Anaerobic Biodegradation of Longer-Chain - ACS Publications

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Anaerobic Biodegradation of Longer-Chain n-Alkanes Coupled to Methane Production in Oil Sands Tailings Tariq Siddique,*,† Tara Penner,‡ Kathleen Semple,§ and Julia M. Foght*,§ †

Department of Renewable Resources, University of Alberta, Edmonton, Alberta T6G 2E3, Canada Syncrude Canada Ltd. Research and Development, Edmonton, Alberta T6N 1H4, Canada § Department of Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9, Canada ‡

bS Supporting Information ABSTRACT: Extraction of bitumen from mined oil sands ores produces enormous volumes of tailings that are stored in settling basins (current inventory g840 million m3). Our previous studies revealed that certain hydrocarbons (shortchain n-alkanes [C6
C10] and monoaromatics [toluene, o-xylene, m-xylene]) in residual naphtha entrained in the tailings are biodegraded to CH4 by a consortium of microorganisms. Here we show that higher molecular weight n-alkanes (C14, C16, and C18) are also degraded under methanogenic conditions in oil sands tailings, albeit after a lengthy lag (∼180 d) before the onset of methanogenesis. Gas chromatographic analyses showed that the longer-chain nalkanes each added at ∼400 mg L
1 were completely degraded by the resident microorganisms within ∼440 d at ∼20 °C. 16S rRNA gene sequence analysis of clone libraries implied that the predominant pathway of longer-chain n-alkane metabolism in tailings is through syntrophic oxidation of n-alkanes coupled with CO2 reduction to CH4. These studies demonstrating methanogenic biodegradation of longer-chain n-alkanes by microbes native to oil sands tailings may be important for effective management of tailings and greenhouse gas emissions from tailings ponds.

’ INTRODUCTION Oil sands tailings, a slurry of slightly alkaline water, sand, silt, clay, and residual hydrocarbons, are byproducts of bitumen extraction from surface mining and processing of oil sands ores.1 Vast oil sands operations in northern Alberta, Canada produce ∼1.31 million barrels of bitumen and generate ∼262,000 m3 of tailings per day (http://www.energy.gov.ab.ca) that are deposited into settling basins (tailings ponds). Tailings accumulate because the producing companies operate under a zero discharge policy. Currently, more than 170 km2 in the oil sands region are covered by tailings ponds containing ∼840 million m3 of fine tailings (http://www.ercb.ca). Mildred Lake Settling Basin (MLSB), the largest tailings pond operated by Syncrude Canada Ltd., currently contains >400 million m3 of fine tailings.2 Challenges associated with tailings ponds include the presence of inorganic and organic contaminants (metals, salts, petroleum hydrocarbons, naphthenic acids, etc.), emission of biogenic greenhouse gases (CH4 and CO2), and very slow consolidation (settling) of fine tailings solids (which segregate from the sand component after deposition). The fine tailings settle by gravity most rapidly during the first 3
4 years after deposition, forming mature fine tailings (MFT; also called fluid fine tailings, FFT) which can then require decades for significant incremental settling. Additional challenges faced by the oil sands r 2011 American Chemical Society

industry include dewatering of MFT (i.e., recovery of water in the tailings slurry for reuse in processing), reducing the stored tailings volumes, and using the consolidated MFT in landscape reconstruction. Biogenic methane emissions from MLSB alone have been estimated at 43,000 m3 day
1.3 Paradoxically, rather than interfering with gravitational settling of MFT, methane production is associated with accelerated settling of tailings solids both in situ and in the laboratory.4 To understand and possibly mitigate methane emissions and to provide a scientific rationale for engineered tailings management, long-term laboratory experiments have been conducted by incubating MFT with hydrocarbons known to be present in the tailings ponds. MFT contains unrecovered bitumen (∼5 wt %) and a small proportion (e0.5 wt %) of fugitive solvent used in bitumen froth treatment such as naphtha, a mixture of aliphatic and aromatic hydrocarbons in the range of C3
C14.5 We previously reported that specific components of the residual naphtha that enters MLSB with fresh tailings were metabolized by the indigenous microbial Received: February 24, 2011 Accepted: May 24, 2011 Revised: May 16, 2011 Published: June 06, 2011 5892

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Environmental Science & Technology communities to CH4.6 Paraffinic (nC6
C10) and aromatic (toluene and xylene isomers) components of naphtha were biodegraded during one year of incubation, but the branched (isoparaffins) and cyclic (naphthenes) aliphatics remained undegraded after one year.5,6 The mass of bitumen-associated n-alkanes entering MLSB can be roughly estimated by using the following values: acyclic alkanes of length nC6, and we wished to determine whether a similar pattern of utilization also pertained to longerchain alkanes (nC14, nC16, and nC18). Finally, a predictive model developed to estimate CH4 flux from MLSB8 was based on anaerobic biodegradation of short-chain n-alkanes (nC6-nC10) and monoaromatic (BTEX) constituents of residual naphtha in the tailings. Because longer-chain alkanes contributed by unextracted bitumen entering the ponds may also be a source of CH4 in MFT, it was necessary to examine the potential contribution of such n-alkanes to tailings pond CH4 emissions. The microbial community in MLSB MFT is now shown to be capable of methanogenic biodegradation of longer-chain alkanes as well as short-chain alkanes.5,6 In contrast, Zengler et al.18 observed methanogenic degradation of nC16 incubated with anoxic ditch sediment after a lag phase of 4 months but did not observe methanogenesis from the short-chain alkanes nC6 or nC10. Long acclimation phases were observed before significant CH4 production occurred from longer-chain alkanes, with the 3-alkane culture enduring a shorter lag phase (∼180 d) than the 2-alkane culture (∼280 d). These results contrast with our previous study5,6 in which the same source of MFT incubated with short-chain alkanes (nC6
C10) consistently produced CH4 after nC8 > nC7 > nC6. However, the overall degradation of longer-chain n-alkanes in MFT agrees with the recent reports of methanogenic degradation of crude oil containing longer-chain alkanes. Townsend et al.19 reported consumption of the n-alkane fraction (C13
C34) of a weathered Alaskan North Slope crude oil by anoxic aquifer samples incubated under methanogenic conditions in laboratory microcosms. Similarly, Jones et al.20 incubated North Sea crude oil with river sediment under methanogenic conditions for 686 d and observed loss of C7
C34 n-alkanes with corresponding CH4 production, and Gieg et al.21 documented methanogenic conversion of crude oil alkanes by an enrichment culture derived from gas condensatecontaminated subsurface sediments.

In the present study, experimental production of CH4 during 438 d incubation approached 80
84% of the theoretical maximum CH4 predicted by stoichiometry. The difference between measured and predicted CH4 values might be accounted for by carbon assimilation into microbial biomass and inefficiency of interspecies H2 transfer in a diverse consortium of anaerobic microorganisms in the MFT. The results support our earlier calculations that 77
79% of predicted CH4 was produced from short-chain n-alkanes,5 whereas Zengler et al.18 observed only 64% of predicted CH4 from metabolism of nC16 in an enrichment culture. Although efforts were made to construct the 16S rRNA gene clone libraries by independently extracting and amplifying replicate subsamples of cultures, interpretation of the clone libraries is speculative because they were derived from single cultures. However, the general community compositions detected are consistent with metagenomic 16S rRNA gene sequences from analogous cultures (unpublished data) and clone library analysis of uncultivated MFT.22 Presumptive identification of sequenced clones pointed to taxa that may be important for anaerobic degradation of longer-chain alkanes, inferred from increased clone abundance in the 3-alkane culture versus the viable baseline control clone libraries. For example, the proportions of clones affiliated with Deltaproteobacteria and unclassified Bacteria significantly increased after incubation with n-alkanes. Of the nine OTUs affiliated with the Deltaproteobacteria, all were presumptively identified as Syntrophus spp., and the dominant sequence (EU522631) was closely related to a Syntrophus sp. associated with methanogenic degradation of hexadecane in an enrichment culture.18 The archaeal clone library constructed from the 3-alkane culture exhibited an increased proportion of hydrogenotrophic methanogens and decreased acetoclastic 5897

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Environmental Science & Technology methanogens versus the baseline control where sequences affiliated with acetoclastic methanogens dominated the clone library. The dominant sequence (EU522625) associated with the hydrogenotrophic Methanomicrobiales was most closely related to methanogens in the brackish and marine ends of an estuary.23 The dominant cloned sequence (EU522627) affiliated with the Methanosarcinales was closely related to acetoclastic Methanosaeta sp. associated with longer-chain alkane degradation.18 The dominance of Syntrophus spp. in the 3-alkane culture supports their proposed role in the activation and fermentation of n-alkanes to acetate and H2 or further acetate conversion to CO2 and H2 as suggested by other recent reports.24,25 Extrapolating the current community structure results from this study suggests that the principal pathway of longer-chain n-alkane metabolism is through syntrophic oxidation of n-alkanes to acetate and H2 mediated by Syntrophus spp., likely followed by CO2 reduction with H2 utilization to CH4 (hydrogenotrophic methanogenesis). This postulated pathway is consistent with the MADCOR process proposed by Jones et al.20 for methanogenic degradation of n-alkanes in crude oil. Using isotopic analysis of CO2 and CH4 produced in degraded oils from the Peace River Oil Sands area of western Canada and laboratory microcosm data, they estimated that 75
92% of methanogenesis in situ occurred through CO2 reduction after syntrophic oxidation of nalkanes by Syntrophus spp. However, because half of the archaeal clone library in our study comprised clones affiliated with acetoclastic methanogens even after incubation with n-alkanes, we cannot discount the potential role of the acetoclastic pathway in methanogenic MFT cultures. This important pathway is supported by the findings of Zengler et al.18 who proposed that acetoclastic CH4 production from nC16 degradation was the dominant pathway in an enrichment culture derived from anoxic sediment. A recent review of anaerobic alkane degradation9 cited studies supporting both hydrogenotrophic and acetoclastic methanogenic degradation of n-alkanes and concluded that the dominant methanogenic pathway was likely dictated by the environment and the provenance of the microbial community. The current study shows that the microbial community in oil sands tailings is capable of utilizing longer-chain n-alkanes under methanogenic conditions. This activity may affect the prediction of CH4 flux from a kinetic model,8 depending on the input of longer-chain alkanes into tailings ponds with unrecovered bitumen. More broadly, the results contribute to knowledge about the potential for methanogenic degradation of longer-chain nalkanes present in other anaerobic environments, such as hydrocarbon-contaminated aquifers and petroleum reservoirs.

’ ASSOCIATED CONTENT

bS

Supporting Information. Tables 1S-3S. 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.) and julia.foght@ ualberta.ca (J.M.F.).

’ ACKNOWLEDGMENT The authors gratefully acknowledge funding from NSERC (PostDoctoral Fellowship to T.S.; Discovery Grants to J.M.F. and T.S.)

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and Syncrude Canada Ltd. for providing tailings. We also thank Jonathan L. Klassen for technical assistance.

’ REFERENCES (1) Schramm, L. L.; Stasiuk, E. N.; MacKinnon, M. Surfactants in Athabasca oil sands slurry conditioning, flotation recovery, and tailings processes. In Surfactants, Fundamentals, and Applications in the Petroleum Industry; Schramm, L. L., Ed.; Cambridge University Press: Cambridge, UK, 2000; pp 365
430. (2) Syncrude Canada Limited. Annual tailings plan submission, Syncrude: Mildred Lake. Submitted to Energy Resources Conservation Board, Alberta. 2010; pp 5
15. (3) 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. (4) 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. (5) Siddique, T.; Fedorak, P. M.; Foght, J. M. Biodegradation of short-chain n-alkanes in oil sands tailings under methanogenic conditions. Environ. Sci. Technol. 2006, 40, 5459–5464. (6) 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. (7) Strausz, O. P.; Morales-Izquierdo, A.; Kazmi, N.; Montgomery, D. S.; Payzant, J. D.; Safarik, I.; Murgich, J. Chemical composition of Athabasca bitumen: the saturate fraction. Energy Fuels 2010, 24, 5053–5072. (8) 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. (9) Mbadinga, S. M.; Wang, L.-Y.; Zhou, L.; Liu, J.-F.; Gu, J.-D.; Mu, B.-Z. Microbial communities involved in anaerobic degradation of alkanes. Int. Biodeterior. Biodegrad. 2011, 65, 1–13. (10) Roberts, D. J. Methods for assessing anaerobic biodegradation potential. In Manual of Environmental Microbiology, 2nd ed.; Hurst, C. J., Crawford, R. L., Knudson, G. R., McInerney, M. J., Stetzenbach, L. D., Eds.; ASM Press: Washington, DC, 2002; pp 1008
1017. (11) 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. (12) 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. (13) DeLong, E. F. Archaea in costal marine environments. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 5685–5689. (14) Cheng, S. M.; Foght, J. M. Cultivation-independent and
dependent characterization of Bacteria resident beneath John Evans Glacier. FEMS Microbiol. Ecol. 2007, 59, 318–330. (15) 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. (16) 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. (17) Benson, D. A.; Karsch-Mizrachi, I.; Lipman, D. J.; Ostell, J.; Wheeler, D. L. GenBank. Nucleic Acids Res. 2007, 35, D21–D25. (18) Zengler, K.; Richnow, H. H.; Rossello-Mora, R.; Michaelis, W.; Widdel, F. Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 1999, 401, 266–269. (19) Townsend, G. T.; Prince, R. C.; Suflita, J. M. Anaerobic oxidation of crude oil hydrocarbons by the resident microorganisms 5898

dx.doi.org/10.1021/es200649t |Environ. Sci. Technol. 2011, 45, 5892–5899

Environmental Science & Technology

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of a contaminated anoxic aquifer. Environ. Sci. Technol. 2003, 37, 5213–5218. (20) 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. (21) 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. (22) Penner, T. J.; Foght, J. M. Mature fine tailings from oil sands processing harbor diverse methanogenic communities. Can. J. Microbiol. 2010, 56, 459–470. (23) 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. (24) Gray, N. D.; Sherry, A.; Hubert, C.; Dolfing, J.; Head, I. M. Methanogenic degradation of petroleum hydrocarbons in subsurface environments: Remediation, heavy oil formation and energy recovery. Adv. Appl. Microbiol. 2010, 72, 137–161. (25) Head, I. M.; Larter, S. R.; Gray, N. D.; Sherry, A.; Adams, J. J.; Aitken, C. M.; Jones, D. M.; Rowan, A. K.; Huang, H.; Roling, W. F. M. Hydrocarbon degradation in petroleum reservoirs. In Handbook of Hydrocarbon and Lipid Microbiology; Timmis, K. N., Ed.; SpringerVerlag: Berlin, Heidelberg, 2010; pp 3097
3109.

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