Microbial Communities Involved in Methane ... - ACS Publications

Aug 15, 2012 - chain n-alkanes and certain monoaromatic hydrocarbons, in oil sands tailings ponds produces large volumes of CH4 in situ. We characteri...
0 downloads 0 Views 1MB Size
Download PDF
Article pubs.acs.org/est

Microbial Communities Involved in Methane Production from Hydrocarbons in Oil Sands Tailings Tariq Siddique,†,* Tara Penner,‡,∥ Jonathan Klassen,‡,⊥ Camilla Nesbø,‡,§ and Julia M. Foght‡,* †

Department of Renewable Resources, University of Alberta, Edmonton, AB, T6G 2E3, Canada Department of Biological Sciences, University of Alberta, Edmonton, AB, T6G 2E9, Canada § Department of Biology, University of Oslo, Oslo NO-0316, Norway ‡

S Supporting Information *

ABSTRACT: Microbial metabolism of residual hydrocarbons, primarily shortchain n-alkanes and certain monoaromatic hydrocarbons, in oil sands tailings ponds produces large volumes of CH4 in situ. We characterized the microbial communities involved in methanogenic biodegradation of whole naphtha (a bitumen extraction solvent) and its short-chain n-alkane (C6−C10) and BTEX (benzene, toluene, ethylbenzene, and xylenes) components using primary enrichment cultures derived from oil sands tailings. Clone libraries of bacterial 16S rRNA genes amplified from these enrichments showed increased proportions of two orders of Bacteria: Clostridiales and Syntrophobacterales, with Desulfotomaculum and Syntrophus/Smithella as the closest named relatives, respectively. In parallel archaeal clone libraries, sequences affiliated with cultivated acetoclastic methanogens (Methanosaetaceae) were enriched in cultures amended with n-alkanes, whereas hydrogenotrophic methanogens (Methanomicrobiales) were enriched with BTEX. Naphtha-amended cultures harbored a blend of these two archaeal communities. The results imply syntrophic oxidation of hydrocarbons in oil sands tailings, with the activities of different carbon flow pathways to CH4 being influenced by the primary hydrocarbon substrate. These results have implications for predicting greenhouse gas emissions from oil sands tailings repositories.

■

INTRODUCTION Northern Alberta, Canada has one of the world’s largest oil sands reserves, producing over 85 million m3 of bitumen in 2011.1 Surface mining of oil sands followed by bitumen extraction using water and hydrocarbon solvent (e.g., naphtha) produces oil sands tailings that are deposited in settling basins (colloquially, “tailings ponds”). These fluid fine tailings comprise a thin slurry of water, sand, silt, clays (∼8−12 wt % solids), residual bitumen (∼5 wt %) and unrecovered solvent (≤0.5 wt %). Sand settles quickly near the inflow to the pond, whereas the fine clay suspensions in the pond slowly consolidate over 5−10 years to become mature fine tailings (MFT) at ∼25−30 wt % solids content. The producers operate under a zero discharge policy: the accumulated inventory of MFT requiring long-term confinement is currently ∼830 million m3.2 All of the settling basins tested to date are microbiologically active, and most produce methane (CH4). Mildred Lake Settling Basin, the largest tailings pond at Syncrude Canada Ltd., contains >400 million m3 of tailings3 and emits an estimated 43 000 m3 CH4 day−1 plus an unknown volume of carbon dioxide (CO2).4 However, precise measurement of stochastic gas bubbling over large surface areas of the settling ponds is technically challenging. Thus, a preliminary kinetic model based on laboratory studies5 was developed to describe and predict in situ methane emissions; additional microbiological data would contribute to refinement of such models. © 2012 American Chemical Society

Microbial production of CH4 accelerates the consolidation of fine clay particles and facilitates the recovery of pore-water from MFT for reuse in ore processing.6 The juxtaposition of this beneficial phenomenon with deleterious greenhouse gas emissions sparked interest in discovering the source of CH4 production in tailings ponds. We determined in previous laboratory studies that naphtha, composed of C3−C14 paraffins (n-alkanes), iso-paraffins (branched alkanes), naphthenes (cycloalkanes), and monoaromatics (benzene, toluene, ethylbenzene, and xylenes; BTEX), supported methanogenesis.7 In particular, the n-alkanes (C6−C10) and certain BTEX compounds (toluene and xylenes) in naphtha, as well as long-chain n-alkanes (C14− C18) associated with residual bitumen in MFT were metabolized to CH4 by indigenous microbes during ∼1 year of incubation.7−9 Microbial communities in native MFT have been characterized using 16S rRNA gene clone libraries,10 revealing high bacterial diversity and lesser archaeal diversity. However, the roles of different microbial community members in particular metabolic pathways could not be assigned. Furthermore, in general the hydrocarbon biodegradation pathways under methanogenic conditions are unproven and remain subjects of Received: Revised: Accepted: Published: 9802

June 1, 2012 August 14, 2012 August 15, 2012 August 15, 2012 dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

Figure 1. Methane (CH4) production due to biodegradation of hydrocarbons in primary enrichment cultures containing mature fine tailings from Mildred Lake Settling Basin. (A) CH4 production from n-alkane-, naphtha- or BTEX-amended cultures or unamended (control) tailings. (B) Biodegradation of short-chain n-alkanes during 46 weeks of incubation with mature fine tailings under methanogenic conditions. The figures contain data reported by Siddique et al.7,8 plus new data (see text). Data points are the mean of triplicate cultures and error bars, where visible, represent the standard deviation.

speculation,11 with a few exceptions (e.g., toluene12). We therefore established primary enrichment cultures by amending MFT with whole naphtha or specific naphtha components and analyzed changes in community structure compared with cultures containing unamended MFT after lengthy anaerobic incubation (36 or 46 weeks) so as to infer the microbes and pathways involved in the degradation of particular substrates. In addition to providing fundamental insights into methanogenic hydrocarbon biodegradation to CH4, the results of this study may facilitate development and refinement of mechanism-based

ecosystem models for predicting CH4 emissions5 and linkage of microbial functions to accelerated dewatering and settling of oil sands tailings for the reclamation and management of tailings ponds.

■

EXPERIMENTAL SECTION

Establishment of Primary Enrichment Cultures. The primary cultures (i.e., uncultivated MFT, newly amended with growth medium and hydrocarbons) analyzed here were previously described by Siddique et al.,7,8 including details of 9803

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

Table 1. Heat Map of 20 Most Abundant OTUs in Four Bacterial 16S rRNA Gene Clone Libraries, Reported as Percent of Sequenced Clones within a Library and Arranged in Decreasing Order of Total Clone Abundance in All Four Libraries; Clones were Constructed from Duplicate Cultures Unamended with Hydrocarbons (Control) or Amended with n-alkanes, BTEX or Naphtha and Incubated under Methanogenic Conditions for 36 or 46 Weeks

a

For % similarity to nearest sequence see SI Tables S2−S5 and S6, and Figures S1, S2.

weeks). The DNA from each aliquot was extracted twice sequentially as described previously13 and subjected to PCR amplification in triplicate using bacterial primers PB36 and PB3814 or archaeal primers 21F and 958R15 and Phusion High Fidelity DNA Polymerase (Finnzymes OY, Finland) to yield near full-length 16S rRNA amplicons. PCR conditions for bacterial 16S rRNA gene amplification consisted of a 3 min initial denaturation at 98 °C, 30 cycles of 98 °C for 30 s, 54 °C for 30 s, and 72 °C for 1 min, and a final extension at 72 °C for 10 min. Amplification of archaeal sequences was essentially the same except using 27 cycles with annealing at 51 °C. The PCR products from each treatment (2 culture bottles × 2 aliquots × 2 pooled sequential lysates × 3 independent PCR reactions) were then pooled for bacterial and archaeal clone library construction. Appropriate parallel positive and negative PCR reagent controls were included during gene amplification, yielding the expected results (not shown). Construction of 16S rRNA gene Clone Libraries and Amplified rDNA Restriction Analysis (ARDRA). Bluntended PCR products were cloned into the pJET1 vector following the manufacturer’s protocol (Fermentas Life Sciences, Burlington ON, Canada) to construct bacterial or archaeal 16S rRNA gene clone libraries. Clone inserts were reamplified using primers targeting the plasmid cloning site: pJETF (5′ - GCC TGA ACA CCA TAT CCA TCC - 3′) and pJETR (5′ - GCA

culture preparation, source and chemical descriptions of MFT and naphtha. Previous results of hydrocarbon biodegradation and CH4 production in triplicate cultures are summarized with additional, new data in Figure 1. Briefly, 50 mL of MFT collected from Mildred Lake Settling Basin at 6 m depth was incubated with 50 mL of methanogenic medium in 158-mL serum bottles under a headspace of O2-free 30% CO2, balance N2 at atmospheric pressure. Cultures were amended with a mixture of four short-chain n-alkanes (C6, C7, C8 and C10), BTEX, or Syncrude naphtha diluent (CAS No. 64742-49-0; Table 1 in Siddique et al.7) at concentrations representing those in tailings ponds, or were unamended (i.e., a “baseline” control containing endogenous substrates) and incubated statically at ∼20 °C in the dark (ca. in situ conditions) until CH4 production reached a plateau (36 weeks for BTEX- and 46 weeks for n-alkane- and naphtha-amended cultures). Headspace methane concentrations were determined using a gas chromatograph with flame ionization detector (GC-FID) and residual alkanes in the primary cultures were measured using GC-FID with a purgeand-trap system, as previously described.7 Genomic DNA Extraction and Polymerase Chain Reaction (PCR). Two 300-μL aliquots of primary culture were collected from each of two replicate primary cultures described above (n-alkane-, BTEX- and naphtha-amended or unamended controls; eight bottles total) at the end of incubation (36 or 46 9804

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

conditions using MFT amended with whole naphtha or naphtha fractions, specifically C6−C10 n-alkanes or BTEX. Unamended MFT served as a baseline control to account for metabolism of endogenous residual hydrocarbons and undefined organics in the MFT. CH4 production is shown in a composite graph (Figure 1A) containing previously published data for naphtha and BTEX7 and additional, new data for n-alkanes, extending the previous methane analysis from 29 weeks8 to 46 weeks. Similarly, new data for n-alkane degradation extends the previous incomplete degradation profile at 25 weeks8 to show complete degradation by 46 weeks (Figure 1B). Degradation of amended naphtha, n-alkanes, toluene, o-xylene, and m- and p-xylenes was observed in live cultures compared with parallel heat-killed controls, and stoichiometric calculations confirmed hydrocarbon metabolism to CH4.7,8 For illustration, complete degradation of short-chain n-alkanes is shown in Figure 1B. Isoparaffins and naphthenes (cycloalkanes), which are the other significant components (27−31 wt %) of naphtha, were not degraded during 46 weeks of incubation.7 Composition of Bacterial Community Based on Cloned 16S rRNA Genes. Sequencing of clones from the bacterial 16S rRNA gene library revealed a diverse bacterial community dominated by sequences belonging to the Firmicutes, Deltaproteobacteria, Chloroflexi and Betaproteobacteria (Supporting Information (SI) Table S1). Firmicutes increased from 9% of the sequenced library in the unamended control to 28− 39% of clones from hydrocarbon-amended cultures. In contrast, Chloroflexi and Betaproteobacteria decreased from 41% to 19− 36% and from 20% to 0−3%, respectively, after hydrocarbon amendment. Deltaproteobacteria (17% of control library) increased only in the BTEX- (31%) and naphtha- (26%) amended cultures whereas candidate phylum OP11-related clones (2% of control) increased to 10% only in the n-alkaneamended culture (SI Table S1). The 20 most abundant OTUs are listed in Table 1, and account for 70−85% of all the sequenced clones comprising the four bacterial clone libraries. The majority of Firmicutes OTUs were related to Clostridia, particularly sulfate-reducing genera such as Desulfotomaculum and Desulfosporosinus in the Peptococcaceae (SI Figure S1, Table S2), despite the primary cultures being methanogenic and sulfate-depleted; these OTUs may be fermenting rather than respiring in the cultures. Among these, the Desulfotomaculum-related OTU EU522652 (SI Table S2) was the most abundant OTU in the alkane and naphtha clone libraries (20% and 22% of clones, respectively) but was rarer in the unamended control culture (6%) and undetected in the BTEX-amended culture. These clones were related to the sequences detected in a BTEX-contaminated aquifer, and to sulfate-reducing bacteria involved in anaerobic hydrocarbon degradation in cold hydrocarbon seep in Weddell Sea, Antarctica.26 Firmicutes clones in OTU EU522654, which are distantly related to a different Desulfotomaculum species found in a biodegraded, mesothermic petroleum reservoir in Alaska27 (SI Figure S1), were the dominant OTU (18% of clones) in the BTEX-amended culture but were undetected in the control and alkane-amended clone libraries (Table 1). The other two notably enriched Firmicutes OTUs affiliated with the Peptococcaceae were most closely related to the fermenter Cryptanaerobacter (HM992532; alkanes and naphtha only) and to sulfate-reducing Desulfosporosinus (EU522658; BTEX and naphtha only, see SI Figure S1; Table S2). They were closely related to sequences detected in a crude oil-degrading methanogenic consortium in a sandstone oil reservoir core.28

GCT GAG AAT ATT GTA GGA GAT C - 3′). Each PCR reaction (25 μL) contained 1X PCR buffer (50 mM Tris-HCl, 1.5 mM MgCl2, 0.4 mM β-mercaptoethanol, 0.1 mM bovine serum albumin, 10 mM (NH4)2SO4, and 200 μM each dNTP), 1 μM of each of the primers pJETF and pJETR, 1.25 U Taq DNA polymerase, and 1 μL of liquid cell culture (grown in 100 μL LB broth overnight at 37 °C). The PCR conditions consisted of an initial denaturation at 94 °C for 4 min followed by 25 cycles of 94 °C for 1:30 min, 56 °C for 45 s and 72 °C for 1 min, followed by a final extension of 72 °C for 10 min. For amplified rDNA restriction analysis (ARDRA), PCR amplicons were digested using HaeIII or CfoI under conditions specified by the supplier (Fermentas Life Sciences) and the fragments were separated by agarose electrophoresis and analyzed using GelPro Analyzer Software version 4.5 (Media Cybernetics, Inc.) to determine fragment sizes. Similar ARDRA patterns were grouped and the near full-length (typically ∼1300−1400 nt) 16S rRNA gene sequence determined as representative for at least one clone from each ARDRA pattern having two or more clones, as follows: DNA was reamplified from the selected clones using pJETF and pJETR primers, purified using a High Pure PCR Purification Kit (Roche Diagnostics), sequenced using BigDye Terminator mix (Applied Biosystems Inc.) and resolved on an Applied Biosystems 373A automated DNA sequencer. Subsequently, clones having singleton ARDRA patterns were randomly selected and sequenced to increase the coverage of the libraries. These additional sequences did not substantially increase the OTU diversity of the libraries (as defined below), but rather recapitulated the distribution of taxa already detected in the libraries. The forward primers PB36 and 21F were used for partial sequencing of the additional bacterial and archaeal clones, respectively (typically 700−800 nt). Phylogenetic and Statistical Analyses. 16S rRNA gene sequences were grouped into operational taxonomic units (OTUs) at ≥97% similarity using DOTUR v1.53.16 These near-full-length and a few partial 16S rRNA gene sequences (excluding 9−15% of total clones identified as chimeric sequences using the Mallard and Pintail programs17,18) were compared to DNA sequences in GenBank nr and GenBank refseq RNA (ref 19, accessed October, 2011) to determine closely related species. The sequences were aligned using the NAST (Nearest Alignment Space Termination) algorithm for creating multiple sequence alignments20 and poorly aligned regions were deleted or edited manually in Geneious Pro 5.5.4 (http://www. geneious.com). Maximum Likelihood phylogenetic trees were estimated using PhyML21 as implemented in Geneious using a GTR + Γ + I model with six rate categories as well as RAxML22,23 on the Bioportal web server (http://www.bioportal.uio.no/). Branch support was assessed applying 100 bootstrap replicates. A chi-square significance test implemented in Excel was used to determine the variation in distribution of clones in different clone libraries. Individual confidence intervals were also calculated to assess the probability of clones falling in hydrocarbon-amended or control libraries. In addition, nonmetric multidimensional scaling (NMDS) analysis was used to discriminate among the clone libraries. The ordination was based on thetayc distances24 calculated based on the OTU profiles in the libraries. Distances and NMDS were calculated in MOTHUR v. 1.24.25

■

RESULTS Biodegradation of Hydrocarbons under Methanogenic Conditions. In previous studies,7,8 replicate primary enrichment cultures were established under methanogenic 9805

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

Figure 2. Nonmetric multidimensional scaling (NMDS) plot of the four clone libraries to illustrate their statistically different compositions. Cont, unamended control; Alk, n-alkane-amended; BTEX, BTEX-amended; Naph, naphtha-amended.

amended cultures (Table 1; SI Table S4). The dominant Chloroflexi OTUs in our study were related to several sequences associated with dechlorination of chlorinated solvents31,36 and detected in uncultivated MFT10 (SI Table S4). Candidate Division OP11 (OTU EU522666) was particularly abundant in libraries derived from the n-alkane-amended cultures (Table 1; SI Table S5) and clones were closely related to sequences in an anaerobic toluene-degrading consortium in a tar oil-contaminated plume.32 Betaproteobacterial OTUs (EU522643 and EU522644, affiliated with Rhodoferax fermentans and Acidovorax def luvii with 97% and 99% similarity, respectively), were particularly abundant in the control library (Table 1; SI Table S5). Other OTUs affiliated with the phyla Actinobacteria, Bacteroidetes, Synergistetes, Caldiserica (formerly Candidate Phylum OP5), Spirochaetes, and unclassified bacteria were detected infrequently in the libraries, with no discernible pattern of occurrence (SI Table S5). Analysis of the bacterial clone libraries using pairwise chi square tests confirmed that the four libraries comprise statistically different communities (SI Table S6), despite the small number of clones in some categories. A NMDS plot of the clone libraries also revealed distinctness of the clone libraries, being located in different quadrants (Figure 2). The R2 and stress values indicate the quality of the ordination: a low stress value (i.e., below 0.10) and an R2 close to 1 in Figure 2 indicate a good fit between the data and the ordination. The plot also indicates that n-alkane and naphtha clone libraries appear to be more similar to each other than to BTEX and control libraries. Thus, incubation of MFT with naphtha or selected components (nalkanes or BTEX) significantly shifted the microbial community composition. Composition of Archaeal Community Based on Cloned 16S rRNA Gene. Archaeal 16S rRNA gene clone libraries

Deltaproteobacteria were dominated by Syntrophobacterales (Table 1; SI Figure S2). OTU EU522631, related to the phylogenetically poorly resolved Syntrophus/Smithella group, was more abundant in libraries from hydrocarbon-amended cultures (9−16% of clones) than in the control (3%), and was closely (98%) related to a clone identified as Syntrophus sp. (AJ133795) by Zengler et al.29 involved in methanogenic nhexadecane degradation. A less abundant OTU (HM992528) likewise grouped with this clone and also with a cloned Smithella sequence (GU996558) that was abundant in methanogenic oildegrading cultures inoculated with estuarine sediment.30 OTUs EU522637 and EU522636 were also found in this Smithella/ Syntrophus cluster (SI Figure S2), which also contains sequences found independently in uncultivated methanogenic oil sands tailings10 and in hydrocarbon- and chlorinated solventcontaminated aquifers.31 Several other Syntrophus/Smithellarelated OTUs were also detected at lower abundances, with notable relative abundance of OTU EU522637 in the BTEXenriched library (SI Table S3). The phylogenetic assignment to Syntrophus or Smithella of 16S rRNA gene sequences recovered from hydrocarbon environments is unclear,30 and the same sequence yields different presumptive identities in searches using Ribosomal Database Project (RDPII) and GenBank databases; hence our use of dual designations for these OTUs. Other notable deltaproteobacterial OTUs (EU522638 and EU522641) belonged to the orders Desulfobacterales and Desulfuromonadales, which include sulfate-reducing bacteria (SI Table S3) detected in tar-oil-contaminated aquifer sediment,32,33 dioxindehalogenating34 and tetrachloroethene-reducing acetate-oxidizing cultures35 (SI Figure S2), suggesting a broad role for such microbes in contaminated anaerobic environments. Chloroflexi sequences were abundant in all the clone libraries, although generally less so in those derived from hydrocarbon9806

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

Table 2. Heat Map of Five Most Abundant OTUs in Four Archaeal 16S rRNA Gene Clone Libraries, Reported as Percent of Sequenced Clones within a Library and Arranged in Decreasing Order of Total Clone Abundance in All Four Libraries; Clones Were Constructed from Duplicate Cultures Unamended with Hydrocarbons (Control) or Amended with Naphtha, n-Alkanes or BTEX Components and Incubated under Methanogenic Conditions for 36 or 46 Weeks

a

For % similarity to nearest sequence see SI Table S7 and Figure S3.

microbial roles in mineral precipitation and dissolution that have a significant effect on recovered pore water quality. Primary enrichment cultures established during long-term studies of hydrocarbon degradation7,8 were used here to characterize the community structure of indigenous tailings microbes involved in methanogenesis and to propose general mechanisms of hydrocarbon degradation in tailings ponds by correlating microbial community structure with hydrocarbon metabolism. In bacterial clone libraries, sequences affiliated with Deltaproteobacteria, particularly Syntrophus/Smithella, were abundant and notably increased in relative abundance in clone libraries derived from hydrocarbon-amended cultures relative to those from the unamended control culture. The dominant OTU EU522631 in hydrocarbon-amended cultures was closely related to a Syntrophus sp. (AJ133795) reported by Zengler et al.29 to be clearly implicated in the methanogenic degradation of nhexadecane. In the current study, sequences related to Syntrophus/Smithella spp. may represent syntrophs involved in the initial anaerobic activation of alkanes and/or subsequent beta-oxidation of their metabolites to acetate.30 This supposition corresponds with the concomitant striking enrichment of acetoclastic methanogens (EU522628) in the n-alkane-amended culture and slightly lesser enrichment in the naphtha-amended culture (where n-alkanes are “diluted” with other hydrocarbons). The greater abundance of OTUs EU522631 and EU522628 in our n-alkane-amended culture suggests a functional pairing of Syntrophus/Smithella and Methanosaeta spp. in n-alkane degradation in MFT, supported by the observations of Zengler et al.29 It does, however, contradict the proposal by Gray et al.30 that Smithella degrades n-alkanes as a syntrophic partner of hydrogenotrophic Methanocalculus through syntrophic acetate oxidation. In the current study acetoclastic Methanosaeta OTU EU522627 decreased in n-alkane- and naphtha-amended cultures (albeit less so in the BTEX culture) compared to the unamended control, indicating some preference of association of certain methanogens with hydrocarbon-degrading bacteria. Another factor to consider may be the differential toxicity of certain hydrocarbons to acetoclastic methanogens. In addition to increased proportions of Syntrophus/Smithella clones, the abundance of clostridial OTU EU522652 in the nalkane-amended clone library suggests their role in n-alkane degradation. The dominant clostridial OTUs in the current study were related to putative sulfate-reducing Desulfotomaculum spp. Notably, both the MFT and the methanogenic medium used in

constructed in parallel to the bacterial clone libraries had comparatively low sequence diversity and generally high similarity (98−99%) to reference genome sequences and/or cultivated methanogens (Table2; SI Figure S3; Table S7). Almost all archaeal sequences were related either to acetoclastic methanogens (order Methanosarcinales) or hydrogenotrophic (hydrogen-oxidizing) methanogens (order Methanomicrobiales); only a few clones in OTU EU522629 grouped with unclassified Euryarchaeotes (SI Figure S3). The proportions of acetoclastic and hydrogenotrophic methanogens in the archaeal libraries differed depending on hydrocarbon amendment in the culture. Most (∼75%) of the archaeal OTUs in the library generated from the control enrichment culture were affiliated with putative acetoclastic methanogens (OTUs EU522627 and EU522628, 97% and 99% similar to Methanosaeta harudinacea and Methanosaeta concilii, respectively). The clone library constructed from the alkane-amended enrichment culture was also dominated (∼90%) by the putatively acetoclastic OTU EU522628 and no hydrogenotrophic methanogen sequences were detected. In contrast, the relative abundance of presumptive hydrogenotrophic methanogenic sequences (OTUs EU522625 and EU522626) increased in clone libraries constructed from BTEX- and naphtha-amended cultures to 62% and 35%, respectively. Although the similarity of these sequences to cultivated methanogens was lower (94−96%; SI Table S7), they clearly belonged to the Methanobacteriales within all known strains of hydrogenotrophic methanogens and were most closely related to methanogens detected in an estuary37 and a saline wetland in northern Chile38 (SI Figure S3; Table S7). Pairwise chi square statistical analyses of the archaeal clone libraries indicated significant differences among the libraries (SI Table S8).

■

DISCUSSION Methanogenic hydrocarbon degradation is an important process in tailings ponds. Not only does it produce greenhouse gas emissions but also beneficially increases the consolidation rate of fine clay particles6 to facilitate rapid recovery of pore water in oil sands tailings ponds for reuse in the bitumen extraction process. Therefore, characterization of the microbial communities in tailings ponds and their metabolic potential for hydrocarbon degradation is important to better enable prediction of greenhouse gas emissions from tailings ponds5 and to investigate 9807

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

contrast with the pairing observed by Gray et al.30 The pattern of methanogen OTU abundance in the naphtha culture is a blend of the dominant patterns in the n-alkane and BTEX cultures, suggesting concurrent acetoclastic and hydrogenotrophic methanogenesis during metabolism of aliphatic and aromatic naphtha components. Although the literature commonly attributes distribution and dominance of acetoclastic and hydrogenotrophic methanogens to environmental conditions such as pH, temperature and nutrient availability,50−52 our results suggest that enrichment of particular functional groups of methanogens in response to different hydrocarbon amendments relates to the bacterial generation of different metabolites, thus influencing terminal pathways of degradation. This report extends our previous understanding of the complex microbial communities in oil sands tailings ponds10 and allows us to begin assigning presumptive functions to microbial taxa involved in metabolism of three groups of hydrocarbons (alkanes, BTEX and naphtha) present in oil sands tailings settling basins.

our primary enrichment cultures were depleted in sulfate, precluding significant sulfate reduction. However, in the absence of adequate sulfate for respiration, many sulfate-reducing bacteria can ferment39 or grow syntrophically with a hydrogenconsuming partner, or are capable of homoacetogenic growth on substrates such as lactate, pyruvate, formate and H2+CO2.40−43 Therefore, Clostridia-affiliated sequences including Desulfotomaculum in our hydrocarbon-amended cultures may utilize hydrocarbons or their intermediates to produce substrates (e.g., acetate) for acetoclastic methanogens. Syntrophus/Smithella spp. and clostridia were also abundant in naphtha- and BTEX- amended cultures where hydrogenotrophic archaeal OTU EU522625 (Methanomicrobiales) was enriched 3-fold in the BTEX culture and 2-fold in the naphtha culture, compared to the control library, emphasizing the other major route of CH4 production (hydrogenotrophic methanogenesis). The increase in this OTU was correlated with enriched Syntrophus/Smithella-related sequences (EU522631) and Desulfotomaculum-related sequences (EU522654 and EU522652) in BTEX- and naphtha-amended cultures, suggesting that these Bacteria participate in converting monoaromatics to H2+CO2 for consumption by the Methanomicrobiales. Although Syntrophus/ Smithella are implicated in the degradation of hydrocarbons and their metabolites to acetate, enrichment of hydrogenotrophic methanogens in the crude oil degrading enrichment culture has been attributed to syntrophic acetate oxidation H2 and CO2.30 Some Syntrophus/Smithella are known to syntrophically degrade organic molecules such as fatty acids or aromatic acids when a hydrogen-consuming partner is present to keep hydrogen partial pressures low.44 Chauhan and Ogram 45 proposed that Syntrophus-like bacteria in soils of the Florida Everglades were involved in syntrophic oxidation of acetate leading to methanogenesis. The abundance of clostridial OTU EU522654 in only BTEX- and naphtha-amended cultures also suggests clostridial involvement in the production of H2+CO2 to support hydrogenotrophic methanogens. The role of other observed bacterial groups in hydrocarbon metabolism in our study is not clear. Chloroflexi are common constituents of hydrocarbon-degrading methanogenic environments28,46,47 and might participate in syntrophic metabolism.47 OP11 sequences have been detected in various contaminated and pristine environments (SI Table S5) but their role in anaerobic hydrocarbon degradation is unknown. Betaproteobacterial sequences in OTU EU522643, closely (97%) related to iron-reducing Rhodoferax fermentans, were dominant only in the control library suggesting that they play a secondary role and are out-competed by species directly involved in hydrocarbon metabolism. Detection of related betaproteobacterial sequences in a consortium that degraded propylbenzene metabolites under iron-reducing conditions48 suggests that this OTU may persist in tailings due to its ability to utilize metabolites that accumulate in native MFT. In summary, the increase in hydrogenotroph-affiliated archaeal clones and concomitant decrease of acetoclast-affiliated clones in our BTEX−amended culture suggests that hydrogenotrophic methanogenesis is the dominant route for CH4 production from BTEX, in agreement with crude oil biodegradation observed by Jones et al.49 Conversely, the absence of sequences assigned to Methanomicrobiales in n-alkane-amended cultures indicates that syntrophic hydrocarbon degradation proceeds primarily in association with acetoclastic methanogens in these cultures. These results are supported by observation of methanogenic hexadecane29 and crude oil degradation28 but, interestingly,

■

ASSOCIATED CONTENT

* Supporting Information S

Tables listing best matches of the 16S rRNA gene OTUs to sequences in GenBank databases as well as phylogenetic trees of the major groups comprising the OTUs, placing the sequences generated here into clades with archived sequences from GenBank. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (T.S.); julia.foght@ualberta. ca (J.F.). Present Addresses ∥

Syncrude Canada Ltd., Edmonton Research and Development Centre, Edmonton, AB, T6N 1H4 ⊥ Department of Bacteriology, University of Wisconsin-Madison, Madison, Wisconsin, 5370, United States Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We gratefully acknowledge NSERC (TS, JF), Genome Canada and Genome Alberta (JF), Alberta Innovates−Energy and Environment Solutions (TS, JF), School of Energy and the Environment (TS, JF) and the Norwegian Research Council (CLN; project no. 180444/V40) for funding, the Molecular Biology Services Unit (Univ. Alberta) for sequencing, and Syncrude Canada Ltd. for providing samples of tailings and naphtha.

■

REFERENCES

(1) Government of Alberta. Total oil sands production graph. 2011. http://environment.alberta.ca/apps/OSIPDL/Dataset/Details/46 (accessed August 21, 2012). (2) Government of Alberta Oil Sands Information Portal; Tailings Ponds Surface Area, 2012. http://environment.alberta.ca/apps/osip (accessed August 21, 2012). (3) Energy Resources Conservation Board, Regulations and Directives: Tailings Plans 2010 and 2011. Mildred Lake and Aurora North Baseline Survey for Fluid Deposits, 2010. http://www.ercb.ca/ oilsands/tailings-plans/Syncrude_2010_BaselineReport.pdf (accessed August 21, 2012).

9808

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

(4) Holowenko, F. M.; MacKinnon, M. D.; Fedorak, P. M. Methanogens and sulfate-reducing bacteria in oil sands fine tailings waste. Can. J. Microbiol. 2000, 46, 927−937. (5) Siddique, T.; Gupta, R.; Fedorak, P. M.; MacKinnon, M. D.; Foght, J. M. A first approximation kinetic model to predict methane generation from an oil sands tailings settling basin. Chemosphere 2008, 72, 1573− 1580. (6) Fedorak, P. M.; Coy, D. L.; Dudas, M. J.; Simpson, M. J.; Renneberg, A. J.; MacKinnon, M. D. Microbially-mediated fugitive gas production from oil sands tailings and increased tailings densification rates. J. Environ. Eng. Sci. 2003, 2, 199−211. (7) Siddique, T.; Fedorak, P. M.; MacKinnon, M. D.; Foght, J. M. Metabolism of BTEX and naphtha compounds to methane in oil sands tailings. Environ. Sci. Technol. 2007, 41, 2350−2356. (8) Siddique, T.; Fedorak, P. M.; Foght, J. M. Biodegradation of shortchain n-alkanes in oil sands tailings under methanogenic conditions. Environ. Sci. Technol. 2006, 40, 5459−5464. (9) Siddique, T.; Penner, T.; Semple, K.; Foght, J. M. Anaerobic biodegradation of longer-chain n-alkanes coupled to methane production in oil sands tailings. Environ. Sci. Technol. 2011, 45, 5892− 5899. (10) Penner, T. J.; Foght, J. M. Mature fine tailings from oil sands processing harbor diverse methanogenic cultures. Can. J. Microbiol. 2010, 56, 459−470. (11) Dolfing, J.; Larter, S. R.; Head, I. M. Thermodynamic constraints on methanogenic crude oil biodegradation. ISME J. 2008, 2, 442−452. (12) Fowler, S. J.; Dong, X.; Sensen, C. W.; Suflita, J. M.; Gieg, L. M. Methanogenic toluene metabolism: Community structure and intermediates. Environ. Microbiol. 2012, 14, 754−764. (13) Foght, J. M.; Aislabie, J.; Turner, S.; Brown, C. E.; Ryburn, J.; Saul, D. J.; Lawson, W. Culturable bacteria in subglacial sediments and ice from two Southern Hemisphere glaciers. Microbial Ecol. 2004, 47, 329− 340. (14) Saul, D. J.; Aislabie, J. M.; Brown, C. E.; Harris, L.; Foght, J. M. 2005. Hydrocarbon contamination changes the bacterial diversity of soil from around Scott Base, Antarctica. FEMS Microbiol. Ecol. 2005, 53, 141−155. (15) DeLong, E. F. Archaea in costal marine environments. Proc. Nat. Acad. Sci. USA. 1992, 89, 5685−5689. (16) Schloss, P. D.; Handelsman, J. Introducing DOTUR, a computer program for defining operational taxonomic units and estimating species richness. Appl. Environ. Microbiol. 2005, 71, 1501−1506. (17) Ashelford, K. E.; Chuzhanova, N. A.; Fry, J. C.; Jones, A. J.; Weightman, A. J. New screening software shows that most recent large 16S rRNA gene clone libraries contain chimeras. Appl. Environ. Microbiol. 2006, 72, 5734−5741. (18) Ashelford, K. E.; Chuzhanova, N. A.; Fry, J. C.; Jones, A. J.; Weightman, A. J. At least 1 in 20 16S rRNA sequence records currently held in public repositories is estimated to contain substantial anomalies. Appl. Environ. Microbiol. 2005, 71, 7724−7736. (19) Benson, D. A.; Karsch-Mizrachi, I.; Lipman, D. J.; Ostell, J.; Wheeler, D. L. GenBank. Nucleic Acids Res. 2007, 35, D21−D25. (20) DeSantis, T. Z.; Hugenholtz, P.; Keller, K.; Brodie, E. L.; Larsen, N.; Piceno, Y. M.; Phan, R.; Andersen, G. L. NAST: A multiple sequence alignment server for comparative analysis of 16S rRNA genes. Nucleic Acids Res. 2006, 34, W394−W399. (21) Guindon, S.; Gascuel, O. A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst. Biol. 2003, 52, 696−704. (22) Stamatakis, A.; Ludwig, T; Meier., H. RAxML-III: A fast program for maximum likelihood-based inference of large phylogenetic trees. Bioinformatics 2005, 21, 456−463. (23) Stamatakis, A.; Hoover, P; Rougemont., J. A rapid bootstrap algorithm for the RAxML Web servers. Syst. Biol. 2008, 57, 758−771. (24) Yue, J. C.; Clayton, M. K. A similarity measure based on species proportions. Commun. Stat.−Theory Methods 2005, 34, 2123−2131. (25) Schloss, P. D.; Westcott, S. L.; Ryabin, T.; Hall, J. R.; Hartmann, M.; Hollister, E. B.; Lesniewski, R. A.; Oakley, B. B.; Parks, D. H.; Robinson, C. J.; Sahl, J. W.; Stres, B.; Thallinger, G. G.; Van Horn, D. J.;

Weber, C. F. Introducing mothur: Open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl. Environ. Microbiol. 2009, 75, 7537−7541. (26) Niemann, H.; Fischer, D.; Graffe, D.; Knittel, K.; Montiel, A.; Heilmayer, O.; Nothen, K.; Pape, T.; Kasten, S.; Bohrmann, G.; Boetius, A.; Gutt, J. Biogeochemistry of a low-activity cold seep in the Larsen B area, western Weddell Sea, Antarctica. Biogeosciences 2009, 6, 2383− 2395. (27) Pham, V. D.; Hnatow, L. L.; Zhang, S.; Fallon, R. D.; Jackson, S. C.; Tomb, J. F.; DeLong, E. F.; Keeler, S. J. Characterizing microbial diversity in production water from an Alaskan mesothermic petroleum reservoir with two independent molecular methods. Environ. Microbiol. 2009, 11, 176−187. (28) Gieg, L. M.; Duncan, K. E.; Suflita, J. M. Bioenergy production via microbial conversion of residual oil to natural gas. Appl. Environ. Microbiol. 2008, 74, 3022−3029. (29) Zengler, K.; Richnow, H. H.; Rossello-Mora, R.; Michaelis, W.; Widdel, F. Methane formation from long-chain alkanes by anaerobic microorganism. Nature 1999, 401, 266−269. (30) Gray, N. D.; Sherry, A.; Grant, R. J.; Rowan, A. K.; Hubert, C. R. J.; Callbeck, C. M.; Aitken, C. M.; Jones, D. M.; Adams, J. J.; Larter, S. R.; Head, I. M. I. M. The quantitative significance of Syntrophaceae and syntrophic partnerships in methanogenic degradation of crude oil alkanes. Environ. Microbiol. 2011, 13, 2957−2975. (31) Dojka, M. A.; Hugenholtz, P.; Haack, S. K.; Pace, N. R. Microbial diversity in a hydrocarbon- and chlorinated-solvent- contaminated aquifer undergoing intrinsic bioremediation. Appl. Environ. Microbiol. 1998, 64, 3869−3877. (32) Winderl, C.; Anneser, B.; Griebler, C.; Meckenstock, R. U.; Lueders, T. Depth-resolved quantification of anaerobic toluene degraders and aquifer microbial community patterns in distinct redox zones of a tar oil contaminant plume. Appl. Environ. Microbiol. 2008, 74, 792−801. (33) Winderl, C.; Penning, H.; Netzer, F. V.; Meckenstock, R. U.; Lueders, T. DNA-SIP identifies sulfate-reducing Clostridia as important toluene degraders in tar-oil-contaminated aquifer sediment. ISME J. 2010, 4, 1314−1325. (34) Bunge, M.; Wagner, A.; Fischer, M.; Andreesen, J. R.; Lechner, U. Enrichment of a dioxin-dehalogenating Dehalococcoides species in twoliquid phase cultures. Environ. Microbiol. 2008, 10, 2670−2683. (35) Sung, Y.; Ritalahti, K. M.; Sanford, R. A.; Urbance, J. W.; Flynn, S. J.; Tiedje, J. M.; Löffler, F. E. Characterization of two tetrachloroethenereducing, acetate-oxidizing anaerobic bacteria and their description as Desulf uromonas michiganensis sp. nov. Appl. Environ. Microbiol. 2003, 69, 2964−2974. (36) Bedard, D. L.; Bailey, J. J.; Reiss, B. L.; Van Slyke Jerzak, G. Development and characterization of stable sediment-free anaerobic bacterial enrichment cultures that dechlorinate Aroclor 1260. Appl. Environ. Microbiol. 2006, 72, 2460−2470. (37) Purdy, K. J.; Munson, M. A.; Nedwell, D. B.; Embley, T. M. Comparison of the molecular diversity of the methanogenic community at the brackish and marine ends of a UK estuary. FEMS Microbiol. Ecol. 2002, 39, 17−21. (38) Dorador, C.; Vila, I.; Remonsellez, F.; Imhoff, J. F.; Witzel, K. Unique clusters of Archaea in Salar de Huasco, an athalassohaline evaporitic basin of the Chilean Altiplano. FEMS Microbiol. Ecol. 2010, 73, 291−302. (39) Raskin, L.; Rittmann, B. E.; Stahl, D. A. Competition and coexistence of sulfate-reducing and methanogenic populations in anaerobic biofilms. Appl. Environ. Microbiol. 1996, 62, 3847−3857. (40) Tasaki, M.; Kamagata, Y.; Nakamura, K.; Mikami, E. Isolation and characterization of a thermophilic benzoate-degrading, sulfate-reducing bacterium, Desulfotomaculum thermobenzoicum sp. nov. Arch. Microbiol. 1991, 155, 348−352. (41) Kuever, J.; Rainey, F. A.; Hippe, H. Description of Desulfotomaculum sp. Groll as Desulfotomaculum gibsoniae sp. nov. Int. J. Syst. Bacteriol. 1999, 49, 1801−1808. (42) Imachi, H.; Sekiguchi, Y.; Kamagata, Y.; Loy, A.; Qiu, Y. L.; Hugenholtz, P.; Kimura, N.; Wagner, M.; Ohashi, A.; Harada, H. Non9809

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810

Environmental Science & Technology

Article

sulfate-reducing, syntrophic bacteria affiliated with Desulfotomaculum cluster I are widely distributed in methanogenic environments. Appl. Environ. Microbiol. 2006, 72, 2080−2091. (43) Schink, B.; Stams, A. J. M. Syntrophism among prokaryotes. In The prokaryotes: Ecophysiology and biochemistry; Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K.-H., Stackebrandt, E., Eds.; SpringerVerlag: New York, 2006; pp 309−335. (44) Jackson, B. E.; Bhupathiraju, V. K.; Tanner, R. S.; Woese, C. R.; McInerney, M. J. Syntrophus aciditrophicus sp. nov., a new anaerobic bacterium that degrades fatty acids and benzoate in syntrophic association with hydrogen- using microorganisms. Arch. Microbiol. 1999, 171, 107−114. (45) Chauhan, A.; Ogram, A. Phylogeny of acetate-utilizing microorganisms in soils along a nutrient gradient in the Florida everglades. Appl. Environ. Microbiol. 2006, 72, 6837−6840. (46) Inagaki, F.; Nunoura, T.; Nakagawa, S.; Teske, A.; Lever, M.; Lauer, A.; Suzuki, M.; Takai, K.; et al. Biogeographical distribution and diversity of microbes in methane hydrate-bearing deep marine sediments on the Pacific Ocean Margin. Proc. Natl. Acad. Sci. 2006, 103, 2815−2820. (47) Sekiguchi, Y.; Takahashi, H.; Kamagata, Y.; Ohashi, A.; Harada, H. In situ detection, isolation, and physiological properties of a thin filamentous microorganism abundant in methanogenic granular sludges: A novel isolate affiliated with a clone cluster, the green nonsulfur bacteria, subdivision I. Appl. Environ. Microbiol. 2001, 67, 5740− 5749. (48) Eriksson, S.; Ankner, T.; Abrahamsson, K.; Hallbeck, L. Propylphenols are metabolites in the anaerobic biodegradation of propylbenzene under iron-reducing conditions. Biodegradation 2005, 16, 253−263. (49) Jones, D. M.; Head, I. M.; Gray, N. D.; Adams, J. J.; Rowan, A. K.; Aitken, C. M.; Bennett, B.; Huang, H.; Brown, A.; Bowler, B. F. J.; Oldenburg, T.; Erdmann, M.; Larter, S. R. Crude oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 2008, 451, 176−180. (50) Kotsyurbenko, O. R.; Friedrich, M. W.; Simankova, M. V.; Nozhevnikova, A. N.; Golyshin, P. N.; Timmis, K. N.; Conrad, R. Shift from acetoclastic to H2-dependent methanogenesis in a west Siberian peat bog at low pH values and isolation of an acidophilic Methanobacterium strain. Appl. Environ. Microbiol. 2007, 73, 2344−2348. (51) Hoj, L.; Olsen, R. A.; Torsvik, V. L. Effect of temperature on the diversity and community structure of known methanogenic groups and other archaea in high arctic peat. ISME J. 2008, 2, 37−48. (52) Galand, P. E.; Fritze, H.; Conrad, R.; Yrjala, K. Pathways for Methanogenesis and diversity of methanogenic archaea in three boreal peatland ecosystems. Appl. Environ. Microbiol. 2005, 71, 2195−2198.

9810

dx.doi.org/10.1021/es302202c | Environ. Sci. Technol. 2012, 46, 9802−9810