Environ. Sci. Technol. 2006, 40, 5459-5464
Biodegradation of Short-Chain n-Alkanes in Oil Sands Tailings under Methanogenic Conditions TARIQ SIDDIQUE, PHILLIP M. FEDORAK, AND JULIA M. FOGHT* Department of Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9, Canada
The biodegradation of a mixture of low molecular weight n-alkanes (C6, C7, C8, and C10) was assessed under methanogenic conditions using mature fine tailings (MFT) produced by the oil sands industry in Alberta, Canada. Microorganisms present in the MFT mineralized the added n-alkane mixture, producing 16.2 ((0.3) or 20.5 ((0.1) mmol of methane in the headspace of microcosms spiked with 0.2% or 0.5% w/v n-alkanes, respectively, during 29 weeks of incubation. The spiked n-alkanes biodegraded in the sequence C10 > C8 > C7 > C6. Degradation of 100% C10, 97% C8, 74% C7, and 44% C6 occurred in a mixture of n-alkanes in the MFT spiked at 0.2% after 25 weeks of incubation. The same pattern of biodegradation was also observed in the MFT spiked with 0.5% n-alkanes. Stoichiometric calculations confirmed the mineralization of the degraded n-alkanes to methane. This study showed that the short-chain n-alkanes, which comprise a significant portion of the unrecovered naphtha used in bitumen extraction and released into the settling basins, can be biodegraded into methane. These findings may influence decisions regarding extraction processes and longterm management of MFT, and they suggest that intrinsic, methanogenic metabolism of these n-alkanes may occur in other anoxic environments.
Introduction Enormous volumes of tailings are produced during the recovery of bitumen from Alberta oils sands. In 2005, total production of crude bitumen reached 1 million barrels per day, accounting for 50% of Canadian oil production (Canadian Association of Petroleum Producers, http://en.wikipedia.org/wiki/Athabasca•Tar•Sands), and it is estimated that by 2015 Canadian oil production may reach 4 million barrels per day. With the extraction of 1 m3 of oil sands, about 4 m3 of tailings waste comprising a slurry of alkaline water, sand, silt, clay, and bitumen is produced (1). Oil sands tailings produced by Syncrude Canada Ltd. contain not only residual bitumen but also a fraction of the organic diluent used in oil extraction processes. The diluents may be naphtha (used by Syncrude) or a mixture of pentanes and hexanes (C5 and C6) (used by Albian Sands Energy, Inc.). After bitumen extraction, tailings are pumped into tailings ponds or settling basins where the sand fraction settles, and most of the aqueous slurry of fines (silts and clays plus residual hydrocarbon) slowly densifies to a suspension called mature fine tailings (MFT). It was estimated that 315 000 tonnes of tailings * Corresponding author phone: (780) 492-3279; fax: (780) 4929234; e-mail:
[email protected]. 10.1021/es060993m CCC: $33.50 Published on Web 08/03/2006
2006 American Chemical Society
per day were pumped into the Mildred Lake Settling Basin (MLSB), Fort McMurry, Alberta, by Syncrude (2) over a period of more than 25 years. Biodegradation of the residual hydrocarbons and densification of MFT are important factors in the long-term management of oil sands tailings. Recently, MLSB, which was a primary tailings pond for Syncrude, began to evolve methane gas, releasing an estimated 108 L of methane per day (3). Because MLSB first became methanogenic in the area receiving diluent-enriched tailings, we hypothesized that a suite of anaerobic bacteria in MLSB utilize the lighter hydrocarbons from the diluent (naphtha) used in the extraction process and thereby support methanogenesis by providing hydrocarbon metabolites to the methanogenic consortium. Recently, Fedorak et al. (1) reported an unexpected increase in the rate of densification of MFT accompanying the microbially mediated production of methane (CH4). Therefore, it is important to determine the source of biogenic gases evolved from MFT because this may lead to engineered options to enhance densification, reduce MFT inventories, and improve reclamation options. Assessing and understanding the role of naphtha inputs on maintaining methanogenesis would be useful in developing strategies for future tailings management decisions. The naphtha used by Syncrude Ltd. Canada (CAS No. 64742-49-0) is a mixture of aliphatic and aromatic compounds (C3-C14) containing a significant portion of n-alkanes (heptane (C7), 1-5% by wt; octane (C8), 5-10% by wt; nonane (C9), 1-5% by wt) in addition to benzene, toluene, ethylbenzene, and xylenes (BTEX) compounds. Biodegradability of alkanes has been studied extensively under aerobic conditions (4-9), but less is known about their biodegradation in anoxic environments. Rueter et al. (10) demonstrated that hydrocarbons in crude oil were used directly by sulfatereducing bacteria growing under strictly anoxic conditions. A moderately thermophilic pure culture isolated from the sediment of the Guaymas Basin (Gulf of California, Mexico) utilized alkanes in oil during sulfate reduction (10). Massias et al. (11) studied in situ anaerobic degradation of petroleum alkanes in marine sediments and reported significant (>50%) depletion of C17, C18, and C30 after 24 months of incubation. There are other reports of anaerobic alkane degradation under sulfate-reducing (12-14) and denitrifying, iron-reducing, and methanogenic conditions (12, 15-17), but attention has been focused on monitoring the degradation of longchain n-alkanes (gC12). In most cases, the fate of a single compound has been studied as a model to elucidate the metabolic pathway, and if more than one compound was used, then biodegradation of each compound was assessed in individual microcosms (12) rather than investigating their biodegradation in a mixture of compounds, which would be more realistic. In the present study, anaerobic degradation of low molecular weight n-alkanes (C6, C7, C8, and C10) added collectively to the MFT was assessed for its contribution to methanogenesis. Stoichiometric calculations were performed to evaluate the complete mineralization of the added n-alkanes to methane. We observed a unique pattern of degradation among the spiked n-alkanes. This is the first demonstration that short-chain n-alkanes can support methanogenesis in MFT and thereby contribute to the CH4 flux from the oil sands tailings ponds.
Experimental Section Chemicals. n-Octane (C8; >99% pure) and n-decane (C10; >99% pure) were purchased from Sigma-Aldrich, Oakville, VOL. 40, NO. 17, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
5459
TABLE 1. Characteristics of Mature Fine Tailings Used in the Experiment General Characteristics solids (% by weight) bitumen (% by weight) naphtha (% by weight) pH conductivity (µS cm-1) alkalinity (as ppm of CaCO3) C8-C10 C10-C12 C12-C16 C16-C21
39.5 4.4 0.4 7.8 4200 1570
Aliphatic Hydrocarbons (mg kg-1 (Dry Weight))
BTEX (mg kg-1 (Dry Weight)) benzene toluene ethylbenzene o-, m- and p- xylenes
160 480 3400 6000 0.26 0.35 13 12
Polycyclic Aromatic Compounds (mg kg-1 (Dry Weight)) naphthalene 99% pure) was from Caledon Laboratories Ltd., Georgetown, Ontario, Canada. n-Hexane (C6; >99% pure), methanol (HPLC grade), and other chemicals (analytical reagent grade) were purchased from Fisher Scientific, Ontario, Canada. Description of the Mature Fine Tailings. Fresh MFT (a slurry with 61% moisture content) was collected by Syncrude Canada Ltd. from MLSB at 6 m depth in July 2005. EnviroTest Laboratories, Edmonton, Canada, analyzed the MFT (Table 1) for aliphatic hydrocarbons (Canadian Council of Ministers of the Environment, 2001; http://www.ccme.ca/assets/ pdf/final_phc_method_rvsd_e.pdf), BTEX compounds (EPA 5030/8260-P&T GC-MS; http://www.epa.gov/epahome/ index/), and polycyclic aromatic hydrocarbons (PAHs) and alkylated PAHs (EPA 3540/8270-GC/MS; http://www. epa.gov/epahome/index/). The MFT was stored at 4 °C for use in the experiments. Mineralization of Added n-Alkanes (C6, C7, C8, and C10). This experiment was conducted in 158 mL microcosms with 50 mL of MFT and 50 mL of methanogenic medium, sealed with butyl rubber stoppers. Methanogenic medium was prepared using inorganic salts, vitamins, a redox indicator (resazurin), and a reducing agent (sulfide), as described by Fedorak and Hrudey (18).The headspace in the microcosms was flushed with O2-free 30% CO2 balance N2, at atmospheric pressure. Before being amended with any carbon source 5460
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 17, 2006
(hydrocarbons or acetate), microcosms were preincubated at 22 °C for 2 weeks in the dark to allow for the activation and growth of a methanogenic community and the reduction of alternative electron acceptors in the MFT (19). Each microcosm was then flushed with O2-free 30% CO2 balance N2 gas to remove any CH4 produced during the preincubation period. A mixture of n-alkanes (n-hexane (C6) 2.89 mL, specific gravity 0.66; n-heptane (C7) 2.74 mL, specifiic gravity 0.68; n-octane (C8) 2.72 mL, specific gravity 0.7; n-decane (C10) 2.58 mL, specific gravity 0.73) was prepared, and 0.29 or 0.72 mL of the mixture was then added to the microcosms to give final concentrations of 0.2% or 0.5% w/v of total n-alkanes, respectively. The microcosms were prepared in triplicate. In addition to these treatments, sodium acetate (∼1400 mg L-1) was added to some microcosms containing 0.5% n-alkanes. Parallel heat-killed sterilized controls were prepared in the same manner as the experimental cultures and autoclaved four times on four consecutive days. Two types of viable controls were also included: a baseline control consisting of unamended MFT to account for any methane production from indigenous substrates in the MFT and a positive control amended with acetate but no alkanes. All microcosms were incubated at 22 °C (ca. in situ temperature in MLSB near the source of the MFT) in the dark without shaking. Samples were withdrawn periodically for chemical analyses. Chemical Analyses. Methane production in the microcosms was measured by removing 0.1 mL of headspace from each microcosm and analyzing by gas chromatography with a flame ionization detector (GC-FID) (3). While the methane concentration in the headspace was monitored over time, a measured volume of headspace was removed from the microcosms periodically to release the very high pressure produced by methane production during alkane degradation and replaced by a measured volume of 30% CO2 balance N2, using a sterile needle and syringe. The volumes of gases removed and replenished were accounted for when calculating the total mass of methane produced in the microcosms. For acetate analysis, 0.1 mL samples drawn from the microcosms were centrifuged in a microcentrifuge (Eppendorf 5415D), then 80 µL of the supernatant was mixed with 10 µL of 4 N phosphoric acid plus 10 µL of propionic acid solution (1500 mg L-1) as an internal standard and analyzed by GC-FID (20). Residual alkanes were analyzed using a GC-FID equipped with a purge and trap system. One milliliter of MFT sampled from the experimental microcosms was shaken for 30 min at room temperature with 10 mL of methanol in a 20 mL EPA glass vial capped with a Teflon-coated septum. Vials were stored at 4 °C for 30 min to allow the sediment particles to settle. Two milliliters of supernatant were then transferred to a 44 mL Teflon-sealed EPA glass vial (Fisher Scientific, catalog no. 03-339-14C) and filled completely with deionized water to avoid any headspace. Vials were sonicated for 2 min in a bath sonicator to mix the solution and put on an autosampler for analyses of alkanes (C6-C10) on a HewlettPackard model HP 6890 GC-FID equipped with purge and trap system. The line temperature in the purge and trap system was set at 180 °C. Analytes were desorbed at 225 °C for 4 min and then heated at 225 °C for 10 min before passing to the GC. The capillary column used was a 30 m DB-1 with 0.53 mm internal diameter and 1.50 µm film thickness (J&W Scientific/Agilent Technologies). The front inlet temperature was maintained at 200 °C with a split ratio of 50:1. The column was initially held at 36 °C for 4 min and was then increased at 15.0 °C min-1 to 350 °C. Helium was used as a carrier gas with a flow rate of 7.4 mL min-1. The FID was kept at 250 °C. Stoichiometry of n-Alkane Mineralization. After quantification of the methane and alkane concentrations in the microcosms by GC analysis, theoretical calculations were
FIGURE 1. Methane production in microcosms containing MFT amended with n-alkanes (C6, C7, C8, and C10) with or without acetate, during 29 weeks of incubation at 22 °C: (b) unamended live tailings, ([) acetate-amended tailings, (2) 0.2% C6-C10 spiked tailings, (/) 0.5% C6-C10 spiked tailings, (9) 0.5% C6-C10 + acetate-amended tailings. Symbols represent the mean from analysis of triplicate microcosms, and error bars (where visible) represent 1 standard deviation. Inset shows methane production during the first 10 weeks of incubation. made to compare the actual and predicted values of methane production upon the mineralization of known amounts of n-alkanes. Calculations were based on the following stoichiometric equations derived from the Symons and Buswell equation (21), which describe the complete oxidation of n-alkanes and acetate to CO2 and CH4 under methanogenic conditions
n-hexane
C6H14 + 2.5H2O f 1.25CO2 + 4.75CH4 (1)
n-heptane
C7H16 + 3.0H2O f 1.50CO2 + 5.50CH4 (2)
n-octane
C8H18 + 3.5H2O f 1.75CO2 + 6.25CH4 (3)
n-decane
C10H22 + 4.5H2O f 2.25CO2 + 7.75CH4 (4)
acetate
CH3COO- + H2O f HCO3- + CH4
(5)
Results Methane Production. Methane production was monitored as a quantitative indicator of anaerobic microbial metabolism of n-alkanes in the MFT. Figure 1 shows the time course of cumulative methanogenesis with and without the addition of n-alkanes. After 29 weeks of incubation, amendment with alkanes yielded a high amount of methane, with 16.2 ((0.3) or 20.5 ((0.1) mmol produced in the microcosms spiked with 0.2% or 0.5% w/v n-alkanes, respectively (Figure 1). In the acetate-amended microcosms, 1.9 ((0.03) mmol of methane was observed by 6 weeks of incubation, which slightly increased to 2.8 ((0.02) mmol by week 29. Autoclaved microcosms did not produce any methane (not shown). Only 0.26 ((0.02) mmol of methane was recorded in the live unamended MFT (baseline control) after 29 weeks of incubation. Methane production in all replicate cultures amended with alkanes started within the first week (Figure 1, inset). A slight delay in methane formation was observed in the microcosms spiked with the higher concentration (0.5%) of alkanes compared with methane produced in 0.2% alkane-amended microcosms (Figure 1, inset). Initially, the addition of 0.5% alkanes to acetate-amended microcosms also reduced methane production (Figure 1, inset) with
FIGURE 2. Acetate concentrations in MFT amended with acetate with or without C6-C10 during 10 weeks of incubation: (b) heatkilled MFT, ([) microcosms amended with acetate only, (9) microcosms amended with 0.5% C6-C10 plus acetate. Symbols represent the mean from analysis of triplicate microcosms, and error bars (where visible) represent 1 standard deviation. significantly lower average methane yields (t-test; P ) 0.0024) from acetate plus 0.5% alkane than acetate-amended MFT at week 5. However, the amounts of methane produced in the acetate-amended microcosms were the same by week 9, with or without 0.5% alkanes. After a plateau of 3 weeks, methane started increasing again in the microcosms with 0.5% alkanes plus acetate, and 17.6 ((0.2) mmol of methane was measured after 29 weeks of incubation (Figure 1). Acetate Consumption. Acetate was added (as a positive control) to the MFT with and without n-alkanes to determine the methanogenic potential of MFT. Depletion of acetate over 10 weeks of incubation is shown in Figure 2. In the acetate-amended microcosms that initially contained 1450 ((15) mg acetate L-1, all acetate was consumed during 5 weeks of incubation. The presence of 0.5% alkanes in acetateamended microcosms delayed acetate consumption, with 540 ((48) mg acetate L-1 (ca. 37%) remaining at week 5. Acetate was depleted faster in the microcosms without n-alkanes than in those with n-alkanes. This is consistent with the faster production of methane in the acetateamended microcosms without n-alkanes (Figure 1, inset). No significant acetate depletion was noted in the sterile acetate-amended controls during the incubation period (Figure 2). n-Alkane Biodegradation. The concentrations of residual n-alkanes recovered after 25 weeks of incubation from the heat-killed and live microcosms were used to calculate the percentage biodegradation of individual alkanes in each culture (Figure 3). In all the treatments, no significant change in the concentrations of the added n-alkanes in the MFT could be detected during the first 5 weeks of incubation. n-Decane (C10) was the first alkane to be depleted, with the concentration at 10 weeks dropping from 800 ((14) to 370 ((21) mg L-1 and from 1900 ((220) to 1440 ((140) mg L-1 in the microcosms spiked with 0.2% and 0.5% w/v of total n-alkanes, respectively. Concentrations of other n-alkanes (C6, C7, and C8) were not significantly changed during 10 weeks of incubation. The biodegradation pattern for the n-alkanes became clear with analysis performed after 20 weeks of incubation, when significant degradation of all four n-alkanes (C6, C7, C8, and C10) was observed in live cultures. The higher molecular weight alkanes in the mixture were preferentially metabolized by the microbial populations in the MFT. In the 25 week analysis, no C10 was detected in microcosms spiked with 0.2% n-alkanes whereas a small amount of C8 (25 ( 2.0 mg L-1), 210 ((32) mg L-1 of C7, and 290 ((23) mg L-1 of C6 were detected, compared to VOL. 40, NO. 17, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
5461
TABLE 2. Predicted and Measured Methane Production in Mature Fine Tailings with Degradation of n-Alkanes after 25 Weeks of Incubation
treatment
substrate consumed (mmol)
theoretical CH4 productiona (mmol)
measured CH4 production (mmol)
0.2% C6-C10 C6 ) 0.26 11.3 ((0.2) 12.9 ((0.1) C7 ) 0.48 C8 ) 0.67 C10 ) 0.42 0.5% C6-C10 C6 ) 0.37 18.7 ((0.8) 14.4 ((0.1) C7 ) 0.49 C8 ) 1.16 C10 ) 0.91 0.5% C6-C10 C6 ) 0.33 14.7 ((0.8) 11.6 ((0.2) + acetate C7 ) 0.34 C8 ) 0.49 C10 ) 0.81 acetate ) 1.95
percent of theoretical production 114
77
79
a Based on eqs 1-5 and on GC quantitation of alkane concentrations. Values represent the mean from analysis of triplicate microcosms ((1 standard deviation).
Discussion
FIGURE 3. Biodegradation of n-alkanes (C6, C7, C8, and C10) in microcosms amended with 0.2%, 0.5%, or 0.5% n-alkanes plus acetate during 25 weeks of incubation at 22 °C. Bars represent the mean from analysis of triplicate microcosms, and error bars (where visible) represent 1 standard deviation. concentrations of 760 ((35), 990 ((69), 810 ((56), and 570 ((39) mg L-1, respectively, in heat-killed controls. GC analyses of the heat-killed alkane-amended microcosms revealed that 82-105% of the spiked C6-C10 were still present in the microcosms after 25 weeks of incubation. The same trend of alkane degradation was also observed in the other treatments (0.5% alkanes and 0.5% alkanes plus acetateamended microcosms) (Figure 3), although alkane degradation was delayed in the 0.5% alkane-amended microcosms containing acetate compared with those containing only 0.5% alkanes. Stoichiometries. The mass of individual alkanes consumed (quantified by GC) during incubation was fit into the respective stoichiometric equation (eqs 1-5) to estimate maximum theoretical methane production resulting from complete mineralization of the added n-alkanes. Results are presented in Table 2. In the microcosms spiked with 0.2% n-alkanes, slightly more CH4 was measured in the headspace than was predicted by the stoichiometric equations. In the treatments where a higher concentration of n-alkanes (0.5%) was present, with and without acetate amendment, less methane was measured for 0.5% and 0.5% plus acetateamended microcosms than the predicted values. 5462
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 17, 2006
Although anaerobic biodegradation of hydrocarbons has been reported and is now accepted as a significant process in natural environments (10, 22-23), relatively little is known about the prevalence of this process. We studied the biodegradation of low molecular weight n-alkanes in oil sands tailings under methanogenic conditions. Currently, there are three oil sands extraction plants in northeastern Alberta, and tailings ponds at each site are methanogenic. The onset of methanogenesis in an oil sands tailings pond requires the depletion of sulfate (which originates from oil sands ore during the extraction process) from the tailings, and suitable carbon sources. Holowenko et al. (3) demonstrated that sulfate concentrations in MFT decreased with depth in the MLSB, which equates to age of deposition in the basin. Sulfate-reducing bacteria and methanogens were detected at each sample depth, and the numbers of methanogens often exceeded the numbers of sulfate-reducing bacteria in samples with depleted sulfate concentrations (3). Thus, over time, microbial activity has depleted sulfate, creating conditions that are suitable for methanogenesis. The carbon sources for sulfate reduction or methanogenesis in the tailings ponds were unknown. The major organic material in the oil sands tailing is bitumen which escapes during the extraction process and comprises approximately 2-5% of the weight of MFT (24) and contains insoluble and complex asphaltenes. Due to its high molecular weight, bitumen is unlikely to undergo rapid biodegradation to contribute to methanogenesis in the short term. Holowenko (25) amended MFT samples with increasing concentrations of bitumen and incubated for nearly 250 days under methanogenic conditions but did not detect any enhanced methane production in bitumen-amended microcosms. Table 1 shows substantial amounts of some potential substrates (aliphatic hydrocarbons; C12-C21) in MFT, but their quantities resulted in only a small amount of methane production in unamended microcosms during the incubation period (Figure 1). The biodegradation of various n-alkanes (13, 26-27) and alicyclic hydrocarbons (28) under sulfate-reducing conditions has been demonstrated, and these may also serve as electron donors to drive MFT methanogenesis, as n-hexadecane (16) and the n-alkanes in crude oil (26) can be biodegraded to yield methane. In the present study, biodegradation of a mixture of short-chain n-alkanes in MFT under methanogenic conditions started producing methane in the microcosms
within one week of amendment. In contrast, consortia derived from a marine-estuarine site heavily polluted with petroleum products were incubated under four different anaerobic conditions (12); there was no methanogenic activity in the enrichment culture amended with octane for more than 13 weeks whereas in the decane-spiked culture methanogenic activity began in 3-8 weeks (12). Therefore, we assume that the MFT is already acclimated to utilization of n-alkanes as a methanogenic substrate. Branched and cyclic alkanes, which might also be present in the MFT, would be far less abundant than the spiked n-alkanes. Thus, only the degradation of the n-alkanes was followed in the current study. Substantial methane production in the n-alkane-amended microcosms (Figure 1) indicates that the alkanes were readily metabolized by anaerobes in the MFT. Similar results have been reported in other studies but for longer-chain n-alkanes. For example, in crude-oil-amended methanogenic incubations, endogenous electron donors resulted in evolution of 314 µmol of methane over 475 days of incubation, with the complete removal of the n-alkane fraction of crude oil (C13-C34) in 13 months by the microorganisms from an anoxic aquifer previously contaminated by natural gas condensate (26). Salminen et al. (29) studied the anaerobic biodegradation of mineral oil in boreal subsurface soil and observed methane production with removal of n-alkanes (C11-C15) during mineral oil degradation. In the acetate-amended MFT where n-alkanes were also added, initially methane production was observed with the metabolism of acetate. After the consumption of acetate, no further increase in the methane production was noted for almost 3 weeks, after which methanogenesis resumed, presumably utilizing the added n-alkanes. This rise in methane after a short lag period may be due to a shift in species dominance after acetate exhaustion to permit metabolism of n-alkanes, a less-preferred substrate. Formation of methane by the resident microflora in the MFT is supported by the work of Penner (30) who reported the presence of active microbial populations in MFT and identified various bacterial and archaeal species potentially in syntrophic relationships involved in methanogenesis in MLSB. Methane production due to n-alkane metabolism in the MFT is consistent with the loss of n-alkanes quantified by GC analyses and agrees with the work of Townsend et al. (26) who reported that the n-alkane fraction of crude oil (C13-C34) was consumed under methanogenic conditions in anoxic aquifer samples. Massias et al. (11) studied in situ anaerobic degradation of petroleum alkanes in marine sediments and found a marked decrease in alkanes (eC25) after 24 months. In the present study, it is interesting that the highest molecular weight compound (C10) started disappearing first and supported methanogenesis. The spiked n-alkanes biodegraded selectively depending on the length of their C-chains in the sequence C10 > C8 > C7 > C6. Degradation of 100% C10, 97% C8, 74% C7, and 49% C6 in a mixture of n-alkanes in the MFT spiked at 0.2% during 25 weeks of incubation revealed a definite pattern of biodegradation. This preferential degradation among spiked nalkanes may be attributed either to their octanol/water partition coefficients (Pow) which increase with increasing chain length (Pow for C6 ) 3.9, C7 ) 4.66, C8 ) 5.18, C10 ) 5.98; International Program on Chemical Safety, 2004, http:// www.ilo.org/public/english/protection/safework/cis/products/icsc/dtasht/) or to selective uptake across cell membranes of the n-alkane-degrading microorganisms as proposed by Kim et al. (31). This sequence of biodegradation is in contrast to the sequence observed by Davidova et al. (27) who studied the biodegradation of C6-C12 n-alkanes under sulfate-reducing conditions. They reported rates of degradation that were C6 > C10 > C12. Some studies have shown that
short-chain n-alkanes tend to be removed faster than longerchain n-alkanes, the latter being removed faster than branched hydrocarbons (32-34), but still there is some debate on the exact order in which different compounds are removed during biodegradation. In general, there is good agreement between the measured and the predicted methane yields based on the stoichiometric conversion of the n-alkanes (Table 2). The reason for slightly greater than expected methane production in the microcosms with 0.2% n-alkanes cannot be explained. However, the methane production values (77% and 79% of predicted) from the other two amendments (0.5% n-alkanes and 0.5% n-alkanes plus acetate) agree with findings of earlier investigators. For example, in studies of benzene biodegradation by methanogenic consortia, Kazumi et al. (35) reported 73% of the predicted methane production, and Weiner and Lovley (36) found 80% of the predicted methane production. Of course, eqs 1-5 do not account for carbon assimilation into biomass involved with the biodegradation of substrates. In addition, the methanogenic biodegradation of hydrocarbons depends on a consortium of anaerobic bacteria, and the overall methane yield will depend on the efficiency of interspecies H2 transfer. Two of the oil sands plants use naphtha (containing some short-chain alkanes), and the third plant uses a mixture of C5- and C6-alkanes to recover bitumen from oil sands ores. The results of this study show that the microbial communities in the MFT are capable of utilizing the added C6-C10 n-alkanes under methanogenic conditions and support the hypothesis that components of unrecovered naphtha in oil sands tailings can sustain the methanogenesis in the settling basins. This would explain the evolution of significant volumes of methane from oil sands tailings and has implications for the management of those wastes regarding future densification of MFT and emission of greenhouse gases. We are currently evaluating the contribution of BTEX components in naphtha and of whole naphtha to methanogenesis to expand this observation. Our study extends observations of anaerobic biodegradation of alkanes to the mineralization of a mixture of short-chain n-alkanes under methanogenic conditions. In addition, it is quite conceivable that these compounds may support methanogenesis in other anaerobic environments, such as petroleum-contaminated aquifers and petroleum reservoirs. Finally, this is the first report that describes the sequential methanogenic biodegradation of n-alkanes in order of their decreasing molecular weights.
Acknowledgments The authors gratefully acknowledge funding from the Natural Sciences and Engineering Research Council of Canada (Postdoctoral Fellowship to T.S.), Syncrude Canada Ltd., and Canadian Natural Resources Ltd. We particularly thank Mike MacKinnon (Syncrude) for his assistance in establishing this research program, helpful discussions, and providing samples. Thanks to Jela Burkus (Civil and Environmental Engineering, University of Alberta) for assistance with GC-FID purge and trap analysis and Debbi Coy for technical advice.
Literature Cited (1) 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, 199211. (2) Mikula, R. J.; Kasperski, K. L.; Burns, R.; MacKinnon, M. D. The nature and fate of oil sands fine tailings. In Suspensions: Fundamentals and Applications in the Petroleum Industry; Schram, L. L., Ed.; American Chemical Society, Washington, DC, 1996; pp 677-723. (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. VOL. 40, NO. 17, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
5463
(4) Amouric, A.; Verhe, F.; Auria, A.; Casalot, L. Study of a hexanedegrading consortium in a biofilter and in liquid culture: Biodiversity, kinetics and characterization of degrading strains. FEMS Microbiol. Ecol. 2006, 55, 239-247. (5) Belhaj, A.; Desnoues, N.; Elmerich, C. Alkane biodegradation in Pseudomonas aeruginosa strains isolated from a polluted zone: Identification of alkB and alkB-related genes. Res. Microbiol. 2002, 153, 339-344. (6) Britton, L. N. Microbial degradation of aliphatic hydrocarbons. In Microbial Degradation of Organic Compounds; Gibson, D. T., Ed.; Marcel Dekker: New York, 1984; pp 89-129. (7) Rahman, K. S. M.; Rahman, T. J.; Kourkoutas, Y.; Petsas, I.; Marchant, R.; Banat, I. M. Enhanced bioremediation of n-alkane in petroleum sludge using bacterial consortium amended with rhamnolipid and micronutrients. Bioresour. Technol. 2003, 90, 159-168. (8) Sekine, M.; Tanikawa, S.; Omata, S.; Saito, M.; Fujisawa, T.; Tsukatani, N.; Tajima, T.; Sekigawa, T.; Kosugi, H.; Matsuo, Y.; Nishiko, R.; Imamura, K.; Ito, M.; Narita, H.; Tago, S.; Fujita, N.; Harayama, S. Sequence analysis of three plasmids harboured in Rhodococcus erythropolis strain PR4. Environ. Microbiol. 2006, 8, 334-346. (9) van Beilen, J. B.; Marin, M. M.; Smits, T. H. M.; Rothlisberger, M.; Franchini, A. G.; Witholt, B.; Rojo, F. Characterization of two alkane hydroxylase genes from the marine hydrocarbonoclastic bacterium Alcanivorax borkumensis. Environ. Microbiol. 2004, 6, 264-273. (10) Rueter, P.; Rabus, R.; Wilkes, H.; Aeckersberg, F.; Rainey, F. A.; Jannasch, H. W.; Widdel, F. Anaerobic oxidation of hydrocarbons in crude oil by new types of sulfate-reducing bacteria. Nature 1994, 372, 455-458. (11) Massias, D.; Grossi, V.; Bertrand, J.-C. In situ anaerobic degradation of petroleum alkanes in marine sediments: preliminary results. C. R. Geosci. 2003, 335, 435-439. (12) So, C. M.; Young, L. Y. Anaerobic biodegradation of alkanes by enriched consortia under four different reducing conditions. Environ. Toxicol. Chem. 2001, 20, 473-478. (13) Kropp, K. G.; Davidova, I. A.; Suflita, J. M. Anaerobic oxidation of n-dodecane by an addition reaction in a sulfate-reducing bacterial enrichment culture. Appl. Environ. Microbiol. 2000, 66, 5393-5398. (14) Aeckersberg, F.; Rainey, F. A.; Widdel, F. Growth, natural relationships, cellular fatty acids and metabolic adaptation of sulfate-reducing bacteria that utilize long-chain alkanes under anoxic conditions. Arch Microbiol. 1998, 170, 361-369. (15) Wilkes, H.; Rabus, R.; Fischer, T.; Armstroff, A.; Behrends, A.; Widdel, F.; Anaerobic degradation of n-hexane in a denitrifying bacterium: Further degradation of the initial intermediate (1methylpentyl) succinate via C-skeleton rearrangement. Arch. Microbiol. 2002, 177, 235-243. (16) Anderson, R. T.; Lovley, D. R. Hexadecane decay by methanogensis. Nature 2000, 404, 722-723. (17) 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. (18) Fedorak, P. M.; Hrudey, S. E. The effects of phenol and some alkyl phenolics on batch anaerobic methanogenesis. Water Res. 1984, 18, 361-367. (19) Lueders, T.; Friedrich, M. W. Effects of amendment with ferrihydrite and gypsum on the structure and activity of methanogenic populations in rice field soil. Appl. Environ. Microbiol. 2002, 68, 2484-2494.
5464
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 17, 2006
(20) Fedorak, P. M.; Coy, D. L.; Salloum, M. J.; Dudas, M. J. Methanogenic potential of tailings samples from oil sands extraction plants. Can J. Microbiol. 2002, 48, 21-33. (21) 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. (22) Rabus, R.; Widdel, F. Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Arch. Microbiol. 1995, 163, 96-103. (23) Widdel, F.; Rabus, R. Anaerobic biodegradation of saturated and aromatic hydrocarbons. Curr. Opin. Biotechnol. 2001, 12, 259-276. (24) MacKinnon, M. Development of the tailings pond at Syncrude’s oil sands plant: 1978-1987. AOSTRA J. Res. 1989, 5, 109-133. (25) Holowenko, F. M. Methanogenesis and fine tailings wastes from oil sands extraction: A microcosm-based laboratory study. M.Sc. Thesis, University of Alberta, 2000. (26) Townsend, G. T.; Prince, R. C.; Suflita, J. M. Anaerobic oxidation of crude oil hydrocarbons by the resident microorganisms of a contaminated anoxic aquifer. Environ. Sci. Technol. 2003, 37, 5213-5218. (27) Davidova, I. A.; Gieg, L. M.; Nanny, M.; Kropp, K. G.; Suflita, J. M. Stable isotope studies of n-alkane metabolism by a sulfatereducing bacterial enrichment culture. Appl. Environ. Microbiol. 2005, 71, 8174-8182. (28) Townsend, G. T.; Prince, R. C.; Suflita, J. M. Anaerobic biodegradation of alicyclic constituents of gasoline and natural gas condensate by bacteria from an anoxic aquifer. FEMS Microbiol. Ecol. 2004, 49, 129-135. (29) Salminen, J. M.; Tuomi, P. M.; Suortti, A. M.; Jorgensen, K. S. Potential for aerobic and anaerobic biodegradation of petroleum hydrocarbons in boreal subsurface. Biodegradation 2004, 15, 29-39. (30) Penner, T. J. Molecular analysis of microbial populations in oil sands tailings. M.Sc. Thesis, University of Alberta, 2005. (31) Kim, I. S.; Foght, J. M.; Gray, M. R. Selective transport and accumulation of alkanes by Rhodococcus erythropolis S+14He. Biotechnol. Bioeng. 2002, 80, 650-659. (32) Peters, K. E.; Moldowan. The Biomarker Guide: Interpreting Molecular Fossils in Petroleum and Ancient Sediments; Prentice Hall: Englewood Cliffs, NJ, 1993. (33) Volkman, J. K.; Alexander, R.; Kagi, R. I.; Rowland, S. J.; Sheppard, P. N. Biodegradation of aromatic hydrocarbons in crude oils from the Barrow Sub-basin of Western Australia. Org. Geochem. 1984, 6, 619-632. (34) Wenger, L. M.; Davis, C. L.; Isaksen, G. H. Multiple controls on petroleum biodegradation and impact on oil quality. SPE Reservoir Eval. Eng. 2001, 5, 375-383. (35) Kazumi, J.; Caldwell, M. E.; Suflita, J. M.; Lovley, D. R.; Young, L. Y. Anaerobic degradation of benzene in diverse anoxic environments. Environ. Sci. Technol. 1997, 31, 813-818. (36) Weiner, J. M.; Lovley, D. R. Rapid benzene degradation in methanogenic sediments from a petroleum-contaminated aquifer. Appl. Environ. Microbiol. 1998, 64, 1937-1939.
Received for review April 25, 2006. Revised manuscript received June 28, 2006. Accepted July 5, 2006. ES060993M
