceA, vcrA, or bvcA genecould be identified (Kaufhold et al. 2012). In anotherstudy, reductive dechlorination of cis- and trans-DCE toethenewas characterized in a constructedwetland suppliedwith groundwater from the Bitterfeld aquifer (Imfeld et al.2010). The above-described dechlorinating systems (i.e.,groundwater, microcosms, enrichment cultures, andconstructed wetland) have the same source groundwater;however, they differ in terms of the prevailing environ-mental conditions, system heterogeneity, and complexity,as well as the stage of dechlorination activities. Thesedifferences may cause the selective enrichment of specificDehalococcoides populations with distinct dehalogenasegenes, which may impact in situ dechlorination activities.On one hand, one Dehalococcoides sp. (based on 16SrRNA gene sequence) may be present in all samples witheither an identical or varying set of RDase genes. On theother hand, multiple distinctDehalococcoides populationswith their respective RDase gene sets may be present.Alternatively, other, thus far unidentified, populationsmay play a role in the removal of lower chlorinatedethenes (DCEs and VC) in the different systems.

In this study, we addressed the following main ques-tions: (1) is the Dehalococcoides sp. enriched in labora-tory cultures as described previously representative of

1768, Page 2 of 12 Water Air Soil Pollut (2013) 224:1768

the Dehalococcoides diversity in the complete investi-gated field site; (2) how do environmental conditionsincluding enrichment conditions affect the diversity ofDehalococcoides and its key dehalogenase genes; and(3) are the vcrA and bvcAgenes both associatedwith thecomplete dehalogenation of chlorinated ethenes? There-fore, we compared the diversity of Dehalococcoides sp.and three key RDase genes (tceA, bvcA, and vcrA) ingroundwater, water from a constructed wetland, micro-cosms, and enrichment cultures established usingBitterfeld groundwater. The 16S rRNA gene sequencesof Dehalococcoides subgroups and the three RDase(tceA, bvcA, and vcrA) genes were sequenced andcompared between the systems. The presence and di-versity of Dehalococcoides sp. and RDase genes wereanalyzed and interpreted with respect to the varied con-ditions and activities in the investigated systems.

2 Materials and Methods

2.1 Description of the Bitterfeld/Wolfen Field Site

Groundwater, constructed wetland, and microcosm sam-ples were all derived from a plume of chlorinated ethenes,located in the industrial Bitterfeld/Wolfen contaminatedmega-site (Saxony-Anhalt, Germany; Fig. 1). The overallgeological setting and the hydrogeological characteristicsof the investigation site were previously described in detail(Heidrich et al. 2004; Wycisk et al. 2003). TheBitterfeld/Wolfen region (Saxony-Anhalt, Germany) com-prises a contaminated area of about 25 km2 with anestimated volume of 200 million m3 of contaminatedgroundwater as a result of the former chemical industry

(Wycisk et al. 2003). Chlorinated aliphatic compounds arethe largest and most relevant group of contaminants in theinvestigated area, although other contaminants, such aschlorinated aromatics, hexachlorocyclohexane, or BTEX,are also present. cis-DCE is a major contaminant whichaccumulates as a result of PCE and TCE dechlorination.

2.2 Experimental Setups and Sampling Procedure

2.2.1 Groundwater and Constructed Wetland

Location of the wells within the cis-DCE plume isprovided in Fig. 1. Groundwater samples from wells1241, 3051, 3062, ML-d7, ML-d4, and BMH werecollected using a submersible peristaltic pump (MP1Grundfos, Bjerringbro, Denmark). Dissolved oxygen,pH, conductivity, redox potential, and temperaturewere measured directly at the field site. To ensurerepresentative sampling, groundwater samples werecollected after the sampling tubes were purged to re-place at least the equivalent of one volume of ground-water and until values of field measurements varied byless than 5 % (Imfeld et al. 2008).

The constructedwetlandwas continuously suppliedwithanoxic groundwater from well BMH, in which cis- andtrans-1,2-DCEwere themain contaminants (Table S1). Thecharacteristics of the constructed wetland have been de-scribed previously (Imfeld et al. 2010).Water samples werecollected from the sand compartment of the wetland usingsterile glass syringes 430 days after the beginning of theexperiment, when anaerobic conditions prevailed in thewetland (Imfeld et al. 2010).

Groundwater and wetland samples were dispensedinto vials that were sealed with Teflon-coated septa

Fig. 1 Schematics of thecis-DCE plume at the con-taminated site (Bitterfeld/Wolfen, Germany) with thelocation of the wells (▲).The concentration at the iso-lines is given in μg l-1. Thebold black arrow indicatesthe overall direction ofgroundwater flow

Water Air Soil Pollut (2013) 224:1768 Page 3 of 12, 1768

(headspace-free) for concentration analysis of volatileorganic compound (VOC), 20 ml high-density poly-ethylene bottles for geochemical analysis, and replicate1 l autoclaved glass bottles (groundwater samples) or120 ml vials (wetland samples) for microbial investi-gations as described previously (Imfeld et al. 2008,2010). The groundwater and wetland samples wereplaced on ice and transported directly to the laboratoryfor chemical analysis. The samples intended for micro-bial analysis were immediately cooled to 4 °C to slowdown further transformation processes and filteredwithin less than 5 h after sampling as described previ-ously (Nijenhuis et al. 2007; Imfeld et al. 2010).

2.2.2 Microcosms and Enrichment Cultures

Microcosms and enrichment cultures were set up usinggroundwater from wells 3051, 588/589, and 5261.Preparation of the microcosms was previously de-scribed in Nijenhuis et al. (2007). Briefly, laboratorymicrocosms were prepared from three differentsources. One set was prepared with groundwater orig-inating from well 3051 that was located at the fringe ofthe PCE and TCE plume but contained high concen-trations of DCE. The second set was prepared from thegroundwater of wells 588/589 with only low chlorinat-ed ethene concentrations (Table S1). The third set wasprepared from the groundwater of well 5261 with VC,cis- and trans-DCE as main contaminants. Microcosmswere amended with an electron donor (lactate or ace-tate) and an electron acceptor (VC, PCE, or trans-DCE; see Table 1).

The enrichment cultures were derived frommicrocosmsprepared from well 3051 as described in Nijenhuis et al.(2007) and further cultivated as described byCichocka et al.(2010; see also Table 1). Briefly, microcosmswere preparedin 120 ml bottles filled with 100 ml of groundwater, closedwith Teflon-coated butyl rubber septa and crimped. Lactate(3mM)was used as electron donor and PCE (100μmol l-1)as electron acceptor. Active microcosms were transferredthree times into mineral medium described by Zinder(1998), and amended with PCE as electron acceptor andlactate as electron donor and carbon source (Cichocka et al.2010). The third transfer was used as inoculum for theenrichment cultures. This study investigated a second cul-ture transfer on VC/lactate, a second culture transfer onVC/H2/acetate, and a third culture transfer withPCE/H2/acetate. Other characteristics of these enrich-ment cultures were also described by Cichocka et al.

(2010), without functional gene analysis. Enrichmentcultures were incubated at 20 °C without shaking. Sam-ples were taken using sterile syringes. Bacterial biomassfor further DNA extraction was obtained from the liquidculture of both microcosms and enrichment cultures bycentrifugation of 1.5 ml at 16,100×g for 30 min.

2.3 Analytical Methods

Quantification of chlorinated ethenes and ethene wasperformed with a gas chromatograph (Varian ChrompackCP-3800,Middelburg, The Netherlands) with flame ioniza-tion detection (GC-FID) equipped with a 30 m×0.53 mmGS-Q column (J&W Scientific, Waldbronn, Germany) asdescribed by Nijenhuis et al. (2007).

2.4 Molecular Analysis

2.4.1 DNA Extraction

DNAwas extracted from the membrane filters (from thegroundwater 2×1 l, and from the wetland a 120 mlsample was filtered) and the microcosm samples bydisrupting microorganisms with a bead beater(FastPrep®, Qbiogene, Irvine, CA, USA), applying aFastDNA® kit for DNA extraction (BIO101, La Jolla,CA, USA) and elution in 50 μl nuclease-free water.DNA concentration and quality were assessed spectro-photometrically (Nanodrop ND 1000; NanoDrop Tech-nologies, DE), and DNA samples were normalized bythe lowest concentration value (1.5 ng μl-1; diluting withnuclease-free water). DNA samples were tested for PCRamplification with universal 16S rRNA gene primers 27 F(5′AGAGTTTGATCMTGGCTCAG 3′; Lane 1991) and1387R (5′ GGGCGGWGTGTACAAGGC 3′; Heueret al. 1997) using the following amplification program:95 °C (15 min), followed by 30 cycles of 95 °C (30 s),52 °C (30 s), and 72 °C (50 s), completed with anadditional 30 min at 72 °C.

2.4.2 Taxon-Specif ic PCR Amplificationand Calibration Processes

Taxon-specific 16S rRNA-based PCR amplification wasused to test the presence of the genera Dehalococcoidesand performed according to Imfeld et al. (2008). Theforward primer DHC1 (5′ GATGAACGCTAGCGGCG3′) and the reverse primer DHC1377 (5′ GGTTGGCACATCGACTTCAA 3′) were used in the first round,

1768, Page 4 of 12 Water Air Soil Pollut (2013) 224:1768

Tab

le1

Sum

maryof

samples,h

ydrochem

istry,contam

inantsanddetectionof

specificDehalococcoides

andfunctio

nalgenesPCRanalysis

Origin(see

Fig.1)

Hyd

rochem

istry

Chlorinated

ethene

(presence)

DHCdehalogenases

Closestdehalogenase

relativ

e

Eh

(mV)

pHTOC

(mg

l-1)

PT

DV

E(p/

c)tceA

bvcA

vcrA

bvcA

(bp)

vcrA

(bp)

Groun

dwater

1241

(plumesource)

−123

4.8

–X

XX

XX

Pn.d.

n.d.

X–

Dehalococcoides

sp.G

T(100

%)CP00

1924

(144

)

3051

(middleof

theplum

e)−2

726.4

–a

XX

XX

Pn.d.

n.d.

X–

Dehalococcoides

sp.G

T(100

%)CP00

1924

(375

)

3062

(fring

eof

theplum

e)−4

126.7

–X

XX

XX

Pn.d.

n.d.

n.d.

–

ML-d7(m

ultilevelwell,36

.5mbS

)11

5.8

<5

XX

XX

XP

n.d.

n.d.

X–

Dehalococcoides

sp.G

T(100

%)CP00

1924

(149

)

ML-d4(m

ultilevelwell,19

.5mbS

)30

5.7

<5

aX

Xa

aP

n.d.

n.d.

n.d.

–

BMH

327

6.7

15.6

XX

XX

XP

n.d.

n.d.

n.d

–

Con

structed

wetland

BMH-W

L(w

ater

samples

from

awetland

system

treatin

gcis-

andtran

s-DCE)

−137

6.9

35.9

aa

XX

XP

n.d.

XX

Dehalococcoides

sp.B

AV1

(100

%)CP00

0688

(460

)Dehalococcoides

sp.G

T(100

%)CP00

1924

(340

)

Microcosm

s58

8/58

9-1(from

grou

ndwater

+3

mM

lactateandVC)

––

–a

aa

XX

Pn.d.

XX

Dehalococcoides

sp.B

AV1

(98%)CP00

0688

(721

)Dehalococcoides

sp.G

T(100

%)CP00

1924

(390

)

588/58

9-2(from

grou

ndwater

+3mM

lactateandtDCE)

––

–a

aX

XX

pn.d.

n.d.

XDehalococcoides

sp.G

T(100

%)CP00

1924

(350

)

5261

(from

grou

ndwater

+3mM

lactateandVC)

––

–a

aa

XX

Pn.d.

n.d.

XDehalococcoides

sp.G

T(100

%)CP00

1924

(377

)

3051

-1(from

grou

ndwater

+3mM

lactateandVC)

––

–a

aa

XX

Pn.d.

XX

Dehalococcoides

sp.B

AV1

(97%)CP00

0688

(422

)Dehalococcoides

sp.G

T(100

%)CP00

1924

(377

)

3051

-2(enrichedDHCcultu

re+H2+acetate/lactateandVC/PCE)

––

–(x)

(x)

(x)

XX

Pn.d.

n.d.

XDehalococcoides

sp.G

T(100

%)CP00

1924

(376

)

Reference

strains

Dehalococcoides

etheno

genes19

5Magnu

sonetal.2

000

Xn.d.

n.d.

Dehalococcoides

sp.F

L2

Heetal.2

005

Xn.d.

n.d.

Dehalococcoides

sp.B

AV1

Krajm

alnik-Brownetal.2

004

n.d.

Xn.d.

Dehalococcoides

sp.G

TSun

getal.2

006

n.d.

n.d.

X

Dehalococcoides

sp.V

SMülleretal.2

004

n.d.

n.d.

X

TOCtotalorganiccarbon

,PPCE,T

TCE,D

DCE,V

VC,E

ethene,D

HCDehalococcoides

sp.,pPinellas,cCornellsubg

roup

,Xpresence,a

absence,n.d.genescouldno

tbedetected,–

respectiv

edataisno

tavailableor

notrelevant,(x)

someenrichmentcultu

resPCEwereaddedandTCE,D

CEwereob

served

astransformationprod

ucts.

Water Air Soil Pollut (2013) 224:1768 Page 5 of 12, 1768

while DHC774 (5′ GGGAGTATCGACCCTCTC 3′)coupled with the reverse primer DHC1212 (5′GGATTAGCTCCAGTTCACACTG 3′) were used inthe second round, in a nested PCR approach (Hendricksonet al. 2002). The PCR products were purified using aQIAquick PCR purification kit (Qiagen, Hilden, Germany)and quantified using aNanoDropND1000 device. Follow-ing the nested PCR, the amplicons were sequenced directlywithout cloning with the primer DHC1212. Big Dye Ter-minator Cycle Sequencing Ready Reaction kit V3.1 (Ap-plied Biosystems, Foster City, CA USA) was applied todetermine the nucleotide sequences. The sequencing reac-tions were performed according to the recommendations ofthe manufacturer. Sequencing products were separated onABI PRISM 3130xl Genetic Analyzer (AppliedBiosystems). Analysis of the sequences was performed byMEGA 5.05 software (Tamura et al. 2011). Similaritysearch of the related sequences was performed using thebasic local alignment search tool program on the NationalCenter for Biotechnology Information (Altschul et al.1997). All obtained sequences were unambiguous.

Based on specific base substitution patterns in the V2and V6 regions of the 16S rRNA sequence, theDehalococcoides genus can be divided into three sub-groups: Cornell, Victoria, and Pinellas (Hendrickson et al.2002). In order to assess the diversity of Dehalococcoidesof the Cornell and Pinellas subgroups by direct sequencingin the different samples, the detection limit was determinedwith templates of different mixtures of the two phylogeneticsubgroups, i.e., D. mccartyi strain 195 and derived fromsample 588/589. The Victoria subgroup was not examinedin this study, since occurrence in Europe has not beenreported yet (Hendrickson et al. 2002). Quantifiedamplicons of Dehalococcoides-specific PCR for thePinellas and Cornell subgroups were mixed in ratios of1:1, 1:5, 1:10, 1:50, 1:100, 5:1, 10:1 50:1, and 100:1. Themixtures of purified amplicons were directly sequencedwith the primer DHC1212 and the detection limits for eachsubgroup present togetherwere established. The sequencingreactions and the analyses of the sequences were performedas described above.

2.4.3 Analysis of RDase genes

Three RDase genes—vcrA, bvcA, and tceA—werePCR amplified and sequenced as described above.For the amplification of vcrA, gene primers vcrAf (5′TGCTGGTGGCGTTGGTGCTCT 3′) and vcrAr (5′TGCCCGTCAAAAGTGGTAAAG 3′; Müller et al.

2004) were used; for bvcA, gene primers bvcAF (5′TGCCTCAAGTACAGGTGGT 3′) and bvcAR (5′ATTGTGGAGGACCTACCT 3′; Krajmalnik-Brownet al. 2004) were used; and for tceA, gene primers 797f(5′ ACGCCAAAGTGCGAAAAGC 3′) and 2490r (5′TAATCTATTCCATCCTTTCTC 3′; Magnuson et al.2000) were used. PCR parameters were 15 min at95 °C, 30 cycles of 45 s at 94 °C, 1 min at 55 °C (vcrA)or 52 °C (bvcA and tceA), 1 min at 72 °C, followed by7 min at 72 °C. PCR products were sequenced withreverse primers as mentioned previously. Plasmids con-taining tceA (from Dehalococcoides sp. FL2), vcrA(from strain GT), and bvcA (from strain BAV1) geneswere used as positive control DNA in PCR. A similaritysearch of the related sequences was performed using theBLAST program on the NCBI (Altschul et al. 1997).Phylogenetic analysis was conducted using the MEGA5.05 software (Tamura et al. 2011).

3 Results and Discussion

Knowledge on the presence and diversity of Dehalo-coccoides sp. and related RDase genes is essential forthe evaluation of the potential for complete removal ofchlorinated ethenes in situ since Dehalococcoides isthe only genus described thus far dehalogenating thechlorinated ethenes past DCE to ethene. We evaluatedthe presence and sequence diversity ofDehalococcoidessp. and three RDase genes in different types of systems(i.e., groundwater, constructed wetland, microcosms,and enrichment cultures), all of them derived from theBitterfeld/Wolfen contaminated site. In all the analyzedsystems, ethene was present as an end product of chlo-rinated ethene dehalogenation. In particular, we evalu-ated variations of both the 16S rRNA and RDase genes,revealing distinct groups produced by selective pres-sures exerted by the specific eco-chemical conditionsthat prevailed in the systems studied.

3.1 16S rRNA Dehalococcoides sp. Genes

Taxon-specific 16S rRNA-based PCR amplification wascarried out to detect the presence of the generaDehalococcoides. The nested PCR amplification yielded438 bp amplicons in all cases, which emphasizes thatDehalococcoides sp. was present in all samples (Table 1).

Since a previous study showed that rare 1–2 bpvariations of Dehalococcoides type sequences were

1768, Page 6 of 12 Water Air Soil Pollut (2013) 224:1768

introduced by the cloning, re-amplification, and se-quencing approach (Cichocka et al. 2010), the diversi-ty of Dehalococcoides sp. was assessed by a noveldirect sequencing approach. With this method, wewere able to differentiate between bias caused by theapplied molecular biological method and true sequencevariants and also for rapid and definitive identificationof the subgroup present. The direct sequencing proce-dure was tested by mixing defined PCR products atvarious ratios. The DNA ratio that enables distinctionof the Cornell and Pinellas Dehalococcoides sub-groups ranged between 1:5 and 1:50 (Fig. 2.). At ratio1:5, both minor and predominant sequences could bedistinguished, whereas at ratio 1:50, minor peaks couldbe still observed, although the peak height was in therange of noise level. Based on this method, all se-quences, either from groundwater, wetland, micro-cosm, or enrichment culture samples, belonged to thePinellas subgroup.

The phylogenetic analysis revealed that the retrievedsequences were identical (100 % sequence similarity) tothose of Dehalococcoides strains GT, FL2, BAV1, andCBDB1 (Table 1). Since all sequences were identical,only the longest sequence was submitted to the EMBLdatabase under accession number HE651153. This se-quence was identical to those previously retrieved fromthe Bitterfeld groundwater (AM399022) and enrich-ment culture BTF08, which was prepared with the samegroundwater (AM981291). These observations suggestthat—based on 16S rRNA gene sequence—only oneribotype of Dehalococcoides sp. was present in theBitterfeld area and that the organisms prevailing in thesource samples were maintained in subsequent enrich-ments, irrespective of the different chemical conditions(contamination, geochemical parameters like Eh, TOC,pH, and electron donor).

Our results also highlight that taxon-specific or gene-specific PCR by direct sequencing may provide suffi-cient information if a given taxon or a specific genevariant predominates with minor populations present at≤2 %. Nevertheless, ambiguous sequences in somecases suggest that a mixed template requires molecularcloning in order to correctly affiliate the microorganismsor genes represented by the mixed sequences.

While the specific 16S rRNA targeted PCR andsubsequent sequencing confirmed the presence ofDehalococcoides-related sequences, they do not pro-vide much information about the dechlorination activ-ity and substrate specificity. Therefore, we also

examined the presence of specific RDase genes inorder to better evaluate and predict the catabolic po-tential of the Dehalococcoides population.

3.2 RDase Genes

The presence and phylogeny of TCE RDase (tceA) andVC RDase (vcrA, bvcA) genes was specifically exam-ined because reductive dechlorination to ethene oc-curred in the investigated systems. Sequences weresubmitted to the EMBL database under accession num-bers HE653967 to HE653970.

3.2.1 tceA Gene

PCR assays targeting the tceA gene resulted in no de-tectable amplicons, although the co-occurrence of highTCE concentrations (11.6–35 mg l-1) with dechlorina-tion products (including DCE, VC, and ethene) wasobserved in groundwater samples from wells 1241(15.9 mg l-1 c-DCE, 4 mg l-1 VC and 0.5 mg l-1 ethene)and 3062 (0.3mg l-1 c-DCE, 0.2 mg l-1 VC and 0.1 mg l-1 ethene; Table S1). Also, the enrichment culture wasshown to dechlorinate TCE (Cichocka et al. 2010). Theapparent absence of this gene would suggest that reduc-tive dehalogenation of TCE in cultures 1241, 3062,3051–2 is performed by microorganisms with differentTCE RDase gene sequences (i.e., Desulfuromonas,Desulfitobacterium, and Dehalobacter), as observedpreviously at the field site (Nijenhuis et al. 2007; Imfeldet al. 2008, 2011). These microorganisms may partiallyor fully contribute to dehalogenation of TCE to cis-DCEin the above-mentioned groundwater and microcosmsamples. Another explanation could be that a varianttceA gene might have been expressed that could not beamplified with the set of primers used in the presentstudy. Previous studies showed that the same primersfailed to detect tceA in TCE-contaminated samples andin TCE-degrading mixed cultures (Waller et al. 2005;Morris et al. 2007) and another study highlighted thatonly part of the tceA genetic diversity might be retrievedwith these primers (Krajmalnik-Brown et al. 2007). In arecent study, TCE and VC RDase genes could not bedetected by assessing the diversity of the putative RDasegenes using highly degenerated primers in enrichmentcultures from Bitterfeld, although dechlorination wasestablished (Kaufhold et al. 2012). However, recentgenome sequencing of the Dehalococcoides mcccartyistrain BTF08 (CP004080) enriched from the same

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contaminated site revealed the presence of the tceA geneand the primers’ target sequences (Pöritz et al. 2013).This suggests that theDehalococcoides populations in oursamples also contained the target gene, but the PCR de-tection failed. A homology search with partial sequences

of the primers revealed that the last 14 nucleotides (3′ end)of primer 2490r have an additional perfect match outsidethe target site and that the last 11 nucleotides of the sameprimer have one additional match on the other DNAstrand. In addition, the last nine bases (3′ end) of the

Fig. 2 Sequencing results of DNA mixtures of Dehalococcoides sp. belonging to the Pinellas and Cornell subgroups. 100 % Pinellas,100 % Cornell or 1:1, 1:5, 5:1, 1:50 and 50:1 mixtures

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797f primer match nine additional sites of the full genome.This partial annealing of the primers outside the targetsequence may initiate primer extension, and thereforereduce the efficiency, or could contribute to the failure ofthe PCR detection of the target gene. Therefore, absence oftceA determined by PCR has to be considered with cau-tion, as activity as well as the gene may be present;however, artefacts during the PCR amplification mayresult in a false negative.

3.2.2 vcrA and bvcA Genes

The vcrA gene was detected in groundwater samples(1241, 3051, and ML-d7), wetland samples, in all micro-cosm samples (588/589-1, 588/589-2, 5261, 3051–1), andin enrichment culture 3051–2. In all these samples, reduc-tive dechlorination to ethene occurred (43.3–526μg l-1), aspreviously described (Nijenhuis et al. 2007; Imfeld et al.2008, 2011; Table S1).

Direct sequencing of the PCR amplicons and furthersimilarity analysis resulted in unambiguous sequences thatwere identical to those of the vcrAgene ofDehalococcoidesstrain GT (Table 1) for which TCE, cis-DCE, and VCwerethe only growth-supporting electron acceptors identified(Sung et al. 2006). This is in line with our findings as wecould detect the vcrAgene in the systems studied where theelectron acceptor was TCE, cis-DCE, or VC.

The bvcA gene was detected where ethene productionwas observed in both the wetland (BMH-WL) treating cis-(1069±260 μg l-1) and trans-DCE (258±98.6 μg l-1) andthe microcosms 558/589-1 and 3051–1 which wereamended with VC. The bvcA genes were all closelyrelated to that of Dehalococcoides strain BAV1 (BMH-WLwith 100% similarity, 588/589-1 with 98% similarityand 3051–1 with 97 % similarity; Table 1). The presenceof the bvcA gene correlates with reductive dechlorinationactivity up to ethene in our samples. However, the bvcAgene could not be detected in the parallel microcosm588/589-2 with trans-DCE as the main electron acceptor(Table 1). This suggests that a different gene may beresponsible for dehalogenation of VC derived fromtrans-DCE. Moreover, our results show that both bvcAand vcrA genes can be present simultaneously, similar torecent observations by Dugat-Bony and co-workers(2012) during a field biostimulation study. However, itremains unclear whether the presence of two genes withina single genome or the co-detection of the two genesindicates the presence of two distinct Dehalococcoidespopulations.

Although no vcrA and bvcA genes were detected ingroundwater sample BMH with trans-DCE (284±36 μg l-1) and cis-DCE (1062±64 μg l-1) as the maincontaminants, their presence was detected in the corre-sponding wetland samples. In the wetland, VC (<5 to 518±13 μg l-1) and ethene (<5 to 102±5 μg l-1) were detectedafter 430 days of groundwater supply, which indicates theoccurrence of reductive dechlorination (Imfeld et al.2010). The occurrence of both vcrA and bvcA genesimply that both genes were involved in this process.

Our results showed that the groundwater, wetlandand microcosms samples (which were characterized bydifferent ecological pressures) contained the sameDehalococcoides mccartyi-related bacteria and dif-fered with respect to their RDase genes. This indicatesthe presence of various functional sub-clusters of thesame ribotype, similar to observations by Futagamiet al. (2011) and Ritalahti et al. (2006).

Only the vcrA gene was present when TCE or DCEwas the main contaminant. Similarly, Morris et al.(2007) found that only vcrAwas expressed in the KB-1 culture and suggested that, due to sequence variation,the bvcA gene sequences were not amplified using theapplied primers. This is also in agreement with previousresults, which emphasize that the VC RDase genes inKB-1 occurred according to the pattern of substrateutilization (Waller et al. 2005).

The bvcA gene showed a considerable variabilitybetween the samples whereas the vcrA gene was iden-tical in all samples (Table 1). This suggests that addi-tional sequence variants of the bvcA gene exist. Thesevariations may also affect the primer binding sites andresult in no PCR amplification with the used primer setunder the applied PCR conditions.

No RDase gene could be amplified in groundwatersamples 3062, ML-d4, and BMH. One possible expla-nation could be that the main contaminants in ML-d4and BMH samples were cis- and trans-DCE, with only asmall amount of VC and ethene, which suggests thatreductive dehalogenation of DCE and VC was occur-ring only at a very low rate. The lack of vcrA and bvcAgenes detected in those samples may be due to very lowinitial target-DNA concentrations, which resulted inamplicon concentrations below the detection limit, orthe presence of RDase sequence variants that could notbe amplified with the method used in this study. Alter-natively, other microorganism that can tolerate chlori-nated hydrocarbons or take part in their dehalogenation,e.g., Geobacter or Dehalobacter (Nijenhuis et al. 2007;

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Imfeld et al. 2008, 2010) might have been present in theanalyzed samples and their RDase genes are not ampli-fied with the applied primer sets and PCR conditions.

4 Conclusions

In this study, we investigated the ubiquity and diversityof Dehalococcoides sp. and its three RDase genes indifferent systems supplied with groundwater originat-ing from the same source in Bitterfeld/Wolfen.

Our study indicates that the enriched Dehalococcoidessp. from the field in Bitterfeld (Cichocka et al. 2010;Kaufhold et al. 2012) is relevant for the field site and ispresent in all investigated systems. Further, both key genesfor VC-reductive dehalogenation to ethene, vcrA, and bvcAwere found in parallel, suggesting that they are both impor-tant for the dehalogenation of chlorinated ethenes to ethene.Additionally, 16S rRNAgene diversity ofDehalococcoideswas not affected by the different environmental or enrich-ment conditions. However, RDase gene diversity wasconfirming that for monitoring purposes, key metabolicgene detection is of greater importance compared to thedetection of the ribosomal genes only for analysis of themetabolic potential. In the case of tceA, absence of a positivePCR amplification may not be relevant, as the same ormany other organisms with similar genes can take overthe metabolic function of TCE dehalogenation. Neverthe-less, tceAwas not found in our studied systems. However,this does not mean that TCE was not dechlorinated, sincethis process was observed in the microcosms; thus, it alsoconfirms that Dehalococcoides spp. with uniform 16S se-quences have different functionality.

Our investigations suggest that one species, evenone ribotype can be present, but behind this ribotypethere must be several genetically different inventories(different set of functional genes) and depending on theenvironmental conditions (especially halogenatedcompounds as electron acceptor) one or the other pop-ulation is getting predominant.

In practice, when the potential for chlorinated ethenedehalogenation to ethene is assessed, evaluating boththe presence and diversity of specific dehalogenasegenes is essential. Nevertheless, the results of this studyrepresent a snapshot view of the taxonomic and func-tional diversity in these systems and thus may not reflectthe entire evolution of the studied system with respect todechlorinating activity. Therefore, a regular monitoringof the geochemical parameters together with the

presence/absence of the taxon-specific and functionalgenes indicative of dechlorination activities might benecessary for a more comprehensive understanding ofmicrobial processes occurring in the various chlorinatedethene contaminated systems.

Acknowledgments É.Mészáros and G. Imfeld were supportedby a European Union Marie Curie Early Stage Training Fellow-ship (AXIOM, contract no. MEST-CT-2004-8332). This workwas supported by the Helmholtz Centre for Environmental Re-search – UFZ. We would like to thank D. Cichocka for gener-ously supplying the microcosm and enrichment culture samplesused and F.E. Löffler and K. Fletcher for providing thedehalogenase clones. The Department of Groundwater Remedi-ation and the SAFIRA II Project, in particular, H. Weiss and R.Trabitzsch, as well as J. Drangmeister and J. Grossmann ofGICON and the Landesanstalt für Altlastenfreistellung desLandes Sachsen-Anhalt – LAF, are acknowledged for supportduring sampling. We would also like to thank I. Mäusezahl andK. Ethner for help with laboratory work.

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