Anaerobic Methyl tert-Butyl Ether-Degrading Microorganisms Identified in Wastewater Treatment Plant Samples by Stable Isotope Probing

Weimin Sun, Xiaoxu Sun, and Alison M. Cupples

Department of Civil and Environmental Engineering, Michigan State University, East Lansing, Michigan, USA

Anaerobic methyl tert-butyl ether (MTBE) degradation potential was investigated in samples from a range of sources. From these 22 experimental variations, only one source (from wastewater treatment plant samples) exhibited MTBE degradation. These microcosms were methanogenic and were subjected to DNA-based stable isotope probing (SIP) targeted to both bacteria and archaea to identify the putative MTBE degraders. For this purpose, DNA was extracted at two time points, subjected to ultracentrifugation, fractioning, and terminal restriction fragment length polymorphism (TRFLP). In addition, bacterial and archaeal 16S rRNA gene clone libraries were constructed. The SIP experiments indicated bacteria in the phyla Firmicutes (family Ruminococcaceae) and Alphaproteobacteria (genus Sphingopyxis) were the dominant MTBE degraders. Previous studies have suggested a role for Firmicutes in anaerobic MTBE degradation; however, the putative MTBE-degrading microorganism in the current study is a novel MTBE-degrading phylotype within this phylum. Two archaeal phylotypes (genera Methanosarcina and Methanocorpusculum) were also enriched in the heavy fractions, and these organisms may be responsible for minor amounts of MTBE degradation or for the uptake of metabolites released from the primary MTBE degraders. Currently, limited information exists on the microorganisms able to degrade MTBE under anaerobic conditions. This work represents the first application of DNA-based SIP to identify anaerobic MTBE-degrading microorganisms in laboratory microcosms and therefore provides a valuable set of data to definitively link identity with anaerobic MTBE degradation.

Methyl tert-butyl ether (MTBE) is a synthetic organic com-pound that was added to gasoline in the late 1970s, following the phaseout of tetra-ethyl lead. Later, the implementation of the Clean Air Act Amendments (1990) caused a significant increase in MTBE use. In 1970, MTBE was the 39th highest produced U.S. organic chemical, whereas by 1998 it ranked 4th. During this time, the aggregate production of MTBE was 60 million metric tons (12). The large-scale use, combined with MTBE’s physiochemical properties, has resulted in severe contamination. MTBE has a high water solubility (51 g liter1), strongly partitions to water from air (dimensionless Henry’s law constant, 0.02) (16), has a low sorption partition coefficient (Koc is 1.035 to 1.091 [17]) and, thus, is highly mobile in water (52). MTBE contamination has been reported in surface waters (2, 3, 9, 12, 17, 26, 46), groundwater (12, 17, 27, 28, 31, 35, 37, 57, 63, 69), and drinking water sources (1, 6, 12, 22, 33, 39, 50, 56, 59). MTBE contamination in drinking water has been reported in 36 states, with removal estimates being $25 billion to $33.2 billion (5).

Biological degradation is becoming increasingly common as a remediation method for groundwater contaminants, either through natural attenuation or enhanced bioremediation. There have been numerous studies on aerobic MTBE biodegradation (29, 45, 61, 62), and microorganisms capable of degrading MTBE under aerobic conditions have been isolated (18, 24, 36, 49). Anaerobic MTBE biodegradation has also been documented (10, 19, 38, 44, 53–55); however, much less is known about the microorganisms involved and, to date, no MTBE-degrading isolates have been obtained. This knowledge gap is a significant limitation to in situ MTBE bioremediation because many contaminated sites are anaerobic. More information on the microorganisms capable of anaerobic MTBE degradation could result in the application of molecular methods to investigate their presence and abundance and, therefore, the potential for MTBE degradation at different sites.

The microbial composition of anaerobic MTBE-degrading enrichment cultures has only recently been investigated. In 2009, the microbial communities of three anaerobicMTBE-degrading cultures derived from MTBE-contaminated aquifer material were examined using 16S rDNA-based amplified ribosomal DNA restriction analysis (ARDRA) and 16S rRNA gene sequencing (60). The cultures were maintained with anthroquinone-2,6-disulfonate (AQDS), sulfate, or fumarate as electron acceptors, and the authors found that the microbial diversity varied under these different conditions. In another recent study, other researchers characterized the community composition of anaerobic enrichment cultures originating from three different contaminated sediments (67). Interestingly, terminal restriction fragment length polymorphism (TRFLP) profiles indicated substantially different community profiles from MTBE-degrading microcosms established from different sediment sources. A third group investigated the microbial community present (using 16S rRNA gene sequencing) when MTBE degradation occurred under sulfate- or iron-reducing conditions or when both electron acceptors were present together and identified five to eight microorganisms in the three consortia (44).

The current study expands on these investigations by applying DNA-based stable isotope probing (SIP) to determine which organisms are responsible for 13C label uptake from MTBE in anaerobic MTBE-degrading microcosms under methanogenic conditions. The SIP method is unique in that it can directly identify the microorganisms responsible for contaminant degradation and therefore offers more targeted information than community analysis alone (e.g., TRFLP, ARDRA, 16S rRNA clone libraries). In 2008, phospholipid-SIP was used to examine MTBE and tert-butyl alcohol (TBA) biodegradation potential using “bio-traps” in gasoline-contaminated aquifers (11). However, to the authors’ knowledge, DNA-based SIP has yet to be used to investigate anaerobic MTBE degradation. The DNA-based SIP method involves exposure of mixed cultures to the labeled compounds of interest (e.g., [13C]MTBE) and DNA extraction over time. The DNA is then subjected to ultracentrifugation, fractionation (to separate label incorporated DNA from the unlabeled DNA), and TRFLP on each fraction. Any TRFLP fragment illustrating an increase in relative abundance in the heavy fraction of the samples (exposed to labeled substrate) compared to the controls (exposed to unlabeled substrate) is identified (using 16S rRNA gene clone libraries) as the putative degrader. The method has been used to identify the microorganisms involved in the degradation of numerous contaminants (4, 15, 21, 34, 40, 51, 58, 64, 65). In the current study, a range of sources (contaminated site sediment, agricultural soils, and wastewater treatment samples) were examined for anaerobic MTBE degradation potential. In the active anaerobic MTBE-degrading microcosms, SIP targeted to both bacteria and archaea was applied to identify the putative MTBE-degrading microorganisms.

MATERIALS AND METHODS
Microcosm construction and analytical techniques. A range of sources were tested for their potential to degrade MTBE (see Table S1 in the supplemental material), including agricultural soil, contaminated site soil, and wastewater treatment plant (WWTP) samples. From all samples tested, only one source (WWTP sample) demonstrated MTBE degradation and was further investigated. Microcosms were prepared under strictly anaerobic conditions in an anaerobic chamber (Coy Laboratory Products Inc., Grass Lake, MI). For each microcosm, an 6-g sample (wet weight) was anoxically incubated in 60-ml serum bottles containing 25 ml of anaerobic basal media (66). The SIP experiments involved triplicate abiotic controls and triplicate unlabeled-MTBE (99%; Sigma-Aldrich, St. Louis, MO)- and triplicate labeled-MTBE ([13C5]MTBE; Sigma-Aldrich, St. Louis, MO)-amended samples. These microcosms were incubated at room temperature (20°C) with reciprocal shaking. MTBE concentrations in headspace gas samples (200 l) were determined with a gas chromatograph (Perkin Elmer) equipped with a flame ionization detector and a capillary column (DB-624; diameter, 0.53 mm; J&W Scientific). Injector and detector temperatures were set at 200°C, and the column temperature was 120°C.
DNA extraction and ultracentrifugation. For the SIP experiments, DNA was extracted at two time points (30% and 70% MTBE removal after 6 and 29 days, respectively) during MTBE depletion from microcosms amended with labeled MTBE. When 70% MTBE was removed, DNA was extracted from microcosms amended with unlabeled DNA. The PowerSoil DNA extraction kit (MO BIO Laboratories, Inc., Carlsbad, CA) was used for total nucleic acid extraction according to the manufacturer’s recommended procedure. Quantified DNA extracts (10 g) were loaded into Quick-Seal polyallomer tubes (13 by 51 mm, 5.1 ml; Beckman Coulter) along with a Tris-EDTA (pH 8.0)-CsCl solution. Prior to sealing (cordless Quick-Seal tube topper; Beckman), the buoyant density (BD) was determined with a model AR200 digital refractometer (Leica Microsystems Inc.) and adjusted by adding small volumes of CsCl solution or Tris-EDTA buffer with a final BD of 1.7300 g ml1. The tubes were centrifuged at 178,000 g (20°C) for 48 h in a StepSaver 70 V6 vertical titanium rotor (8 by 5.1 ml capacity) within a Sorvall WX 80 Ultra Series centrifuge (Thermo Scientific). Following centrifugation, the tubes were placed onto a fraction recovery system (Beckman), and fractions (150 l) were collected. The BD of each fraction was measured, and CsCl was removed by glycogen-assisted ethanol precipitation. For each sample, approximately 20 fractions were obtained; however, TRFLP was performed (see below) only on fractions (between 6 to 13 fractions for each sample) that successfully produced an amplicon.

PCR, TRFLP, and sequencing of 16S rRNA genes. The density-resolved fractions from [12C]- and [13C]MTBE-amended microcosms were PCR-amplified using 27F-FAM (5=-AGAGTTTGATCMTGGCTCAG, 5= end-labeled with carboxyfluorescein) and 1492R (5=-GGTTACCTTGTT ACGACTT) for generating bacterial amplicons and A109F-FAM (5=-AC KGCTCAGTAACACGT) and A934R (5=-GTGCTCCCCCGCCAATT CCT) for archaeal amplicons (Operon Biotechnologies). The presence of PCR products was confirmed by 1.5% agarose gel electrophoresis and the subsequent staining of the gels with ethidium bromide. PCR products were purified with a QIAquick PCR purification kit (Qiagen Inc.) following the manufacturer’s instructions, and approximately 150 ng was digested with HaeIII (New England BioLabs) with a 6-hour incubation period. Three additional digests (MspI, RsaI, MseI) for TRFLP analyses in a number of heavy labeled fractions were included to correlate the TRFLP fragment lengths to the in silico cut sites of the cloned 16S rRNA gene sequences. DNA fragments were separated by capillary electrophoresis (ABI Prism 3100 genetic analyzer; Applied Biosystems) at the Research Technology Support Facility (RTSF) at Michigan State University. Data were analyzed with GeneScan software (Applied Biosystems), and the percent abundance of each fragment was determined. Clone libraries of the 16S rRNA genes were constructed using total genomic DNA amplified with 27F/1492R and A109F/A934R as above except the forward primer was unlabeled, and the final extension time was extended to 15 min. To reduce sequencing redundancy, restriction fragment length polymorphism (RFLP) analysis was performed and specific operational taxonomic units (OTU) were selected for sequencing. The PCR products were purified with a QIAquick PCR purification kit (Qiagen Inc.) and cloned into Escherichia coli TOP10 vector supplied with a TOPO TA cloning kit (Invitrogen Corporation). E. coli clones were grown on Luria-Bertani (LB) medium solidified with 15 g agar liter1with 50g ampicillin liter1 for 16 h at 37°C. Colonies with inserts were verified by PCR with primers M13 F (5=-TGTAAAACGACGGCCAGT-3=) and M13 R (5=-AACAGCT ATGACCATG-3=), plasmids were extracted from the positive clones with a QIAprep miniprep system (Qiagen, Inc.), and the insertions were sequenced at RTSF. The Ribosomal Database Project (RDP) (Center for Microbial Ecology, Michigan State University) analysis tool “classifier” was utilized to assign taxonomic identity. Phylogenetic trees for the partial 16S rRNA gene sequences of the putative MTBE degraders along with the closest matches in GenBank were obtained by the neighbor-joining method using MEGA 5.0 software.

Nucleotide sequence accession numbers. The 16S rRNA gene clones libraries obtained have been deposited in GenBank under accession numbers JQ423059 to JQ423114.

RESULTS AND DISCUSSION
From 22 experimental setups, anaerobic MTBE biodegradation was noted in microcosms constructed from only one source and only under methanogenic conditions (see Table S1 in the supplemental material). Microcosms for the SIP experiment were constructed using freshly sampled material from the East Lansing (MI) WWTP. MTBE degradation occurred in both labeled MTBE-amended and unlabeled-MTBE-amended samples but not in the abiotic controls (Fig. 1). Methane production was observed following 2 days of incubation and increased rapidly as MTBE was degraded (data not shown). Bacterial microbial communities were profiled by SIP and TRFLP when 30% and 70% MTBE was
degraded, whereas archaeal communities were only profiled when 70% was degraded. Assimilation of 13C-labeled MTBE was detected by comparing TRFLP profiles of DNA derived from labeled treatments with DNA from unlabeled treatments. Specifically, the organisms responsible for 13C assimilation were identified by the comparison of relative abundances of specific terminal restriction fragments (TRFs) between the labeled and unlabeled gradient fractions (see example TRFLP profiles in Fig. S1 in the supplemental material).

Labeled bacterial DNA was successfully amplified in the heavier fractions (higher buoyant density or BD values) in DNA extracted from [13C]MTBE-amended microcosms from both time points (30 and 70% degraded). Two bacterial TRFs (67 bp and 215 bp) were highly enriched in the heavy 13C fractions, while such enrichment was not seen in the corresponding 12C fractions with similar buoyant densities (Fig. 2A and B). A high (30%) relative abundance (RA) of both TRFs was noted in the heavier 13C fractions, and their RA increased over time, as MTBE was degraded (30% and 70% MTBE degraded). When 30% MTBE had been degraded, the maximum RA of the 215-bp TRF from the samples ([13C]MTBE-amended samples) was 15% in the heavy fractions, and this increased to 50% at the second time point (Fig. 2A). The maximum RA of the 67-bp TRF in the heavy fractions of the samples ([13C]MTBE amended) at the first time point was 20%, and again this increased to 50% at the later time point (Fig. 2B). An analysis of the archaeal SIP TRFLP profiles (from 70% MTBE degraded) indicated two TRFs (132 bp and 162 bp) were relatively more dominant in the labeled fractions than in the controls (Fig. 3A and B). However, these enrichment patterns were less pronounced (compared to the bacterial enrichment patterns); therefore, it is likely these organisms played only a syntrophic role during MTBE degradation.

Clone libraries for both bacteria (see Table S2 in the supplemental material) and archaea (Table S3) were generated to both investigate the diversity of the microbial community within the MTBE-degrading microcosms and to identify the putative MTBE degraders represented by the TRFs discussed above. The most dominant bacterial phylum was the Proteobacteria (59 from 122 clones), and the second most dominant phylum was the Firmicutes(29/122), which primarily consisted of Clostridia (26/29). The genera of each clone determined from RDP are also shown, with a significant number (13/26) from being unclassified to the genus level (Table S2). The archaeal community was less diverse with only 7 different phylotypes (Table S3), all within the class Methanomicrobia (phylum Euryarchaeota). Two phylotypes could not be classified to the genus level. The bacterial and archaeal sequences were digested in silico to identify the TRFs enriched in the heavy fractions as discussed above. The two bacterial TRFs dominant in the heavy fractions, and thus the two putative MTBE degraders, were identified as Clostridia (215 bp; GenBank accession no. JQ428827) and Alphaproteobacteria (67 bp; GenBank accession no. JQ428826). The putative identification of the bacterial TRFs was confirmed with additional digests on the heavy fractions (Table S4). The Clostridia-related phylotype could be classified to the family Ruminococcaceae and was most similar (98% sequence identity) to uncultured clones from Arctic permafrost soil from northern Norway (GenBank accession no. EF034844.1, EF034735.1, and EF34712.1) (23) and to an uncultured Acetivibrio sp. clone ZZ-S2G3 (95% sequence identity) from amixedmicrobial communitymineralizing benzene under sulfate-reducing conditions (GenBank accession no. EF613411.1) (30) (Fig. 4). The isolate with the highest level of similarity (88% sequence identity) was Clostridium sp. strain P6 isolated from sludge-treating brewery wastewater (GenBank accession no. AY949857) (14). The other most similar sequences were all uncultured bacteria from a range of environments, including bulking sludge (GenBank accession no. HQ538642), a polychlorinated biphenyl (PCB) dechlorinating community (GenBank accession no. GU325884), other clones from the above permafrost study (GenBank accession no. EF034651, EF03415, EF034551, EF034481, EF034335), another clone from the above benzene study (GenBank accession no. EF613469), a clone from an SIP benzene-degrading study (GenBank accession no. FN824839), a clone from subsurface coal beds and methanogenic coal enrichment cultures (GenBank accession no. EU073776), and a clone from sediments polluted with nitrobenzene (GenBank accession no. EF590062). The putativeMTBEClostridia-related phylotype identified in the current study had a low level of similarity to these uncultured clones (between 89% and 95%). The closest type strain to the Clostridia-related putative MTBE degrader was Clostridium thermocellum ATCC 27405 (GenBank accession number CP000568.1).

However, in this case, the partial 16S rRNA gene of the putative MTBE degraderwas only 86% similar (539from 630 bpwere similar) to C. thermocellum ATCC 27405.

The Alphaproteobacteria-related clone was classified to the genus Sphingopyxis and exhibited a high level of similarity (98 to 99% sequence identity) to many uncultured clones (Fig. 5). These uncultured clones were also obtained from a diverse range of sources, including other wastewater samples (GenBank accession no. HQ158685, HQ385511, HQ592561), communities involved in anaerobic digestion of sludge (GenBank accession no. CU926829, CU926428, CU925739, CU927137, CU926644, CU927293) (47), cyanobacterial water blooms (GenBank accession no. AM989067) (7), a stream community (GenBank accession no. JF697411), sulfidic spring water (GenBank accession no. JF747974), sediment enriched with brominated flame retardant (GenBank accession no. JF345300, JF345291, JF345333) and a membrane bioreactor (GenBank accession no. FN827250). The closest type strain to the Alphaproteobacteria-related putative MTBE degrader was Sphingopyxis alaskensis RB2256 (GenBank accession number CP000356.1). The partial 16S rRNA gene of the putative MTBE degrader was 95% similar (649 from 682 bp were similar) to S. alaskensis RB2256.

The archaeal 132-bp and 162-bp TRFs belonged to the genera Methanosarcina and Methanocorpusculum, respectively. However, the level of enrichment was low in the archaeal phylotypes in the heavy fractions, and so it is unlikely that these organisms (genera Methanosarcina and Methanocorpusculum) were the dominant degraders. The Methanosarcina phylotype may have incorporated labeled acetate (from [13C]MTBE biodegradation), as these organisms have previously been described as acetate and H2 scavengers (8). The Methanocorpusculum phylotype may have incorporated 13CO2 (also from [13C]MTBE biodegradation), as previous research indicated that members of the genus Methanocorpusculumproduce methane using H2-CO2 (71, 72). It is possible that the identified archaeal phylotypes were responsible for minor amounts of MTBE degradation. However, they more likely played a syntrophic role during MTBE biodegradation.

Interestingly, other researchers have identified Clostridia as dominant organisms in their anaerobic MTBE-degrading enrichments. From the three enrichments (AQDS, sulfate, or fumarate reducing) developed from MTBE-contaminated aquifer material, Interestingly, other researchers have identified Clostridia as dominant organisms in their anaerobic MTBE-degrading enrichments. From the three enrichments (AQDS, sulfate, or fumarate reducing) developed from MTBE-contaminated aquifer material, the sulfate-reducing enrichment contained 19.3% assigned to the order Clostridiales and the fumarate-reducing enrichment contained a dominant clone (related to Clostridium sp. Kw12) (22.8%) also belonging to the phylum Firmicutes (60). Similarly, following continual enrichment of an anaerobic MTBE-degrading consortium, researchers reduced the community to three dominant phylotypes belonging to Deltaproteobacteria, Chloroflexi, and Firmicutes (67). In addition, Clostridia were found in anaerobic MTBE-degrading consortia under sulfate- and iron-reducing conditions (44). These previous studies, combined with the data in the current study, indicate that organisms in the phylum Firmicutes are important for the anaerobic degradation of MTBE. Although these organisms are all within the same phylum, the putative MTBE degrader in the current study represents the first link of this particular phylotype to MTBE degradation.

Microorganisms associated with the family of Ruminococcaceae (phylum Firmicutes) are predominant members of mammalian gut microbial flora. The presence of these organisms in the enrichments is therefore not surprising given the source of the inocula (WWTP sample). Members of Ruminococcaceae isolated from human gut were previously correlated with the biodegradation of complex polysaccharides such as starch or xylan (32). In addition, Ruminococcaceae isolated from rumen or human guts were proven to be able to degrade cellulose (13, 20, 48). Further, Chassard et al. reported Ruminococcaceaewere responsible for cellulose biodegradation in the fecal samples of methane-excreting subjects while the main cellulose-degrading bacteria belong essentially to Bacteroidetes in non-methane-excreting subjects (13), indicating a possible link between methane production and Ruminococcaceae-associated cellulose biodegradation. This observation is consistent with the current study in that no MTBE biodegradation under the sulfate- and nitrate-amended conditions (no methane was produced), although they were seeded from the same inocula. Ruminococcaceae were also linked with 2,4,6-trinitrotoluene (TNT) degradation in a recent study (42).

The other putative MTBE degrader identified in the current study (TRF, 67 bp) was classified within the genus Sphingopyxis (phylum Alphaproteobacteria, family Sphingomonadaceae). Interestingly, another genus (Sphingomonas) within the same family contains previously reported MTBE degraders. For example, a Sphingomonassp. isolate was shown to grow aerobically on MTBE as the sole source of carbon and energy (41). Further, Sphingomonas spp. have also been associated with aerobic MTBE biodegradation in bioreactors (43, 70). Recently, a 16S rRNA gene sequence classifying as a Sphingomonas sp. was found in an MTBE-degrading enrichment culture amended with iron and sulfate as electron acceptors (44). The results of the current study indicate the Sphingopyxis phylotype may be a novel anaerobic MTBE degrader within this family that warrants further investigation for potential anaerobic remediation applications.

The presence of two enriched TRFs suggests more than one microorganism may be responsible for MTBE biodegradation (the Clostridia-related phylotype, which classified to the family Ruminococcaceae, and the Alphaproteobacteria-related clone, which classified to the genus Sphingopyxix). Youngster et al. (68) suggested that anaerobic MTBE biodegradation required the interaction of a consortium. However, the exact mechanism of MTBE anaerobic biodegradation in the current study has yet to be elucidated. Others have reported TBA as an MTBE degradation metabolite (24, 25, 54), indicating that the cleavage of the ether bond is the initial step of MTBE biodegradation. Unfortunately, TBA was not investigated in the current study. The C—O—C bond found in MTBE also occurs in cellulose; therefore, one might hypothesize that Ruminococcaceae, which can degrade cellulose, may be responsible for the initial step of MTBE biodegradation.

ACKNOWLEDGMENTS

Funding for this work was provided by a grant awarded to A. Cupples from the National Science Foundation (0853249). We thank Catherine Garnham for her help on sampling the activated sludge from the East Lansing wastewater treatment plant.

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