Methanogenic biodegradation of iso-alkanes and cycloalkanes during long-term incubation with oil sands tailings

Environmental Pollution 258 (2020) 113768

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Methanogenic biodegradation of iso-alkanes and cycloalkanes during long-term incubation with oil sands tailings* Tariq Siddique a, *, Kathleen Semple b, Carmen Li c, Julia M. Foght b a

Department of Renewable Resources, University of Alberta, Edmonton, AB T6G 2G7, Canada Department of Biological Sciences, University of Alberta, Edmonton, AB T6G 2E9, Canada c Department of Biological Sciences, University of Calgary, Calgary, AB T2N 1N4, Canada b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 28 June 2019 Received in revised form 7 December 2019 Accepted 7 December 2019 Available online 12 December 2019

Microbes indigenous to oil sands tailings ponds methanogenically biodegrade certain hydrocarbons, including n-alkanes and monoaromatics, whereas other hydrocarbons such as iso- and cycloalkanes are more recalcitrant. We tested the susceptibility of iso- and cycloalkanes to methanogenic biodegradation by incubating them with mature fine tailings (MFT) collected from two depths (6 and 31 m below surface) of a tailings pond, representing different lengths of exposure to hydrocarbons. A mixture of five iso-alkanes and three cycloalkanes was incubated with MFT for 1700 d. Iso-alkanes were completely biodegraded in the order 3-methylhexane > 4-methylheptane > 2-methyloctane > 2-methylheptane, whereas 3-ethylhexane and ethylcyclopentane were only partially depleted and methylcyclohexane and ethylcyclohexane were not degraded during incubation. Pyrosequencing of 16S rRNA genes showed enrichment of Peptococcaceae (Desulfotomaculum) and Smithella in amended cultures with acetoclastic (Methanosaeta) and hydrogenotrophic methanogens (Methanoregula and Methanoculleus). Bioaugmentation of MFT by inoculation with MFT-derived enrichment cultures reduced the lag phase before onset of iso-alkane and cycloalkane degradation. However, the same enrichment culture incubated without MFT exhibited slower biodegradation kinetics and less CH4 production, implying that the MFT solid phase (clay minerals) enhanced methanogenesis. These results help explain and predict continued emissions of CH4 from oil sands tailings repositories in situ. © 2019 Elsevier Ltd. All rights reserved.

Keywords: Anaerobic hydrocarbon biodegradation iso-Alkane Cycloalkane Methanogenesis

1. Introduction Aqueous extraction of bitumen from surface-mined oil sands ores in Alberta, Canada generates huge volumes of fluid fine tailings (FFT) that are deposited in tailings ponds. The current inventory has reached >1.2 billion m3 of fluid tailings (AER, 2019) pending reclamation. At the time of deposition into ‘tailings ponds’, the FFT is
10 wt % solids. Within 2e4 years, typically, these FFT undergo gravitational settling to become an anaerobic colloidal suspension defined as mature fine tailings (MFT) at >30 wt % solids. These MFT comprise slightly alkaline water, silt, clay, unrecovered bitumen (~5 wt %) and residual hydrocarbon diluent (<1 wt %) such as naphtha; the latter represents the small proportion of the light

* This paper has been recommended for acceptance by Charles Wong. * Corresponding author. 348F South Academic Building, University of Alberta, Edmonton, Canada.. E-mail address: [email protected] (T. Siddique).

https://doi.org/10.1016/j.envpol.2019.113768 0269-7491/© 2019 Elsevier Ltd. All rights reserved.

hydrocarbon mixture used during bitumen extraction and recovery from ores that is not recovered from tailings for re-use. All such tailings ponds examined to date support complex communities of indigenous microbes capable of anaerobically biodegrading a range of diluent hydrocarbons to yield greenhouse gases, particularly methane (CH4) and carbon dioxide (CO2) (Foght et al., 2017; Siddique et al., 2018). The largest of the extant ponds, Mildred Lake Settling Basin (MLSB; Syncrude Canada Ltd.), has been estimated to emit ~43e93 million L CH4 day1 (Holowenko et al., 2000; Siddique et al., 2008; Burkus et al., 2014). The hydrocarbons in MFT include unrecovered bitumen (insoluble and complex asphaltenes; ~5% w/w), which is considered resistant to biodegradation (Holowenko et al., 2000) and fugitive process diluent (mixtures of low molecular weight aliphatic and monoaromatic hydrocarbons, primarily in C6eC10 range, specific to each operators; Burkus et al., 2014). The naphtha diluent in MLSB comprises a high proportion of iso-alkanes (31 wt %) plus cycloalkanes (27%), n-alkanes (18%) and monoaromatics (toluene, ethylbenzene and xylenes; 15%) (Siddique et al., 2007). Of these, n-

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alkanes (C6eC10) and some monoaromatic components previously were shown to be biodegraded by Syncrude MFT microbes within 1 year of incubation, but iso- and cycloalkanes were not biodegraded (Siddique et al., 2006; Siddique et al., 2007; Siddique et al., 2012). Subsequent experiments with MLSB-derived enrichment cultures revealed methanogenic biodegradation of some C7eC8 iso-alkanes (Abu Laban et al., 2014) and methylcyclopentane (McyC5; Tan et al., 2015). Experiments incubating MFT from different tailings ponds adapted to different diluents (Canadian Natural Upgrading Ltd. [CNUL] formerly Shell Albian; and Canadian Natural Resources Ltd. [CNRL]) demonstrated methanogenic biodegradation of additional iso-alkanes and some cycloalkanes present in naphtha during longterm incubations of ~1600 d (>4 y) (Mohamad Shahimin and Siddique, 2017a). However, the full range of diluent hydrocarbons susceptible to biodegradation has yet to be established. On the basis of these biodegradation studies, we developed a mathematical model to predict in situ CH4 emissions from MLSB (Siddique et al., 2008) that has been expanded to include recalcitrant hydrocarbons and other oil sands tailings ponds and tailings repositories proposed for reclamation purposes (Kong et al., 2019). Such models are important because they inform regulatory decisions regarding greenhouse gas emissions and therefore may influence tailings pond management practices; also, methanogenic biodegradation of residual hydrocarbons indirectly affects the physical and chemical behavior of MFT in working ponds and in reclamation scenarios. To refine the predictive model, we need to know the biodegradation potential of indigenous microorganisms residing in different strata of a tailing pond and the suite of susceptible hydrocarbons in situ. Given the evidence that long incubation times are needed to observe biodegradation of recalcitrant hydrocarbons, here we used MFT from two strata in MLSB, hypothesizing that the biodegradation potential of microbial communities would differ with age (i.e., duration of exposure to diluent), for which depth in the pond is a proxy. Supporting this hypothesis, previous work revealed that the residual diluent in situ had been progressively and selectively degraded with depth (i.e., time) in MLSB (Fig. S2 in Foght et al., 2017): labile hydrocarbons such as n-alkanes and monoaromatics were degraded first, followed by some iso- and cycloalkanes with others persisting for an unknown timespan. Therefore, here we have used MFT collected from 6 m below the pond surface to represent young tailings deposits, likely <5 years old, and MFT from 31 m deep, likely representing tailings ~30 years old, to assess biodegradation potential for recalcitrant iso-alkanes and cycloalkanes, the predominant diluent fractions remaining in deeper strata, which may resist biodegradation or may eventually be metabolized to greenhouse gases. Recognizing the potential utility of using bioaugmentation to stimulate biodegradation of recalcitrant hydrocarbons for remediation purposes, here we also investigated the effect of inoculating MFT with hydrocarbon-degrading enrichment cultures, also derived from MFT. Cultures were incubated for 1700 d (>4.5 y) while monitoring CH4 production coincident with hydrocarbon biodegradation and microbial community changes. 2. Materials and methods 2.1. Chemicals and materials 3-Methylhexane (hereafter, 3-MC6; Cat #M49801; >99% purity), 3-ethylhexane (3-EC6; Cat #R289396; 99% purity), 2methylheptane (2-MC7; Cat #M47949; 98% purity), 4methylheptane (4-MC7; Cat # 111,023; 99% purity), 2methyloctane (2-MC8; Cat # 68,170; >99% purity), ethylcyclopentane (EcyC5; Cat # 110,752; 98% purity), methylcyclohexane (McyC6; Cat # 300,306; >99% purity) and ethylcyclohexane

(EcyC6; Cat #E19154; >99% purity) were purchased from SigmaAldrich, Oakville, ON, Canada. Two samples of MFT were collected from different depths in the MLSB tailings pond (Syncrude Canada Ltd.) in Alberta, Canada. One MFT sample was collected at 6 m below the pond surface; it had a relatively high concentration of background naphtha diluent (0.4 wt%) (Siddique et al., 2006) and is estimated to have been deposited in the pond
5 years previously. The other MFT sample was collected at 31 m below surface; it was depleted in naphtha diluent (0.01 wt%) and was estimated to have been deposited 30 years ago. The MFT samples were stored at 4  C in the dark in sealed containers until used in the experiment. 2.2. Assessing methanogenic biodegradation of iso-alkanes and cycloalkanes Eight hydrocarbons were selected based on their significant proportions in Syncrude naphtha. The naphtha diluent (C6eC10) comprises n-alkanes (~18%), monoaromatics (~15%), iso-alkanes (~31%), cycloalkanes (~27%) and other hydrocarbons (9%) (Table 1 in Siddique et al., 2007). Of these, we chose five constituent iso-alkanes (3-MC6, 3-EC6, 2-MC7, 4-MC7 and 2-MC8) and three constituent cycloalkanes (EcyC5, McyC6 and EcyC6) for incubation in culture bottles with MFT for 1700 d. A mixture comprising equal masses of these eight hydrocarbons was prepared in a vial sealed with a Teflon liner for amending experimental cultures. Five sets of experiments were established, each consisting of replications of various combinations of live or heat-killed MFT, sterile methanogenic culture medium and/or inoculum (Table S1). The microcosms (158 mL serum bottles) in the set designated MFT6m received 50 mL MFT collected from 6 m depth and 50 mL sterile methanogenic medium (per Siddique et al., 2006). Set MFT-6m-N comprised 50 mL of MFT collected from 6 m depth, 40 mL methanogenic medium and 10 mL inoculum retrieved from a primary methanogenic MFT enrichment culture “N” grown on naphtha diluent (C6eC10; Siddique et al., 2007) for ~270 d at the time of inoculation. Set MFT-6m-P consisted of 40 mL of MFT collected from 6 m depth, 40 mL methanogenic medium and 20 mL inoculum from a primary methanogenic MFT enrichment culture “P”, grown for ~500 d on paraffinic diluent (C5eC6, primarily comprising npentane, n-hexane, isopentane, 2-methylpentane and 3methylpentane; Mohamad Shahimin and Siddique, 2017b). Set MFT-31 m comprised 50 mL of MFT collected from 31 m depth plus 50 mL of methanogenic medium. The final set, No-MFT-P, comprised the inoculum culture “P” (20 mL) plus sterile medium (80 mL) without addition of any MFT. The microcosms were sealed under a headspace of 30% O2-free CO2, balance N2 using procedures described by Siddique et al. (2006). Randomly selected microcosms in each set were autoclaved each day for 4 consecutive days to serve as heat-killed controls. All microcosms were then pre-incubated for 2 weeks in the dark to adapt to culture conditions and reduce the amount of endogenous carbon. After that acclimation period, the headspace in all microcosms was flushed with N2/CO2 to remove any gas produced during the pre-incubation. Immediately thereafter, for each experimental set three treatments were imposed, which were replicated thrice: (1) Unamended treatment where no hydrocarbon mixture was added to the prepared microcosms, serving as a baseline control for methanogenic degradation of endogenous substrates; (2) Amended treatment where the mixture of eight iso-alkanes and cycloalkanes was added via syringe through the Teflon-lined seal (to minimize evaporative losses during handling) to achieve a concentration of 200 mg of each compound L1 total culture volume; and (3) Sterile amended control treatment (using heat-killed cultures) to account for any abiotic loss of the added hydrocarbons and enabling

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calculation of the percent hydrocarbon biodegradation in live cultures. The microcosms were incubated stationary in the dark at room temperature (~20  C) and periodically sampled for gas chromatographic (GC) analyses of headspace CH4 and hydrocarbons as well as molecular (16S rRNA gene) analysis for microbial characterization, as described below. 2.3. Chemical analyses CH4 production was determined by using a sterile needle and syringe to remove 0.1 mL headspace gas from sealed microcosms. These samples were directly injected into a GC equipped with a flame ionization detector (Fedorak et al., 2003). Methanogenic biodegradation of hydrocarbons was quantified by measuring depletion of volatile hydrocarbons in 20e50 mL culture samples of headspace injected directly into an Agilent 5673 GC-mass spectrometry (GC-MS) system equipped with a HP-MS capillary GC column (Tan et al., 2015). Column temperature was maintained at 35  C for 7 min, then increased to 100  C at 10  C min1. The carrier gas was helium at a flow rate of 1.1 mL min1, splitless. The biodegradation of individual compounds was determined by analyzing GC-MS peak area using selected quantification (Q) ions for each compound at specific retention time. We used methylcyclohexane (McyC6), a hydrocarbon in our mixture, as the conserved internal marker because this compound did not degrade over a 5 year period in this study. We confirmed the nondegradability of McyC6 by comparing its peak area with that of 1,1,3-trimethylcyclohexane, a known conserved internal marker (Prince and Douglas, 2005; Townsend et al., 2004), which was present as one of the background hydrocarbons in the 6-m MFT. We calculated the percent remaining hydrocarbons using the following equation:

% Remaining hydrocarbon ¼ ðPARL =PARS Þx100

(1)

where PARL and PARS are the peak area ratios of target hydrocarbon in live and sterile culture bottles, respectively. Peak area ratio (PAR) is calculated using the following equation:

PAR ¼ PATH =PAIM

(2)

where, PATH and PAIM are peak areas of target hydrocarbons and the internal marker, respectively. All calculations are based on the average of three replications per treatment. Residual hydrocarbon concentrations in cultures were quantified using GC-MS analysis as described above, and apparent biodegradation was calculated by difference between live test and heat-treated cultures, thus accounting for any abiotic losses. The theoretical maximum yield of CH4 from hydrocarbon biodegradation was calculated per Mohamad Shahimin and Siddique (2017a) and used in mass balance calculations (Table S2). Any production of CH4 from endogenous substrates in MFT was accounted for by subtracting CH4 measured in live cultures that did not receive hydrocarbon amendments. 2.4. Characterization of microbial communities At intervals during incubation, microbial community structure was assessed by pyrosequencing 16S rRNA genes (Siddique et al., 2014b). In brief, total genomic DNA from each culture sample was extracted from triplicate 300-ml sub-samples (Foght et al., 2004), precipitated and pooled. Each pool was amplified by PCR in triplicate 25-mL reactions using universal primers (An et al., 2013) for Bacteria and Archaea. The following thermocycle program was used: initial 5-min denaturation at 95  C followed by 10 cycles of

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(30 s at 95  C, 30 s at 60  C, decreasing by 0.5  C per cycle, then 30 s at 72  C); 30 cycles of (30 s at 95  C, 30 s at 55  C, and 30 s at 72  C); and a final extension of 5 min at 72  C. The primers and extraction and amplification protocols were developed and validated by An et al. (2013) using a logarithmic-scale mock community consisting of 11 prokaryotic species (Fig. S1 in that publication) and 160 hydrocarbon-impacted environments. Control reactions included DNA-free (reagent-only and PCR reagent-only) negative controls; these were conducted with every set of samples to ensure quality and control during DNA extraction and PCR amplification. Amplification products were examined for quality using agarose gel electrophoresis and those reactions passing scrutiny were purified using a QIAquick (Qiagen) PCR purification kit. Purified products were sequenced (Genome Quebec Innovation Centre; Montreal, QC, Canada) using a GS FLX Titanium Series XLR70 kit (Roche Diagnostics Corporation). Phoenix 2, a SSU rRNA data analysis pipeline that includes quality control and chimera detection, was used to analyze raw pyrosequence data per (Soh et al. (2013)). Pyrotag sequences from samples comprising >2000 to ~10,000 reads per sample were submitted to the NCBI Short Read Archive (http://www.ncbi.nlm.nih.gov/sra) under run numbers SRR617776-91, SRR629423-34 and SRR090182-86. Sequences exceeding quality assurance thresholds were compared to the SILVA 102 database (http://www.arb-silva.de) and clustered into Operational Taxonomic Units (OTUs) at <5% distance. This threshold was selected based on prior results (An et al., 2013) including the observation that many of the bacterial amplicons were affiliated with uncultivated taxa that could be identified only to Family level, and that archaeal sequences were readily identified to genus or species level at this distance. 3. Results 3.1. CH4 production during metabolism of alkanes CH4 production by MFT either unamended or amended with a mixture of iso- and cycloalkanes and either uninoculated or inoculated with MFT-derived enrichment cultures was monitored during 1700 d (~4.7 y) incubation. Amended MFT-6m cultures produced significantly more CH4 than unamended MFT-6m after a lag phase of ~630 d (Fig. 1A). Thereafter, CH4 production increased continuously during incubation, with amended MFT-6m producing ~5.9 ± 0.1 mmol compared to ~3.4 ± 0.3 mmol CH4 in unamended MFT-6m at the last analysis time of 1700 d. A similar lag phase was observed for MFT-31 m, after which amended MFT-31 m yielded a maximum of 3.6 ± 0.3 mmol CH4 versus unamended MFT-31 m (0.03 ± 0.01 mmol) at ~1550 d (Fig. 1B). The difference in CH4 yield between the two MFT samples and between amended and unamended cultures for both MFT samples is interpreted to result from the greater amount of endogenous labile hydrocarbon in the shallower (younger) sample: 6-m MFT has a high background of naphtha (0.4%; Siddique et al., 2006) whereas the deeper sample contains a much lower concentration of biodegraded naphtha (Fig. S2 in Foght et al., 2017). The presence and degradation of nheptane (n-C7), a labile and significant n-alkane hydrocarbon in naphtha diluent, in Fig. 1C, S1C and S1D indicates the presence of endogenous labile naphtha hydrocarbons in 6-m MFT that contributed to greater CH4 production in unamended 6-m MFT (Fig. 1A) compared to unamended 31-m MFT (Fig. 1B). Replicate samples of MFT-6m were inoculated with cultures derived from MFT that had been enriched by growth on either a paraffinic diluent (MFT-6m-P) or a naphtha diluent (MFT-6m-N). Shorter lag phases before onset of CH4 production and slightly faster rates of CH4 production (Figs. S1A and S1B) were observed for both sets of inoculated cultures versus the corresponding

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Fig. 1. Cumulative CH4 production by alkane-amended and unamended (A) MFT-6m or (B) MFT-31 m. Hydrocarbon biodegradation was calculated by reference to parallel sterile MFT controls and expressed as % residual hydrocarbon in alkane-amended (C) MFT-6m or (D) MFT-31 m cultures. No methane was produced in sterile amended controls (not shown). Data points are mean values (n ¼ 3) and error bars, where visible, represent the standard deviation. The abbreviations are: 3-MC6 (3-methylhexane), 3-EC6 (3-ethylhexane), 2-MC7 (2-methylheptane), 4-MC7 (4-methylheptane), 2-MC8 (2-methyloctane), EcyC5 (ethylcyclopentane), McyC6 (methylcyclohexane), EcyC6 (ethylcyclohexane) and n-C7 (nheptane).

uninoculated MFT-6m cultures, with or without amendment (Fig. 1A). Alkane-amended MFT-6m-P produced significantly more CH4 after a lag of 180 d and produced ~5.7 ± 0.2 mmol CH4 compared to ~2.7 ± 0.1 mmol in unamended MFT-6m-P at 1700 d when CH4 was still being produced (Fig. S1A). Alkane-amended MFT-6m-N exhibited a lag phase of ~490 d after which significantly more CH4 was produced (5.3 ± 0.3 mmol at 1700 d) than by the unamended MFT-6m-N (~3.6 ± 0.1 mmol) (Fig. S1B). Amended inoculated medium that did not receive any MFT (No-MFT-P) also produced CH4 (~1.2 ± 0.2 mmol) (Fig. S1A) but less than that in cultures containing MFT. 3.2. Biodegradation of iso- and cycloalkanes Once CH4 production in alkane-amended MFT-6m and MFT31 m cultures significantly exceeded that in the corresponding unamended cultures, concentrations of the eight target alkanes in culture headspace were measured at intervals. Complete biodegradation of the iso-alkanes 3-MC6, 2-MC7, 4-MC7 and 2-MC8 was observed, with only partial depletion of 3-EC6. Among the cycloalkanes, only EcyC5 was partially depleted over 1700 d, leaving McyC6 and EcyC6 completely undegraded (Fig. 1CeD and

Figs. S1CeE). Specifically, in amended MFT-6m cultures three isoalkanes (3-MC6, 4-MC7, and 2-MC8) were completely biodegraded during 1700 d incubation, while ~65% degradation of 2-MC7 was measured. Partial depletion (~30%) of 3-EC6 and EcyC5 also occurred but two cycloalkanes (McyC6 and EcyC6) did not show any degradation during incubation (Fig. 1C). In the amended MFT-31 m, complete biodegradation of four iso-alkanes (3-MC6, 4-MC7, 2-MC8 and 2-MC7) was observed, whereas one iso-alkane (3-EC6) exhibited no degradation during incubation (Fig. 1D). Among cycloalkanes, EcyC5 was 50% degraded but McyC6 and EcyC6 remained undegraded in the amended 31-m MFT (Fig. 1D). Similar patterns of biodegradation susceptibility were observed when 6-m MFT was inoculated with one of two enrichment cultures (P or N). In the amended MFT-6m-P cultures, biodegradation started with a shorter lag and at a slightly faster rate than in uninoculated MFT-6m cultures due to addition of active hydrocarbondegrading microbes to the indigenous microbes. When the first GC analysis was performed at 425 d, >60% of 3-MC6 and 4-MC7 had already been degraded, surpassing the uninoculated MFT-6m cultures (Fig. 1A). By the end of incubation 100% of 3-MC6, 4-MC7, 2MC8 and 2-MC7 were degraded along with 30% of 3-EC6 and 45% of EcyC5 by MFT-6m-P, although no degradation of McyC6 and

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EcyC6 was observed (Fig. S1C). The MFT-6m-N cultures also completely degraded 3-MC6, 4-MC7, 2-MC8 and 2-MC7 plus 35% of 3-EC6. Greater degradation (~65%) of EcyC5 was observed in this culture but McyC6 and EcyC6 showed no measurable degradation during 1700 d incubation (Fig. S1D). Similar biodegradation patterns were observed in the experimental set containing inoculum P without MFT (No-MFT-P), albeit with a slightly longer lag phase and slower degradation rate (Fig. S1E), indicating that inoculation accelerated degradation but did not change the hydrocarbon susceptibility. Phase transfer of carbon from amended hydrocarbons to CH4 through microbial metabolism was determined by first calculating the theoretical maximum CH4 expected from the mass of biodegraded hydrocarbon (the difference between initial and residual hydrocarbon concentrations measured in amended live cultures and sterile control incubations). This value was compared with CH4 measurements in live amended cultures after mathematically subtracting any CH4 produced from endogenous substrates as measured in parallel live, unamended control cultures. The measured CH4 in amended cultures attributable to alkane amendment was 56e73% of the predicted theoretical maximum CH4 at 1700 d, whereas less conversion of hydrocarbons to CH4 (43% of theoretical maximum) was observed in the No-MFT-P set (Table S2). 3.3. Microbial community structure during metabolism of alkanes Samples of amended and unamended cultures were collected at intervals for 16S rRNA gene amplification and sequencing to study community structure during incubation. Community membership was generally similar in all cultures but the proportions of taxa changed with treatment and time. At 120 d, prior to significant CH4 production (Fig. 1A), the bacterial community in MFT-6m comprised only 15% of total prokaryotic reads regardless of alkane amendment (Fig. S2) with the remainder almost exclusively affiliated with methanogenic genera of the Archaea. However, at 815 d, during active alkane degradation (Fig. 1C), bacterial reads in the amended culture had risen to 23% of total reads whereas the unamended MFT-6m bacterial reads had decreased to 10%. The proportions of bacterial taxa in amended and unamended MFT-6m were similar at 120 d, comprising Proteobacteria (~51e53%), Chloroflexi (~13e15%), Firmicutes (~11e14%, including 2e3% Peptococcaeae), Bacteroidetes (~5e6%), Actinobacteria (~3e4%), Spirochaetes (~3e5%) and Synergistetes (<1%). By 815 d of incubation the bacterial communities in amended and unamended MFT-6m had diverged, with the amended cultures experiencing a dramatic increase in Peptococcaceae (62%; 60% Desulfotomaculum alone) versus 18% in unamended MFT-6m (Fig. 2A, Table S3). The proportions of archaeal taxa remained quite stable regardless of amendment and incubation time, except for an increase in Methanomicrobiaceae at 815 d (Fig. 2B). The prokaryotic communities in MFT-31 m cultures differed from those in MFT-6m by having a much greater proportion of bacterial reads. At 182 d and 420 d, before the onset of methanogenesis, Bacteria represented 61e78% of the total pyrosequencing reads in amended and unamended cultures (Fig. S2) versus 10e25% in MFT-6m. Only at 815 d, after alkane biodegradation and CH4 production had begun, did the proportion of bacterial reads in amended MFT-31 m decrease to 28% to resemble that in amended MFT-6m. In contrast, the unamended MFT-31 m cultures maintained dominance of Bacteria throughout the sampling period (Fig. S2). The bacterial community structure was more diverse in MFT-31 m than in MFT-6m cultures. Notably, ‘rare’ OTUs present individually at <1% initially comprised >20% of bacterial reads in unamended MFT-31 m; this proportion remained high in

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Fig. 2. Microbial community structure determined by 16S rRNA gene pyrosequencing of unamended and amended MFT-6m samples during microbial metabolism of isoalkanes and cycloalkanes to CH4. Graphs show proportions of (A) bacterial taxa and (B) archaeal taxa at 120 and 815 d incubation. “Other Bacteria” and “Other Archaea” are the sums of taxa individually present at
1% abundance.

unamended MFT-31 m but decreased with time in amended MFT31 m as presumptive hydrocarbon-degrading OTUs were enriched. Proteobacterial taxa initially dominated the bacterial reads at >70% (Fig. 3A and Table S4) but by 420 d of incubation the bacterial community composition had shifted in the amended MFT-31 m even though neither CH4 production nor the proportion of bacterial:archaeal reads had changed significantly, and despite GC analysis showing onset of biodegradation of only one iso-alkane (3MC6) by that time (Fig. 1D). At 420 d, Firmicutes dominated the bacterial community of amended MFT-31 m by contributing ~47% of bacterial reads (Peptococcaceae, almost exclusively Desulfotomaculum) compared to only ~3% in unamended MFT-31 m (Table S4). Desulfotomaculum maintained its dominance (~46%) in amended MFT-31 m at 815 d incubation during the most active phase of biodegradation, whereas Proteobacteria (~40%) continued to dominate the unamended MFT-31 m community. Another significant change in bacterial community structure noted at 815 d incubation was the increase in Smithella in the amended MFT-31 m (~28%) compared to unamended MFT-31 m (~3%) (Table S4). MFT-6m cultures inoculated with a paraffin-degrading enrichment culture (MFT-6m-P) also exhibited high proportions of Peptococcaceae (24e57%, particularly Desulfotomaculum) regardless of amendment and sample time (Fig. S3A and Table S5); this is surely due to the actively alkane-degrading inoculum. In addition, the proportion of Smithella increased in amended MFT-6m-P from 3% at 420 d to 48% of bacterial reads by 815 d. The archaeal community in all MFT-containing cultures and all treatments at all sampling times was dominated by Methanomicrobia including Methanoregula. In MFT-6m, Methanoregula (hydrogenotrophic methanogens) and Methanosaeta (acetoclastic methanogens) constituted ~60e63% and 32e35% of the archaeal community, respectively, in both unamended and amended MFT at 120 d before significant CH4 production was observed. During active methanogenesis in both the alkane-amended and unamended MFT at 815 d incubation, the proportion of Methanoregula had decreased while a different hydrogenotrophic methanogen (Methanoculleus) had increased 5-fold to 10e11% and Methanosaeta was almost unchanged (Fig. 2B and Table S6). The same archaeal taxa were dominant in MFT-31 m cultures: in unamended MFT31 m, Methanoregula (~40%) and Methanosaeta (~43%) remained

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Fig. 3. Microbial community structure determined by 16S rRNA gene pyrosequencing of unamended and amended MFT-31 m samples during microbial metabolism of iso-alkanes and cycloalkanes to CH4. Graphs show proportions of (A) bacterial taxa and (B) archaeal taxa at 182, 420 and 815 d incubation. “Other Bacteria” and “Other Archaea” are the sums of taxa individually present at
1% abundance.

unchanged at 182, 420 and 815 d incubation. In amended MFT31 m, Methanoregula decreased in proportion during active hydrocarbon metabolism from 38% to 9% by 815 d incubation as another hydrogenotroph (Methanoculleus) increased from 4% to 27%. Methanosaeta increased during active hydrocarbon metabolism in amended MFT-31 m to constitute 62% of archaeal reads by 815 d incubation (Fig. 3B and Table S7). In the inoculated MFT-6m-P culture, there was little difference between unamended and amended MFT at both 420 and 815 d, with Methanoregula present at 64e68% and Methanosaeta at ~30% (Fig. S3B and Table S8). In the amended No-MFT-P, microcosms Methanoculleus outnumbered Methanoregula (rather than vice versa in uninoculated MFT cultures) and constituted ~53% of archaeal reads by 815 d.

4. Discussion 4.1. Duration of lag times Lengthy lag times have been observed repeatedly with oil sands tailings cultures (Siddique et al., 2015) and have been attributed to various causes including inhibition of methanogenesis by volatile hydrocarbons, sequestration of labile hydrocarbons by residual bitumen in MFT, and time required for assemblage (metabolically or perhaps physically) of synergistic consortia. The similar duration of lag phases exhibited by both MFT-6m and MFT-31 m is interesting, as it indicates that biodegradation capability was retained in aged MFT despite the depleted background naphtha in the deeper MFT sample and that the factor(s) affecting lag time were similar in both MFT samples. Also interesting is the proportion of Bacteria and Archaea in both amended and unamended cultures during the lag phase: methanogenic Archaea dominated the total pyrosequencing reads (>80%) in MFT-6m whereas MFT-31 m was dominated by Bacteria (~70% of total reads). By the end of lag phase, with onset of biodegradation and methanogenesis from added alkanes, amended MFT-31 m had shifted to an Archaea-dominated community (~70%), resembling MFT-6m and the inoculated cultures (Fig. S2). The latter cultures (MFT-6m-P and MFT-6m-N) exhibited shorter lag phases (180 and 490 d, respectively), likely due to the addition

of acclimated hydrocarbon-degrading microbial consortia, particularly those adapted to the paraffinic diluent (P culture).

4.2. Patterns of iso-alkane degradation Patterns of susceptibility of iso-alkanes were similar in each of the experimental sets, in the order of preference 3-MC6 > 4-MC7 > 2-MC8 > 2-MC7 and concomitant with CH4 production exceeding that of background (endogenous substrate methanogenesis). In contrast, biodegradation of 3-EC6 was delayed. A major n-alkane (nheptane; n-C7) from endogenous naphtha in MFT-6m was measured along with the added alkanes, to serve as a proxy for other labile endogenous hydrocarbons (Siddique et al., 2006). Its biodegradation, plotted with that of the iso-alkanes (Figs. 1C, 1D, S1C, S1D) to establish the biodegradation sequence, clearly shows that 3-EC6 was partially depleted in MFT-6m cultures only when labile substrates such as endogenous n-C7 were still present, after which 3-EC6 biodegradation ceased (e.g., Figs. 1C and S1D). Notably, 3-EC6 was not degraded in either the MFT-31 m or the no-MFT-P culture, both of which lacked endogenous alkanes (Figs. 1D and S1E). These observations imply that 3-EC6 is degraded via cometabolism as suggested by Mohamad Shahimin and Siddique (2017a). Supporting the current study, Abu Laban et al. (2014) also observed biodegradation of 3-MC6 and 4-MC7 in a naphthadegrading enrichment culture derived from MLSB MFT amended with a mixture of iso-alkanes. Abu Laban et al. (2014) and Tan et al. (2015) additionally observed biodegradation of 2-MC5, probable cometabolism of 3-MC5 and 2-MC6, and possible diauxic degradation of 2-MC5. Subsequently, a comprehensive study of naphtha biodegradation in MFT from other oil sands operators (Mohamad Shahimin and Siddique, 2017a) revealed biodegradation of a broader range of iso-alkanes (2-MC5, 3-MC6, 2-MC7, 4-MC7, 2-MC8, 3-MC8, 4-MC8, and 2-MC9), corroborating the biodegradability of iso-alkanes in Syncrude MFT collected from different depths and tested in this study. 3-MC5 has been shown either to resist biodegradation (Tan et al., 2015) or to be only slowly depleted in alkane mixtures (Siddique et al., 2015; Tan et al., 2015). Recently, Chen et al. (2019) also reported methanogenic biodegradation of 2-

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MC8, 3-MC8, and 4-MC8 in enrichment cultures derived from a high-temperature petroleum reservoir. We compiled and scrutinized biodegradation observations from the current and previously published studies but did not discern an obvious pattern of susceptibility of iso-alkanes to methanogenic biodegradation based on molecular weight, methyl- or ethylsubstitution position or physical characteristics such as vapour pressure and water solubility. Thus, defining the suite of labile isoalkanes supporting methanogenesis remains an empirical task, to which the current study contributes. 4.3. Patterns of cycloalkane degradation Among the added cycloalkanes in all four sets of experiments in this current study, only EcyC5 was partially depleted whereas McyC6 and EcyC6 remained undegraded during 1700 d incubation. Biodegradation of McyC5 (likely diauxic) was also reported previously in an enrichment culture derived from oil sands tailings (Tan et al., 2015) and also during incubation of naphtha with CNUL MFT where McyC5 and EcyC5 were biodegraded by indigenous microorganisms, leaving McyC6 and EcyC6 undegraded during ~1600 d incubation (Mohamad Shahimin and Siddique, 2017a). In the current study, EcyC5 degradation was delayed until after depletion of n-C7 and/or labile iso-alkanes (Figs. 1C, 1D, S1C, S1D) suggesting a diauxic effect. Our long-term studies (1600 d) on the biodegradation of naphtha and paraffinic diluent in CNRL and CNUL MFT (Mohamad Shahimin and Siddique, 2017a, 2017b) likewise have shown sequential biodegradation of n- > iso- > cycloalkanes. 4.4. The stimulatory effect of MFT on methanogenesis An interesting aspect of this current study is the slower kinetics of CH4 production and alkane degradation in the enrichment culture devoid of added MFT (No-MFT-P) compared with cultures containing MFT. For example, the amended No-MFT-P culture produced less CH4 than that produced by either the amended or the unamended MFT-6m-P cultures. This could be explained partially by methanogenesis from endogenous hydrocarbons in the MFT, but mathematically adding the methane produced by the No MFT-P culture (i.e., CH4 from added alkanes) to that from unamended MFT-6m-P (i.e., CH4 from endogenous substrates) still does not equal CH4 production from MFT-6m-P (i.e., CH4 from endogenous plus added alkanes), suggesting that the presence of MFT further enhances methanogenesis from the iso- and cycloalkane mixture beyond simple provision of endogenous substrates, perhaps by increasing rates of biodegradation without affecting the range of substrates biodegraded. Mechanisms could include: sequestration of toxic hydrocarbons by organic components of MFT (e.g., residual bitumen); providing solid phase iron (Fe) minerals that may facilitate methanogenesis in MFT through inter-species electron transfer (Jiang et al., 2013; Bray et al., 2017; Yao et al., 2017) or that simply provide surfaces facilitating assembly of consortia; providing trace elements or co-factors for metabolism; providing residual hydrocarbons that co-solvate amended alkanes to increase their biological availability; enhancing co-metabolism of residual substrates through the introduction of fresh labile substrates; and/ or provision of endogenous microbiota, e.g., a quorum of suitable species. The use of qPCR might have provided some insight into the absolute microbial density in MFT-amended versus MFT-free cultures to help discern the importance of the latter mechanism, versus the relative abundances provided by non-quantitative PCR. 4.5. Theoretical maximum CH4 production from amended alkanes Our stoichiometric calculations (Table S2) suggest that most iso-

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alkanes are completely metabolized rather than undergoing partial oxidation to organic metabolites because the amended MFT produced 56e73% of the predicted theoretical maximum CH4. These results are similar to our previous report on the complete biodegradation of 2-methylpentane (iso-alkane) under methanogenic conditions (Siddique et al., 2015). Our previous studies also reported 75e85% of theoretical maximum CH4 production during biodegradation of easily biodegradable fractions (n-alkanes and monoaromatics) (Siddique et al., 2006; Siddique et al., 2007) and 73e76% for the biodegradation of whole naphtha (Mohamad Shahimin and Siddique, 2017a). In contrast, the lower value of measured CH4 (1.19 mmol) versus the theoretical maximum CH4 (2.8 mmol) in No-MFT-P cultures (Table S2) suggests the incomplete mineralization of some of the iso-alkanes, highlighting the apparent beneficial role of solid phase (Fe minerals) in methanogenesis. 4.6. Bacterial community structure and effect of inoculation Pyrosequencing of 16S rRNA genes revealed conspicuous changes in the microbial community structure during hydrocarbon metabolism. Microbial communities actively biodegrading hydrocarbon (i.e., MFT-6m, No-MFT-P and eventually all amended cultures during methanogenesis) had high proportions of Archaea whereas the initially ‘dormant’ (aged) MFT-31 m had a small proportion of archaeal reads. Similar observations have been reported by Mohamad Shahimin and Siddique (2017a, 2017b) who investigated biodegradation of paraffinic solvent and naphtha diluent in two different oil sands tailings, and by Foght et al. (Fig. S2 in Foght et al., 2017) who characterized microbial community structure in depth profile samples from MLSB. From these results, we can speculate that it is the assembly of a sufficiently large proportion of methanogens with appropriate functional diversity that determines the time of onset of methanogenesis, and it seems that bacterial breakdown of the hydrocarbons is not a primary ratelimiting step in methanogenesis. Prior to active depletion of amended hydrocarbons, the bacterial communities were dominated by Proteobacteria but, during active biodegradation and methanogenesis, the community shifted to Firmicutes as members of Peptococcaceae increased to constitute >45% of the bacterial reads (Tables S3 and S4). In particular, the genus Desulfotomaculum comprised >95% of the Peptococcaceae, indicating their prominence in iso- and cycloalkane degradation as noted recently by Toth and Gieg (2018) during methanogenic biodegradation of short-chain n-alkanes, cyclohexane and monoaromatics. Similar results were observed during methanogenic biodegradation of C7eC8 iso-alkanes in an enrichment culture derived from Syncrude MFT (Abu Laban et al., 2014), C5eC6 iso-alkanes in Syncrude and Shell Albian MFT (Siddique et al., 2015) and a naphtha diluent containing a significant fraction of iso-alkanes in Shell Albian MFT (Mohamad Shahimin and Siddique, 2017a). A partial single-cell amplified genome sequence of an uncultivated Peptococcaeae member most closely affiliated with Desulfotomaculum has been published using cells sorted from an alkanedegrading MFT-derived enrichment culture, documenting the presence of putative alkylsuccinate synthase (ass) genes for anaerobic alkane metabolism (Tan et al., 2014a). Phylotypes of Peptococcaceae have also been implicated in n-alkane (C5eC10) and naphtha biodegradation in Syncrude MFT (Siddique et al., 2012); nalkane (C5eC6) and paraffinic diluent biodegradation in CNRL MFT (Mohamad Shahimin et al., 2016; Mohamad Shahimin and Siddique, 2017b); and n-alkane (C5eC10) and naphtha biodegradation in Shell Albian MFT (Mohamad Shahimin et al., 2016; Mohamad Shahimin and Siddique, 2017a). In addition to Desulfotomaculum, the proportion of Smithella reads were greater in

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amended than unamended MFT-31 m (28% versus 3%), as we have previously reported for Syntrophus/Smithella during shorter nalkane (C5eC10), longer n-alkanes (C14eC18) and naphtha biodegradation in Syncrude and CNRL MFT (Siddique et al., 2011, 2012; Mohamad Shahimin et al., 2016; Mohamad Shahimin and Siddique, 2017a) supporting the inferred role of Syntrophus/Smithella in the biodegradation of longer-chain hydrocarbons (e.g., Gray et al., 2011; Tan et al., 2014b). These bacteria were also found to be co-dominant with sulfate reducers and methanogens in naphtha-receiving tailings ponds during pyrotag surveys (Foght et al., 2017) and in oilfield produced water cultures biodegrading crude oil (Toth and Gieg, 2018). In contrast, the low proportions of bacterial taxa such as Chloroflexi and Actinobacteria suggest that these groups are not directly involved in iso-alkane degradation but rather might scavenge otherwise inhibitory metabolites (e.g., fermentation products) and/or provide co-factors or substrates such as H2þCO2 and/or acetate for subsequent utilization by methanogens.

4.7. Archaeal community structure Co-dominance of hydrogenotrophic and acetoclastic methanogens during hydrocarbon metabolism in MFT (Figs. 2 and 3, S3, and Tables S6eS8) suggests the existence of both acetoclastic and hydrogenotrophic pathways in the cultures, as previously inferred (Siddique et al., 2011, 2012; 2015; Mohamad Shahimin and Siddique, 2017a; 2017b; Abu Laban et al., 2014). However, the proportions of genera changed during incubation. Pyrosequencing reads affiliated with hydrogenotrophic (Methanoregula) and acetoclastic (Methanosaeta) methanogens dominated the archaeal community in unamended and amended MFT, but during active hydrocarbon metabolism in MFT, Methanoregula decreased with a corresponding increase (10e27%) in other hydrogenotrophic methanogens (Methanoculleus). The proportion of archaeal community affiliated with Methanosaeta either remained unchanged or increased during hydrocarbon metabolism. Different results were observed, however, in the amended inoculated medium devoid of MFT solid phase (No-MFT-P) where Methanoculleus (53%) outnumbered Methanoregula (Table S8) correlating with the presence of a solid phase, as discussed above.

5. Conclusions The current study highlights three aspects of methanogenesis: (1) iso-alkanes, a group of hydrocarbons that are considered relatively recalcitrant to anaerobic metabolism, are biodegradable under methanogenic conditions; (2) solid phase (clay minerals) helps maintain the kinetics of hydrocarbon biodegradation; and (3) bioaugmentation (using acclimated hydrocarbon-degrading microbes) reduces the long lag phase generally observed during methanogenic biodegradation of hydrocarbons. Our observations extend the previously known suite of susceptible hydrocarbons upon which a preliminary model for predicting greenhouse gas emissions from tailings ponds was based (Siddique et al., 2008), and allows further improvement and refinement of the model (Kong et al., 2019). Moreover, the current study provides insight into biogeochemical processes that affect tailings management, since it is known that methanogenensis increases tailings consolidation by altering tailings chemistry (Siddique et al., 2014a, 2014b) as well as releasing and transporting contaminants from underlying MFT to overlying cap water in end-pit lakes. Extrapolating the results can also increase understanding of in situ methanogenesis in oil reservoirs, and inform strategies such as bioaugmentation for remediating hydrocarbon impact in anoxic environments.

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