SEARCH

SEARCH BY CITATION

Keywords:

  • natural attenuation;
  • benzene;
  • anaerobic degradation;
  • syntrophy;
  • Desulfotomaculum subcluster Ih

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

The microbial communities of in situ reactor columns degrading benzene with sulfate as an electron acceptor were analyzed based on clone libraries and terminal restriction fragment length polymorphism fingerprinting of PCR-amplified 16S rRNA genes. The columns were filled with either lava granules or sand particles and percolated with groundwater from a benzene-contaminated aquifer. The predominant organisms colonizing the lava granules were related to Magnetobacterium sp., followed by a phylotype affiliated to the genera Cryptanaerobacter/Pelotomaculum and several Deltaproteobacteria. From the sand-filled columns, a stable benzene-degrading consortium was established in sand-filled laboratory microcosms under sulfate-reducing conditions. It was composed of Delta- and Epsilonproteobacteria, Clostridia, Chloroflexi, Actinobacteria and Bacteroidetes. The most prominent phylotype of the consortium was related to the genus Sulfurovum, followed by Desulfovibrio sp. and the Cryptanaerobacter/Pelotomaculum phylotype. The proportion of the latter was similar in both communities and significantly increased after repeated benzene-spiking. During cultivation on aromatic substrates other than benzene, the Cryptanaerobacter/Pelotomaculum phylotype was outcompeted by other community members. Hence, this organism appears to be specific for benzene as a growth substrate and might play a key role in benzene degradation in both communities. Based on the possible functions of the community members and thermodynamic calculations, a functional model for syntrophic benzene degradation under sulfate-reducing conditions is proposed.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Aromatic hydrocarbons such as benzene, toluene, ethylbenzene and xylenes (BTEX) make up a significant percentage of gasoline. They are toxic and, due to their relatively high water solubility and volatility, mobile in the saturated and vadose zone of an aquifer. Benzene is the most mobile and most toxic BTEX compound (Aksoy, 1985). All BTEX compounds are easily biodegraded under oxic conditions by ubiquitous bacteria (van Agteren et al., 1998). However, due to the low water solubility and rapid microbial consumption of oxygen, contaminant plumes become generally anoxic. Anaerobic benzene degradation was observed in laboratory enrichment cultures under methanogenic (Vogel & Grbic-Galic, 1987; Kazumi et al., 1997), nitrate-reducing (Nales et al., 1998; Burland & Edwards, 1999), iron-reducing (Lovley et al., 1996; Kazumi et al., 1997; Nales et al., 1998; Jahn et al., 2005; Botton & Parsons, 2006; Kunapuli et al., 2007) and sulfate-reducing conditions (Edwards & Grbic-Galic, 1992; Lovley et al., 1995; Phelps et al., 1996; Kazumi et al., 1997; Nales et al., 1998; Phelps & Young, 1999; Vogt et al., 2007; Musat & Widdel, 2008). Nevertheless, benzene is considered the most recalcitrant of all BTEX compounds, as the majority of laboratory and field studies failed to demonstrate anaerobic benzene degradation (Aronson & Howard, 1997; Johnson et al., 2003). Results from microcosm studies suggest that anaerobic benzene degraders are not ubiquitous in subsurface sediments (Kazumi et al., 1997; Nales et al., 1998; Weiner & Lovley, 1998; Phelps & Young, 1999). Recently, pure cultures of benzene-metabolizing, facultatively anaerobic nitrate reducers were isolated (Kasai et al., 2006). However, most sulfate-reducing or methanogenic benzene-degrading enrichment cultures were still phylogenetically diverse after cultivation for many years (Phelps et al., 1998; Ulrich & Edwards, 2003). This suggests that under strictly anoxic conditions, benzene is degraded by consortia rather than by single organisms. The biochemical pathway of anaerobic benzene degradation is currently not elucidated (Caldwell & Suflita, 2000; Ulrich et al., 2005).

We studied benzene degradation in an anoxic BTEX-contaminated aquifer polluted by a former hydrogenation and benzene production plant, where sulfate is the main electron acceptor. Previously, in situ benzene degradation was demonstrated and quantified at the site by compound-specific stable isotope fractionation analysis (Vieth et al., 2005; Fischer et al., 2007). Additionally, stable benzene-degrading enrichment cultures were established under sulfate-reducing conditions using material from in situ microcosms (Herrmann et al., 2008) or benzene-degrading columns as inoculum (Vogt et al., 2007). These columns were percolated with sulfidic groundwater from the site containing benzene as the main source of carbon and energy. The fill material of the columns consisted of sand particles or lava granules, two materials with distinctly different physico-chemical properties. In both sand- and lava-filled columns, benzene was mineralized with sulfate as an electron acceptor, as demonstrated by calculation of mass balances for benzene, sulfate and bicarbonate. The benzene degradation rate was on average twice as high in the sand-filled columns (up to 36 μM day−1) as compared with the lava-filled columns (up to 18 μM day−1) (Vogt et al., 2007).

Here, we investigated the phylogenetic composition and dynamics of the benzene-degrading bacterial communities colonizing both fill materials, in order to elucidate the structure and function of these communities. Microorganisms colonizing the lava granules from the columns were analyzed whereas the sand particles from the columns were used as the inoculum for laboratory microcosms and subsequently analyzed by cloning and sequencing of 16S rRNA genes. After shifting the microcosms on other substrates representing potential intermediates, alterations of the community composition were analyzed by community fingerprinting. Our study aimed at identification of key organisms involved in anaerobic benzene degradation and elucidation of their possible interactions.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Description of the field site

The contaminated aquifer examined is located on the site of a former coal hydrogenation and benzene production plant near Zeitz (Saxony-Anhalt, Germany). The main contaminant is benzene, which reached the aquifer by several leakages and accidents during the operation of the plant between 1960 and 1990. Benzene concentrations are as high as 13 mM in the source zone. At the site, an upper and a lower aquifer are separated by a lignite and a clay layer. Both aquifers are heterogeneous and hydrogeologically connected due to discontinuities of the lignite–clay layer. The aquifer matrix is composed of river gravel and sand sediments, which contain more than 95% quartz. Both aquifers are anoxic, and sulfate is the main electron acceptor (Vieth et al., 2005). Natural attenuation (NA) processes proceeding in the upper aquifer have been investigated intensively during the past couple of years (Vieth et al., 2005; Fischer et al., 2006, 2007; Schirmer et al., 2006; Stelzer et al., 2006; Alfreider & Vogt, 2007). Gödeke et al. (2006) showed by means of a reactive tracer test that toluene is oxidized with sulfate as an electron acceptor in the lower aquifer downstream of the source zone.

In order to investigate the anaerobic benzene bioremediation potential of the lower aquifer, two column systems consisting of four columns connected in series (25 cm diameter and 6 m length, each) were set up at an experimental plant in 2002, and continuously percolated with anoxic, sulfidic groundwater from the lower aquifer. One set of four columns was filled with sand and the other one with lava granules. The sand was taken from a nearby aerobic sand pit and sieved before use, resulting in a grain size between 2 and 3.15 mm. The lava is a volcanic rock from the Eifel under-saturated with silicon oxide. The percolating groundwater contains on average 300 μM benzene. Toluene, ethylbenzene and xylenes are present in trace amounts (≤1 μM). Furthermore, 4 mM sulfate, 300 μM sulfide, 120 μM ammonium, 5 μM orthophosphate, 150 μM potassium, 2.3 mM magnesium, 2.2 mM sodium and 6.1 mM calcium are present. A more detailed description of the column system was given by Vogt et al. (2007).

Cultivation and sampling of the benzene-degrading consortium ZzBs1-4

Microcosms were set up in 240-mL serum bottles (Glasgerätebau Ochs GmbH, Bovenden, Germany) inside a glove box (Coy Laboratory Products Inc.) containing an atmosphere of 95% nitrogen and 5% hydrogen. Each bottle was filled with 110 mL of sand particles taken from the sand-filled columns of the in situ reactor and filled with anoxic mineral salt medium described elsewhere (Vogt et al., 2007) to a total volume of c. 235 mL. Benzene (0.3 mM), phenol (0.3 mM), toluene (0.3 mM) or benzoate (1 mM) was added as sources of carbon and energy from stock solutions prepared in anoxic demineralized water. For each substrate two replicates were set up. Furthermore, two controls were prepared without a substrate. Sterile controls were set up using the same bottles and volumes of sand particles and medium. They were autoclaved three times at intervals of at least 24 h. Then the medium was exchanged by fresh medium and the substrates were added at the concentrations given above.

By default, growth was monitored by weekly determination of sulfide and substrate concentrations. Degradation rates were determined for the cultures used for terminal restriction fragment length polymorphism (T-RFLP) analyses. For this purpose, sulfide and substrate concentrations were measured twice a week. When a substrate was completely degraded, it was replenished at the concentrations mentioned above. When the sulfide concentration exceeded 4 mM, the medium was removed and exchanged by fresh medium. The microcosms were incubated statically at 22 °C in the dark outside of the glove box.

Chemical analyses

Sulfide was determined spectrophotometrically according to Cline (1969) with the following modifications: samples (25–200 μL) were dissolved in 1 mL zinc acetate dihydrate solution (20 g L−1) for fixing sulfide immediately after sampling. Subsequently, 4 mL distilled water and 400 μL N,N-dimethyl-p-phenylendiammoniumdichloride (DMPD) were added. After a 20-min reaction time absorption values were measured. Concentrations were calculated using standards prepared from an anoxic sulfide stock solution.

Benzene and toluene were analyzed by automated headspace GC using a Varian 3800 gas chromatograph (Varian, Palo Alto) equipped with a CP SIL 5 CB capillary column (film thickness, 0.12 μm; inside diameter, 0.25 mm; length, 25 m) and a flame ionization detector. The chromatographic conditions were as follows: injector temperature, 250 °C (split 1 : 50); detector temperature, 260 °C; and an oven temperature program consisting of 70 °C for 2 min, followed by an increase at a rate of 10 °C min−1 up to 90 °C and then followed by a further increase at a rate of 60 °C min−1 until 220 °C was reached. Helium (1 mL min−1) was used as a carrier gas. Liquid test samples (diluted 1 : 10 or 1 : 20 in 1.6 mM H2SO4; final volume, 10 mL) were prepared in 20-mL glass vials. The samples were incubated for 30 min at 70 °C in an agitator (rotation regime, 250 r.p.m. for 5 s and no rotation for 2 s) before analysis, and 1 mL of each sample's headspace was injected. For calibration, diluted standards of benzene and toluene prepared from stock solutions were treated in the same way as the samples. The stock solutions were prepared in pure methanol.

Phenol and benzoate were analyzed by HPLC (Shimadzu) using a UV/VIS detector. Separation was performed at 23 °C and a flow rate of 0.6 mL min−1 using a Nucleosil 100 C18 column (3 mm × 250 mm, 5 μm size; Knauer GmbH, Berlin, Germany) as the stationary phase and a buffer containing 60% (v/v) NaH2PO4 (131.5 mM, pH 2.8) and 40% (v/v) acetonitrile as the liquid phase. Samples were prepared by centrifugation (10 min, 18 900 g, 4 °C) and diluted 10-fold in distilled water. Twenty microliters of each sample were injected. The compounds were detected at 271 nm wavelength.

Thermodynamic calculations

Changes in Gibbs free energy inline image were calculated from the data of Thauer et al. (1977). ΔG0298 values for benzene and benzoate were taken from Kaiser & Hanselmann (1982).

DNA preparation, 16S rRNA gene cloning and sequencing

DNA was extracted according to Maher et al. (2001) directly from the sand and lava granules. Agarose gel electrophoresis and spectrophotometry were used for quality check and quantification. Bacterial 16S rRNA gene fragments were PCR-amplified and cloned as described elsewhere (Kleinsteuber et al., 2006) using the universal primers 27F and 1492R (Lane, 1991). Clone libraries were screened by amplified ribosomal DNA restriction analysis (ARDRA) with HaeIII and hierarchical cluster analysis of the ARDRA patterns as described by Kleinsteuber et al. (2006). Partial DNA sequencing of representative clones was performed using the BigDye RR Terminator AmpliTaq FS Kit 1.1 (Applied Biosystems, Weiterstadt, Germany) and the sequencing primers 27F and 519R (Lane, 1991). For almost complete sequencing of selected clones, sequencing primers 27F, 357F, 530F, 926F, 1114F, 519R, 1100R and 1492R (Lane, 1991) were used. Capillary electrophoresis and data collection were carried out on an ABI PRISM 3100 Genetic Analyzer (Applied Biosystems). Data were analyzed using abi prism dna sequencing analysis software, and 16S rRNA gene sequences were assembled by abi prism autoassembler software. The blastn tool (http://www.ncbi.nlm.nih.gov/BLAST) (Altschul et al., 1990) was used to search for similar sequences in the GenBank database, and the Seqmatch tool was used to search for similar sequences compiled by the Ribosomal Database Project – II Release 9.4 (http://rdp.cme.msu.edu) (Cole et al., 2005). The phylogenetic analysis was accomplished with arb software, version Linux Beta 030822 (http://arb-home.de) (Ludwig et al., 2004). The determined sequences were initially aligned to the SILVA SSURef database Release 90 (http://www.arb-silva.de) (Pruesse et al., 2007) by the arb Positional Tree Server and added to the SILVA tree using the Quick Add Parsimony tool and applying a Bacteria-specific filter. The alignment was verified by comparison with the next relative sequences and corrected manually. The final position within the arb tree and bootstrap values were calculated by the Parsimony Interactive tool.

The determined 16S rRNA gene sequences were deposited in the GenBank database under accession numbers EF613368EF613487.

T-RFLP analyses

Bacterial 16S rRNA gene fragments were PCR-amplified with the primers 27F-FAM (labeled at the 5′ end with phosphoramidite fluorochrome-5-carboxyfluorescein) and 1492R (Lane, 1991). Oligonucleotides were purchased from MWG Biotech (Ebersberg, Germany) or from biomers.net (Ulm, Germany). PCR was performed in 25 μL samples containing 3 μL of 1 : 100 diluted template DNA (equivalent to 1–2.5 ng), 5 pmol of each primer and 12.5 μL Taq Master Mix (Qiagen, Hilden, Germany). PCR cycle conditions were as decribed previously (Kleinsteuber et al., 2006). PCR products were purified using the Wizard® SV PCR Clean-Up System (Promega, Mannheim, Germany) and quantified after agarose gel electrophoresis and ethidium bromide staining using the genetools program (Syngene, Cambridge, UK). Purified PCR products were digested with the restriction endonucleases AluI, BstUI, HaeIII or RsaI, respectively (New England Biolabs, Schwalbach, Germany). A 10 μL reaction contained 2.5 ng DNA (for T-RFLP analyses of single clones) or 20 ng DNA (for T-RFLP analyses of the whole consortium) and 10 U of restriction enzyme. Samples were incubated at the appropriate temperature for 3 h and then precipitated with sodium acetate (pH 5.5) and ethanol. Dried DNA samples were resuspended in 20 μL HiDi formamide containing 1.5% (v/v) GeneScan-500 ROX standard (Applied Biosystems). Samples were denatured at 95 °C for 5 min and chilled on ice. The fragments were separated by capillary electrophoresis on an ABI PRISM 3100 Genetic Analyzer (Applied Biosystems). The lengths of the fluorescent terminal restriction fragments (T-RF) were determined using the genemapper V3.7 software (Applied Biosystems), and their relative peak areas were determined by dividing the individual T-RF area by the total area of peaks within the threshold of 50–500 bp. Only peaks with relative fluorescence intensities of at least 50 U were included in the analysis. Theoretical T-RF values of the dominant phylotypes represented in the clone library were calculated based on the determined partial 16S rRNA gene sequences using the NEBcutter V2.0 (http://tools.neb.com/NEBcutter2/index.php), and the calculated T-RF values were verified experimentally using the corresponding clones as templates. The relative T-RF abundances of representative phylotypes were determined based on the relative peak areas of the corresponding T-RF.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Bacterial community associated with lava granules

The lava granules were taken from a column in which benzene is mineralized with sulfate as an electron acceptor (Vogt et al., 2007). A 16S rRNA gene clone library of 814 clones was generated from DNA isolated from the lava granules and screened by ARDRA fingerprinting. Based on a hierarchical cluster analysis, 29 clones representing eight dominant operational taxonomic units (OTUs) were selected for partial sequencing. The numbers of clones representing the identified phylotypes (i.e. displaying identical ARDRA patterns) were recorded using a Phoretix database that has been set up from the ARDRA patterns. In total 31% of the clones were assigned. The phylogenetic composition of the consortium according to these data is shown in Fig. 1. The dominant phylotype comprising 18.2% of all clones was a member of the genus Magnetobacterium (phylum Nitrospira). The second-most prominent phylotype with a proportion of 6.1% (Fig. 1) was a clostridium belonging to the Desulfotomaculum cluster I of the Peptococcaceae, which also comprises the genera Cryptanaerobacter and Pelotomaculum. The phylogenetic position of the clostridial clones within the Desulfotomaculum cluster I is illustrated in Fig. 2. Deltaproteobacteria were found in low frequency, with only 2.3% of all clones belonging to the genus Desulfobacca and 1.7% belonging to the genus Syntrophus (Fig. 1). The phylogenetic relationship of these sequences with other Deltaproteobacteria is shown in Fig. 3. According to the phylogenetic analyses using the arb program, the other identified phylotypes comprising only minor proportions of the clone library were affiliated to the phyla Chloroflexi, Chlorobi and Fusobacteria (data not shown).

image

Figure 1.  Phylogenetic affiliation of the 16S rRNA gene sequences retrieved from the benzene-degrading consortium ZzBs1-4 and from the lava granulate taken from a benzene-degrading column.

Download figure to PowerPoint

image

Figure 2.  Phylogenetic relationship of clostridial clones retrieved from the benzene-degrading consortium ZzBs1-4 (bold, denominated as ZZ-S …) and from the lava granulate (bold, denominated as ZZ-L …) with the Cryptanaerobacter/Pelotomaculum group of Desulfotomaculum cluster I (Peptococcaceae). The determined partial 16S rRNA gene sequences were aligned to the SILVA SSURef database (release 90); the final positions within the tree and bootstrap values were calculated using the ARB Parsimony Interactive tool. Only bootstrap values above 50% are shown. Scale bar=10% nucleotide substitution.

Download figure to PowerPoint

image

Figure 3.  Phylogenetic relationship of deltaproteobacterial clones retrieved from the benzene-degrading consortium ZzBs1-4 (bold, denominated as ZZ-S …) and from the lava granulate (bold, denominated as ZZ-L …) with representatives of the Deltaproteobacteria. The determined partial 16S rRNA gene sequences were aligned to the SILVA SSURef database (release 90); the final position within the tree and bootstrap values were calculated using the ARB Parsimony Interactive tool. Only bootstrap values above 50% are shown. Scale bar=10% nucleotide substitution.

Download figure to PowerPoint

Bacterial community derived from sand microcosms

Several anaerobic benzene-degrading enrichment cultures (named ZzBs1-4) were established under sulfate-reducing conditions in laboratory microcosms using sand particles as inoculum taken from the benzene-degrading column system described elsewhere (Vogt et al., 2007). As shown in Fig. 4, 0.24 mM benzene was consumed within 1100 h (46 days) coupled with sulfide production. It was not possible to grow a sediment-free culture of ZzBs1-4; instead, the cells were attached to the sand particles as confirmed by microscopic observation. After the cells were detached by pyrophosphate or gentle mechanical treatment, the cultures no longer degraded benzene (data not shown).

image

Figure 4.  Conversion of aromatic substrates by the benzene-degrading sulfate-reducing consortium ZzBs1-4. The time course of substrate depletion and sulfide production during growth on benzene (a), toluene (b), phenol (c) and benzoate (d) is shown after cultivation for 120 (a, b) and 200 (c, d) days, respectively.

Download figure to PowerPoint

A 16S rRNA gene clone library of 1121 clones was generated from the consortium. Based on the cluster analysis of ARDRA patterns, 91 clones representing 36 OTUs were partially sequenced. The numbers of clones displaying identical ARDRA patterns were recorded with the phoretix program; as a result, about 51% of the clones were assigned. The phylogenetic composition of the consortium is shown in Fig. 1. In contrast to the clone library derived from the bacterial community associated with the lava granules, the most dominant phylotype of the sand consortium clone library with 14.7% of all clones was an Epsilonproteobacterium related to the genus Sulfurovum, followed by Desulfovibrio with 10.5%. The phylogenetic position of the Desulfovibrio sequences and other deltaproteobacterial sequences is shown in Fig. 3. The third prominent phylotype in the clone library with a proportion of 6.5% was the member of the Cryptanaerobacter/Pelotomaculum that had also been found on the lava granules (Fig. 2). The remaining phylotypes identified in the clone library of ZzBs1-4 were assigned to several Deltaproteobacteria (see Fig. 3) and to the Actinobacteria (Coriobacteriaceae, Rubrobacteraceae, Cellulomonadaceae), Chloroflexi, Bacteroidetes (Sphingobacteriaceae), Clostridia (Acetivibrio), Epsilonproteobacteria (Campylobacteraceae), Chlorobi, Caldithrix and the candidate phyla OP3, OP8, OP11 and WS1 (data not shown).

The phylogenetic composition of the consortium derived from the ARDRA patterns was verified by T-RFLP analysis applying three different restriction enzymes. As shown in Fig. 5, the T-RFLP analysis confirmed that the most dominant phylotype was the Sulfurovum-like organism, which was even under-represented in the clone library. The relative T-RF abundances of several Deltaproteobacteria were also higher than the percentages of these phylotypes according to the clone library. This bias might be explained by the use of different restriction enzymes and by the different phylogenetic resolution of ARDRA and T-RFLP analysis. As ARDRA relies on all restriction sites within the 16S rRNA gene amplicon, it also distinguishes highly similar but not identical sequence types, leading to an underestimation of phylogenetic groups with high micro-heterogeneity. Nevertheless, the proportion of the Cryptanaerobacter/Pelotomaculum phylotype was consistently determined to be 5% and 7% with both methods.

image

Figure 5.  Relative T-RF abundances of predominant phylotypes in the benzene-degrading consortium ZzBs1-4 according to partial sequencing of 16S rRNA genes and ARDRA fingerprinting (clone library) as well as T-RFLP analysis with the restriction enzymes AluI, BstUI and RsaI. The relative T-RF abundances of Desulfovibrio, Desulfobulbus, Desulfuromonas and Desulfobacca were summarized as the T-RF values of these phylotypes could not be distinguished by all enzymes.

Download figure to PowerPoint

Community dynamics of consortium ZzBs1-4 upon shifts to different aromatic substrates

Besides benzene, the consortium derived from sand columns degraded toluene, all xylene isomers, phenol, benzoate or acetate if added as single substrates, coupled with the production of sulfide (data not shown). Benzoate, phenol and toluene are putative intermediates of the anaerobic degradation pathway(s) of benzene. For this reason, we set up microcosms spiked repeatedly with either benzene, phenol, benzoate or toluene as the sole source of carbon and energy and monitored the developing community compositions. All compounds were degraded under sulfate-reducing conditions (Fig. 4). No sulfide was formed in the absence of substrates, and no substrate depletion was observed in the sterile controls (data not shown). The stoichiometry of oxidized substrate and produced sulfide indicated that phenol was mineralized. For benzene and benzoate as substrates, sulfide production was slightly lower than expected for complete substrate oxidation; for toluene, which had been added at the lowest concentration, no production of sulfide was observed. However, care must be taken when interpreting the stoichiometries of the oxidized substrate and sulfide produced in our microcosms, because sulfide can precipitate as FeS or Fe2S coatings on sand particles, as observed previously (Herrmann et al., 2008; Vogt et al., 2007). Benzene was degraded at a rate of 5.22 μM day−1. Benzoate and phenol were degraded considerably faster at rates of 33.68 and 17.5 μM day−1, respectively. Likewise, toluene was depleted faster than benzene (7.1 μM day−1).

After 120 and 270 days of cultivation on the different substrates, DNA was extracted from the sand material and used for T-RFLP analyses to assess changes in the phylogenetic composition of the consortium due to the degradation of the different carbon sources. As shown in Fig. 6, the Cryptanaerobacter/Pelotomaculum phylotype decreased in its relative T-RF abundance after cultivation on toluene, benzoate and phenol, and significantly increased when benzene was used as a substrate. The relative T-RF abundances of the Sulfurovum-like phylotype were also higher with benzene compared with the other substrates. Members of the Desulfobacteraceae became more dominant when benzoate, phenol and toluene were used as substrates. Furthermore, a larger proportion of other phylotypes (T-RF that were not assigned) was present in the cultures grown on benzoate, toluene or phenol, compared with the culture grown on benzene. It therefore appeared that the Cryptanaerobacter/Pelotomaculum- and Sulfurovum-like phylotypes were outcompeted by other groups when shifted from benzene to other aromatic substrates, whereas the Cryptanaerobacter/Pelotomaculum phylotype was further enriched after prolonged cultivation on benzene (270 days).

image

Figure 6.  Relative T-RF abundances of predominant phylotypes in the benzene-degrading consortium ZzBs1-4 after 120 and 270 days of cultivation on different aromatic substrates as determined by T-RFLP analysis with the restriction enzymes BstUI (a) and RsaI (b). Owing to accidental loss, the benzoate-grown culture could be analyzed only after 120 days. The relative T-RF abundances of Desulfovibrio, Desulfobulbus, Desulfuromonas and Desulfobacca were summarized as the T-RF values of these phylotypes could not be distinguished.

Download figure to PowerPoint

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Community composition and implications for syntrophic benzene degradation under sulfate-reducing conditions

Despite their common origin from a benzene-contaminated aquifer and their shared ability to degrade benzene with sulfate as an electron acceptor at comparable rates (Vogt et al., 2007), bacterial communities established on sand particles and lava granules differed remarkably. The only phylotype present in similarly substantial proportions on both materials (Figs 1 and 6) was a member of the Cryptanaerobacter/Pelotomaculum group. Sequences retrieved from both the lava granules and the sand consortium ZzBs1-4 are highly similar and form a distinct cluster within Desulfotomaculum cluster I (Fig. 2). The cultivated species of the genera Pelotomaculum and Cryptanaerobacter are syntrophic, heterotrophic bacteria, which live closely associated with hydrogen-consuming organisms (interspecies hydrogen transfer). Some of them were shown to degrade aromatic compounds like benzoate, phenol or phthalate (Qiu et al., 2003, 2006; Juteau et al., 2005). In contrast to members of the related genus Desulfotomaculum, Pelotomaculum and Cryptanaerobacter species do not use sulfate as an electron acceptor. It has been hypothesized that these genera, also termed Desulfotomaculum subcluster Ih, have lost their ancestral ability of sulfate respiration as they adopted a syntrophic lifestyle to thrive in methanogenic environments (Imachi et al., 2006). Assuming that the Cryptanaerobacter/Pelotomaculum phylotype detected in the lava- and sand-associated communities has conserved the ability to reduce sulfate, it might be able to oxidize benzene completely. Although we cannot rule out this possibility altogether, it appears rather unlikely considering the communities' composition. As explained below, we suggest that this Pelotomaculum/Cryptanaerobacter-like phylotype represents a syntrophic organism responsible for the initiation of the observed anaerobic benzene degradation.

The dominant phylotype in the clone library generated from the lava granules was a member of the Nitrospira phylum related to the genus Magnetobacterium. Because most relatives described hitherto are environmental clones that have not been cultivated so far, the possible physiological function of the Magnetobacterium-like phylotype in the lava community remains unknown. The closest relatives are environmental clones retrieved from sulfate-containing wetlands (Acc. nos. DQ137961, DQ137982), a forested wetland impacted by reject coal (Brofft et al., 2002) and a PCB-contaminated salt marsh sediment (Acc. no. AF286037). The next cultured relatives are sulfur reducers of the genus Thermodesulfovibrio. Candidatus ‘Magnetobacterium bavaricum’ has been suggested to be a chemolithoautotroph with an iron-dependent mode of energy conservation (Spring et al., 1993). Recently, in a stable isotope probing experiment with 13C-labeled acetate, Magnetobacterium-like organisms were found to metabolize acetate in a methanogenic sediment (Schwarz et al., 2007).

The physiological role of the Sulfurovum-like phylotype, which dominated the sand-derived microcosms, is not clear, either. As for the Magnetobacterium group, most sequences found for Sulfurovum relatives stem from environmental clones and not from cultivated species. Members of this genus were often found in sulfidic habitats (Campbell et al., 2006). A highly similar phylotype was retrieved from a phenol-degrading, methanogenic bioreactor (Zhang et al., 2005). Besides clones from various sulfidic habitats such as hydrothermal vents, springs and cave streams, a phylotype retrieved from a benzene-mineralizing consortium (Phelps et al., 1998) was also affiliated to this group.

In the clone library from the sand particles, sequences affiliated to several genera of sulfate-reducing Deltaproteobacteria were detected. For most of these phylotypes, related cultivated species have been described (Fig. 3). The most abundant Deltaproteobacterium belongs to the genus Desulfovibrio, which comprises physiologically versatile sulfate reducers, known for their capability to consume hydrogen very efficiently (Cord-Ruwisch et al., 1988; Bak & Pfennig, 1991). Hence, Desulfovibrio spp. are typical hydrogenotrophs in syntrophic associations under sulfate-reducing conditions (e.g. Elshahed et al., 2001; Jackson & McInerney, 2002). Also, Desulfobulbus relatives are generally very versatile and can grow on a wide range of substrates including hydrogen (e.g. Laanbroek et al., 1982; Lien et al., 1998). Desulfovibrio and Desulfobulbus species are not known for acetate oxidation (incomplete oxidizers). Desulfobacca-like sequences were detected on both sand particles and lava granules. The only described species, Desulfobacca acetoxidans, is specialized on the metabolization of acetate, and can even outcompete acetoclastic methanogens (Oude Elferink et al., 1999). Furthermore, the occurrence of several sequences related to Desulfuromonas acetexigens in the sand consortium indicates a function in acetate degradation, because D. acetexigens uses exclusively acetate as the sole source of carbon and energy (Finster et al., 1994). Several phylotypes of the sand consortium were related to Desulfobacterium and Desulfobacula spp. Species of these genera are known to degrade aromatics completely; hence, they are also able to oxidize acetate. Desulfobacteraceae are involved in the degradation of other aromatic substrates used by the sand consortium ZzBs1-4, as reflected by higher relative T-RF abundances after growth on toluene, phenol and benzoate (Fig. 6). As explained below, we suggest that the mentioned phylotypes affiliated to sulfate-reducing Deltaproteobacteria are mainly responsible for hydrogen and acetate oxidation coupled to sulfate reduction in the course of anaerobic benzene degradation.

Sequences affiliated to the genus Syntrophus were mainly detected in the lava-derived clone library. Syntrophus spp. have been shown to grow syntrophically on benzoate in coculture with hydrogenotrophic sulfate reducers or methanogens (Mountfort et al., 1984; Elshahed et al., 2001), but possibly they are also capable of syntrophic acetate oxidation as shown recently in microcosm experiments with 13C-labeled acetate (Chauhan & Ogram, 2006).

Phylogenetic diversity of the lava- and sand-associated consortia compared with other benzene-degrading consortia

Bacterial communities of sediment-free benzene-degrading enrichment cultures were examined by several authors. Phelps et al. (1998) and Musat & Widdel (2008) investigated cultures originally enriched from marine sediments under sulfate-reducing conditions. The consortium of Musat & Widdel (2008) consists only of Deltaproteobacteria, and the dominant phylotype is a Desulfobacterium species. The authors postulate that this organism is solely responsible for benzene mineralization in their culture. A clone closely related to this phylotype, named SB-21, had been described before by Phelps et al. (1998) in a sulfate-reducing, benzene-degrading consortium. Ulrich & Edwards (2003) investigated the community structure of a methanogenic consortium originally enriched from the sediment of a contaminated aquifer, which first used sulfate as an electron acceptor before switching to methanogenesis. A Desulfobacterium species and a member of the Peptococcaceae were the dominant phylotypes; however, it was not clear which of both species attacks benzene initially. Recently, Kunapuli et al. (2007) characterized the community structure of a benzene-degrading iron-reducing consortium by stable isotope probing. A member of the Peptococcaceae was shown to assimilate benzene primarily, but also a member of the Desulfobulbaceae and members of the Actinobacteria were prominent, indicating a syntrophic relationship during the course of benzene degradation.

Compared with the consortia described above, the sand- and lava-associated communities are more diverse and comprise only a small proportion of Desulfobacteraceae. In the consortium ZzBs1-4, the proportion of Desulfobacteraceae increased after the shift from benzene to toluene, phenol or benzoate (Fig. 6). In contrast to the consortia described above, ZzBs1-4 contains a large proportion of other Deltaproteobacteria like Desulfovibrio, which are not assumed to use aromatic substrates, and other bacteria not known as sulfate reducers. Similarly, in the lava-associated community only a small proportion of Deltaproteobacteria was detected. Except for the Cryptanaerobacter/Pelotomaculum phylotype, both communities are very different in their phylogenetic composition although they were inoculated with the same groundwater. These results form the basis for our hypothesis that benzene is not degraded by single sulfate reducers in the sand- and lava-associated communities, but by syntrophic interactions of several different organisms. We suggest that only one phylotype, originating from the groundwater percolating the column system, can initially attack benzene in our communities. If this phylotype could metabolize benzene completely, one would expect an aromatic-degrading organism using sulfate as an electron acceptor (likely a phylotype related to other typical aromatic-degrading sulfate reducers). Because no other organisms could grow under such circumstances in considerable amounts besides the benzene degrader, the respective phylotype should be dominant (≥50%) in community analyses, as observed for example by Musat & Widdel (2008)– but this is not the case in our consortia. Therefore, we postulate that a member of the Cryptanaerobacter/Pelotomaculum group is responsible for the initial attack of benzene in our enrichments. Interestingly, a member of the family Peptococcaceae dominated in the aquifer-derived methanogenic benzene-degrading enrichment culture examined by Ulrich & Edwards (2003). It might be capable of attacking benzene as well, as postulated recently for a member of the Peptococcaceae in a consortium growing on benzene under iron-reducing conditions (Kunapuli et al., 2007). Aquifers are rather oligotrophic environments and are characterized by conditions that favor slow-growing organisms with low energy demands. The capability to survive long starvation periods as well as an independence from specific electron acceptors might be favorable. Spore-forming fermenters degrading aromatic compounds in syntrophic association with methanogens, sulfate-reducers or iron-reducers might be indigenous in subsurface sediments and thus predestined to develop in benzene-polluted aquifers, at least in the sulfidic, methanogenic or iron-reducing zones.

Thermodynamic considerations and a functional model for syntrophic benzene degradation under sulfate-reducing conditions

The mineralization of benzene with sulfate as an electron acceptor is thermodynamically favorable as it allows synthesizing up to three ATP at 60 kJ mol−1 (Schink, 1997):

  • image(1)

As reasoned above, we assume that a syntrophic association rather than a single member of our communities mineralizes benzene. Support for the involvement of syntrophic associations in benzene degradation by the consortium ZzBs1-4 emerges from the observation that cell detachment and disaggregation at intensities that commonly do not lead to cell damage resulted in the loss of the communities' ability to degrade benzene.

The initial step of benzene degradation under anoxic conditions might be a methylation, a hydroxylation or a carboxylation reaction (Caldwell & Suflita, 2000; Ulrich et al., 2005). Benzoate is a very likely central intermediate in the anaerobic degradation pathway of benzene, because all known degradation pathways of aromatics under anoxic conditions are channeled into benzoate or benzoyl-coenzyme A (Widdel & Rabus, 2001; Boll et al., 2002). Benzoate has also been found in benzene-degrading consortia under iron-reducing (Botton & Parsons, 2007), nitrate-reducing, sulfate-reducing and methanogenic conditions (Caldwell & Suflita, 2000; Ulrich et al., 2005). However, the conversion of benzene to benzoate is thermodynamically unfavorable.

  • image(2)

Hence, a putative anaerobic benzene degrader cannot grow at the expense of reaction (2), but has to oxidize benzoate further to gain energy for ATP synthesis. We assume that acetate and hydrogen are central intermediates at least in the consortium ZzBs1-4, due to the presence of several phylotypes related to acetate- and hydrogen-consuming sulfate reducers, as discussed above. Therefore, we suggest benzene is oxidized according to the following equations:

  • image(3)
  • image(4)

The fermentation of benzene to acetate and hydrogen (reaction 3) is thermodynamically not feasible under standard conditions, but becomes exergonic in the case of low hydrogen and acetate concentrations, allowing growth of a putative benzene-fermenting organism. The energy demand of syntrophic bacteria can be very low: for Syntrophus species, inline image of −5 kJ mol−1 was observed as being sufficient for growth (Jackson & McInerney, 2002). Acetate and hydrogen are key intermediates during the anaerobic conversion of many organic substrates by the anaerobic food chain; both compounds are readily consumed by different types of anaerobes (Schink, 1997; Zengler et al., 1999).

Alternatively, benzene might be oxidized to acetate by a single sulfate reducer according to the following equation:

  • image(5)

However, in such a scenario, the dominance of Desulfovibrio sp. in the sand consortium cannot be explained. It should also be mentioned that all currently known aromatic-degrading sulfate-reducing bacteria are complete oxidizers, probably due to the high energy value of acetate.

Acetate might be oxidized by sulfate reducers related to Desulfobacca (sand, lava granules), Desulfuromonas (sand) and Desulfobacterium (sand), and possibly by other groups (Magnetobacterium- and Sulfurovum-like), according to the following equation:

  • image(6)

We assume that in our communities, the member of the Cryptanaerobacter/Pelotomaculum group ferments benzene to acetate according to equation (4), thus being responsible for the initial transformation reactions of the benzene degradation pathway. As shown for the sand-associated consortium ZzBs1-4 during growth on aromatic substrates other than benzene, the relative T-RF abundance of this phylotype decreased or fell below the detection limit (Fig. 6). By contrast, after prolonged cultivation on benzene (270 days) the relative T-RF abundance of Cryptanaerobacter/Pelotomaculum increased remarkably, indicating a further enrichment of this organism within the consortium (Fig. 6). Although limitations of T-RFLP analyses, e.g., differences in amplification efficiency or non-PCR-based biases, might impair the elucidation of real community structures, relative changes in community composition are reliably reflected by T-RFLP profiles (Hartmann & Widmer, 2008). A significant increase of the relative T-RF abundance of the Cryptanaerobacter/Pelotomaculum phylotype after cultivation on benzene was consistently detected with different restriction enzymes (Fig. 6) and with different amounts of template DNA in the range of 1–5 ng (data not shown). Moreover, in a toluene-grown culture of ZzBS1-4 that had been shifted back to benzene as the sole carbon source, neither benzene degradation nor sulfide production could be restored even after 90 days of incubation on benzene (data not shown), indicating outcompetition of the benzene-attacking key organism. Therefore, we conclude that this organism is responsible for the initiation of the anaerobic benzene degradation on both sand and lava granules.

Whereas we provide strong indications at the functional role of the Cryptanaerobacter/Pelotomaculum phylotype and the Deltaproteobacteria in the benzene-degrading communities, the physiological role of the Magnetobacterium- and Sulfurovum-related phylotypes and their possible involvement in syntrophic interactions still remain to be elucidated. These phylotypes were the predominant members of the communities colonizing either sand or lava granules, respectively. Obviously different properties of sand particles and lava granules gave rise to different community compositions. However, theoretically, both groups of organisms might occupy the same ecological niche in the two habitats. According to the hypothesized degradation pathway and regarding the high relative T-RF abundance of the Sulfurovum-like phylotype, they might also act as hydrogen consumers or acetate oxidizers.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

This work is integrated in the internal research and development program of the UFZ as well as the SAFIRA project. The authors thank Jörg Ahlheim, Ralf Trabitzsch and Werner Kletzander of the Department of Groundwater Remediation for help regarding sampling of sand and lava material from the columns. Special thanks are due to Stephanie Hinke and Ute Lohse for excellent technical assistance.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information
  • Aksoy M (1985) Benzene as a leukemogenic and carcinogenic agent. Am J Ind Med 8: 920.
  • Alfreider A & Vogt C (2007) Bacterial diversity and aerobic biodegradation potential in a BTEX contaminated aquifer. Water Air Soil Pollut 183: 415426.
  • Altschul SF, Gish W, Miller W, Myers EW & Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215: 403410.
  • Aronson D & Howard PH (1997) Anaerobic biodegradation of organic chemicals in groundwater: A summary of field and laboratory studies. Final Report. Environmental Science Center.
  • Bak F & Pfennig N (1991) Sulfate-reducing bacteria in littoral sediment of Lake Constance. FEMS Microbiol Ecol 85: 4352.
  • Boll M, Fuchs G & Heider J (2002) Anaerobic oxidation of aromatic compounds and hydrocarbons. Curr Opin Chem Biol 6: 604611.
  • Botton S & Parsons JR (2006) Degradation of BTEX compounds under iron-reducing conditions in contaminated aquifer microcosms. Environ Toxicol Chem 25: 26302638.
  • Botton S & Parsons JR (2007) Degradation of BTX by dissimilatory iron-reducing cultures. Biodegradation 18: 371381.
  • Brofft JE, McArthur JV & Shimkets LJ (2002) Recovery of novel bacterial diversity from a forested wetland impacted by reject coal. Environ Microbiol 4: 764769.
  • Burland SM & Edwards EA (1999) Benzene biodegradation linked to nitrate reduction. Appl Environ Microbiol 65: 529533.
  • Caldwell ME & Suflita JM (2000) Detection of phenol and benzoate as intermediates of anaerobic benzene biodegradation under different terminal electron-accepting conditions. Environ Sci Technol 34: 12161220.
  • Campbell BJ, Engel AS, Porter ML & Takai K (2006) The versatile ɛ-proteobacteria: key players in sulphidic habitats. Nature Rev Microbiol 4: 458468.
  • Chauhan A & Ogram A (2006) Phylogeny of acetate-utilizing microorganisms in soils along a nutrient gradient in the Florida Everglades. Appl Environ Microbiol 72: 68376840.
  • Cline JD (1969) Spectrophotometric determination of hydrogen sulfide in natural waters. Limnol Oceanogr 14: 454458.
  • Cole JR, Chai B, Farris RJ, Wang Q, Kulam SA, McGarrell DM, Garrity GM & Tiedje JM (2005) The Ribosomal Database Project (RDP-II): sequences and tools for high-throughput rRNA analysis. Nucleic Acids Res 33 (Database Issue): D294D296.
  • Cord-Ruwisch R, Seitz H-J & Conrad R (1988) The capacity of hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen depends on the redox potential of the terminal electron acceptor. Arch Microbiol 149: 350357.
  • Edwards EA & Grbic-Galic D (1992) Complete mineralization of benzene by aquifer microorganisms under strictly anaerobic conditions. Appl Environ Microbiol 58: 26632666.
  • Elshahed MS, Bhupathiraju VK, Wofford NQ, Nanny MA & McInerney MJ (2001) Metabolism of benzoate, cyclohex-1-ene carboxylate, and cyclohexane carboxylate by “Syntrophus aciditrophicus” strain SB in syntrophic association with H2-using microorganisms. Appl Environ Microbiol 67: 17281738.
  • Finster K, Bak F & Pfennig N (1994) Desulfuromonas acetexigens sp. nov., a dissimilatory sulfur-reducing eubacterium from anoxic freshwater sediments. Arch Microbiol 161: 328332.
  • Fischer A, Bauer J, Meckenstock RU, Stichler W, Griebler C, Maloszewski P, Kästner M & Richnow HH (2006) A multitracer test proving the reliability of Rayleigh equation-based approach for assessing biodegradation in a BTEX contaminated aquifer. Environ Sci Technol 40: 42454252.
  • Fischer A, Theuerkorn K, Stelzer N, Gehre M, Thullner M & Richnow HH (2007) Applicability of stable isotope fractionation analysis for the characterization of benzene biodegradation in a BTEX-contaminated aquifer. Environ Sci Technol 41: 36893696.
  • Gödeke S, Richnow HH, Weiß H, Fischer A, Vogt C, Borsdorf H & Schirmer M (2006) Multi component-reactive tracer test for the implementation of enhanced in-situ bioremediation at a BTEX-contaminated megasite. J Cont Hydrol 87: 211236.
  • Hartmann M & Widmer F (2008) Reliability for detecting composition and changes of microbial communities by T-RFLP genetic profiling. FEMS Microbiol Ecol 63: 249260.
  • Herrmann S, Kleinsteuber S, Neu T, Richnow HH & Vogt C (2008) Enrichment of anaerobic benzene degrading microorganisms by in situ microcosms. FEMS Microbiol Ecol 63: 94106.
  • Imachi H, Sekiguchi Y, Kamagata Y, Loy A, Qiu Y-L, Hugenholtz P, Kimura N, Wagner M, Ohashi A & Harada H (2006) Non-sulfate-reducing, syntrophic bacteria affiliated with Desulfotomaculum cluster I are widely distributed in methanogenic environments. Appl Environ Microbiol 72: 20802091.
  • Jackson BE & McInerney MJ (2002) Anaerobic microbial metabolism can proceed close to thermodynamic limits. Nature 415: 454456.
  • Jahn MK, Haderlein SB & Meckenstock RU (2005) Anaerobic degradation of benzene, toluene, ethylbenzene, and o-xylene in sediment-free iron-reducing enrichment cultures. Appl Environ Microbiol 71: 33553358.
  • Johnson SJ, Woolhouse KJ, Prommer H, Barry DA & Christofi N (2003) Contribution of anaerobic microbial activity to natural attenuation of benzene in groundwater. Engin Geol 70: 343349.
  • Juteau P, Côté V, Duckett M-F, Beaudet R, Lépine F, Villemur R & Bisaillon J-G (2005) Cryptanaerobacter phenolicus gen. nov., sp. nov., an anaerobe that transforms phenol into benzoate via 4-hydroxybenzoate. Int J Syst Evol Microbiol 55: 245250.
  • Kaiser J-P & Hanselmann KW (1982) Fermentative metabolism of substituted monoaromatic compounds by a bacterial community from anaerobic sediments. Arch Microbiol 133: 185194.
  • Kasai Y, Takahata Y, Manefield M & Watanabe K (2006) RNA-based stable isotope probing and isolation of anaerobic benzene-degrading bacteria from gasoline-contaminated groundwater. Appl Environ Microbiol 72: 35863592.
  • Kazumi J, Caldwell ME, Suflita JM, Lovley DR & Young LY (1997) Anaerobic degradation of benzene in diverse anoxic environments. Environ Sci Technol 31: 813818.
  • Kleinsteuber S, Riis V, Fetzer I, Harms H & Müller S (2006) Population dynamics within a microbial consortium during growth on diesel fuel in saline environments. Appl Environ Microbiol 72: 35313542.
  • Kunapuli U, Lueders T & Meckenstock RU (2007) The use of stable isotope probing to identify key iron-reducing microorganisms involved in anaerobic benzene degradation. ISME J 1: 643653.
  • Laanbroek HJ, Abee T & Voogd IL (1982) Alcohol conversions by Desulfobulbus propionicus Lindhorst in the presence and absence of sulfate and hydrogen. Arch Microbiol 133: 178184.
  • Lane DJ (1991) 16S/23S rRNA sequencing. Nucleic Acid Techniques in Bacterial Systematics (StackebrandtE & GoodfellowM, eds), pp. 115175. Wiley, Chichester.
  • Lien T, Madsen M, Steen IH & Gjerdevik K (1998) Desulfobulbus rhabdoformis sp. nov., a sulfate reducer from a water-oil separation system. Int J Syst Bacteriol 48: 469474.
  • Lovley DR, Coates JD, Woodward JC & Phillips EJP (1995) Benzene oxidation coupled to sulfate reduction. Appl Environ Microbiol 61: 953958.
  • Lovley DR, Woodward JC & Chapelle FH (1996) Rapid anaerobic benzene oxidation with a variety of chelated Fe(III) forms. Appl Environ Microbiol 62: 288291.
  • Ludwig W, Strunk O, Westram R et al. (2004) ARB: a software environment for sequence data. Nucleic Acids Res 32: 13631371.
  • Maher N, Dillon HK, Vermund SH & Unnasch TR (2001) Magnetic bead capture eliminates PCR inhibitors in samples collected from the airborne environment, permitting detection of Pneumocystis carinii DNA. Appl Environ Microbiol 67: 449452.
  • Mountfort DO, Brulla WJ, Krumholz LR & Bryant MP (1984) Syntrophus buswellii gen. nov., sp. nov.: a benzoate catabolizer from methanogenic ecosystems. Int J Syst Bacteriol 34: 216217.
  • Musat F & Widdel F (2008) Anaerobic degradation of benzene by a marine sulfate-reducing enrichment culture, and cell hybridization of the dominant phylotype. Environ Microbiol 10: 1019.
  • Nales M, Butler BJ & Edwards EA (1998) Anaerobic benzene biodegradation: a microcosm survey. Biorem J 2: 125144.
  • Oude Elferink SJWH, Akkermans-van Vliet WM, Bogte JJ & Stams AJM (1999) Desulfobacca acetoxidans gen. nov., sp. nov., a novel acetate-degrading sulfate reducer isolated from sulfidogenic granular sludge. Int J Sys Bacteriol 49: 345350.
  • Phelps CD & Young LY (1999) Anaerobic biodegradation of BTEX and gasoline in various aquatic sediments. Biodegradation 10: 1525.
  • Phelps CD, Kazumi J & Young LY (1996) Anaerobic degradation of benzene in BTX mixtures dependent on sulfate reduction. FEMS Microbiol Lett 145: 433437.
  • Phelps CD, Kerkhof LJ & Young LY (1998) Molecular characterization of a sulfate-reducing consortium which mineralizes benzene. FEMS Microbiol Ecol 27: 269279.
  • Pruesse E, Knittel K, Fuchs BM, Ludwig W, Peplie J & Glöckner FO (2007) SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA data compatible with ARB. Nucleic Acids Res 35: 71887196.
  • Qiu Y-L, Sekiguchi Y, Imachi H, Kamagata Y, Tseng IC, Cheng SS, Ohashi A & Harada H (2003) Sporotomaculum syntrophicum sp. nov., a novel anaerobic, syntrophic benzoate-degrading bacterium isolated from methanogenic sludge treating wastewater from terephthalate manufacturing. Arch Microbiol 179: 242249.
  • Qiu Y-L, Sekiguchi Y, Hanada S, Imachi H, Tseng IC, Cheng SS, Ohashi A, Harada H & Kamagata Y (2006) Pelotomaculum terephthalicum sp. nov. and Pelotomaculum isophthalicum sp. nov.: two anaerobic bacteria that degrade phthalate isomers in syntrophic association with hydrogenotrophic methanogens. Arch Microbiol 185: 172182.
  • Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61: 262280.
  • Schirmer M, Dahmke A, Dietrich P, Dietze M, Gödeke S, Richnow HH, Schirmer K, Weiß H & Teutsch G (2006) Natural attenuation research at the contaminated megasite Zeitz. J Hydrol 328: 393407.
  • Schwarz JIK, Lueders T, Eckert W & Conrad R (2007) Identification of acetate-utilizing Bacteria and Archaea in methanogenic profundal sediments of Lake Kinneret (Israel) by stable isotope probing of rRNA. Environ Microbiol 9: 223237.
  • Spring S, Amann R, Ludwig W, Schleifer K-H, Van Gemerden H & Petersen N (1993) Dominating role of an unusual magnetotactic bacterium in the microaerobic zone of a freshwater sediment. Appl Environ Microbiol 59: 23972403.
  • Stelzer N, Büning C, Pfeifer F, Dohrmann AB, Tebbe CC, Nijenhuis I, Kästner M & Richnow HH (2006) In situ microcosms to evaluate natural attenuation potentials in contaminated aquifers. Org Geochem 37: 13941410.
  • Thauer RK, Jungermann K & Decker K (1977) Energy conservation in chemotrophic anaerobic bacteria. Bacteriol Rev 41: 100180.
  • Ulrich AC & Edwards EA (2003) Physiological and molecular characterization of anaerobic benzene-degrading mixed cultures. Environ Microbiol 5: 92102.
  • Ulrich AC, Beller HR & Edwards EA (2005) Metabolites detected during biodegradation of 13C6-benzene in nitrate-reducing and methanogenic enrichment cultures. Environ Sci Technol 39: 66816691.
  • Van Agteren MH, Keuning S & Janssen DB (1998) Handbook on Biodegradation and Biological Treatment of Hazardous Organic Compounds. Kluwer Academic Publishers, Dordrecht, the Netherlands.
  • Vieth A, Kästner M, Schirmer M, Weiß H, Gödeke S, Meckenstock RU & Richnow HH (2005) Monitoring in situ biodegradation of benzene and toluene by stable carbon isotope fractionation. Environ Toxicol Chem 24: 5160.
  • Vogel TM & Grbic-Galic D (1987) Transformation of toluene and benzene by mixed methanogenic cultures. Appl Environ Microbiol 53: 254260.
  • Vogt C, Gödeke S, Treutler HC, Weiß H, Schirmer M & Richnow HH (2007) Benzene oxidation under sulfate-reducing conditions in columns simulating in situ conditions. Biodegradation 18: 625636.
  • Weiner JM & Lovley DR (1998) Anaerobic benzene degradation in petroleum-contaminated aquifer sediments after inoculation with a benzene-oxidising enrichment. Appl Environ Microbiol 64: 775778.
  • Widdel F & Rabus R (2001) Anaerobic biodegradation of saturated and aromatic hydrocarbons. Curr Opin Biotechnol 12: 259276.
  • Zengler K, Richnow HH, Rosselló-Mora M, Michaelis W & Widdel F (1999) Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 401: 266269.
  • Zhang T, Ke SZ, Liu Y & Fang HP (2005) Microbial characteristics of a methanogenic phenol-degrading sludge. Water Sci Technol 52: 7378.

Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1. Phylogenetic relationship of the Magnetobacterium-related clones (bold-faced) retrieved from lava granulate to representatives of the phylum Nitrospira.

Fig. S2. Phylogenetic relationship of the Sulfurovum-related clones (bold-faced) retrieved from the benzene-degrading consortium ZzBs1-4 to representatives of the Epsilonproteobacteria.

This material is available as part of the online article from: http://www.blackwell-synergy.com/doi/abs/10.1111/j.1574-6941.2008.00536.x (this link will take you to the article abstract).

Please note: Blackwell Publishing is not responsible for the content or functionality of any supplementary materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.

FilenameFormatSizeDescription
FEM_536_sm_figS1.ppt154KSupporting info item
FEM_536_sm_figS2.ppt151KSupporting info item

Please note: Wiley Blackwell is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.