- Original article
- Open Access
Methanogenic biodegradation of C9 to C12n-alkanes initiated by Smithella via fumarate addition mechanism
AMB Express volume 10, Article number: 23 (2020)
In the present study, a methanogenic alkane-degrading (a mixture of C9 to C12n-alkanes) culture enriched from production water of a low-temperature oil reservoir was established and assessed. Significant methane production was detected in the alkane-amended enrichment cultures compared with alkane-free controls over an incubation period of 1 year. At the end of the incubation, fumarate addition metabolites (C9 to C12 alkylsuccinates) and assA genes (encoding the alpha subunit of alkylsuccinate synthase) were detected only in the alkane-amended enrichment cultures. Microbial community analysis showed that putative syntrophic n-alkane degraders (Smithella) capable of initiating n-alkanes by fumarate addition mechanism were enriched in the alkane-amended enrichment cultures. In addition, both hydrogenotrophic (Methanocalculus) and acetoclastic (Methanothrix) methanogens were also observed. Our results provide further evidence that alkanes can be activated by addition to fumarate under methanogenic conditions.
Methanogenic biodegradation of crude oil is a prevalent process occurring in subsurface petroleum reservoirs and has adverse effect on oil quality (Head et al. 2003, 2006; Jones et al. 2008). However, it has been postulated that methanogenic crude oil degradation can be applied for energy recovery in depleted petroleum reservoirs by bioconversion of residual oil to methane (Gieg et al. 2008). In addition to energy recovery, methanogenic degradation of crude oil is also a major process for bioremediation in the oil-contaminated environments after the depletion of electron acceptors (Amos et al. 2005; Feisthauer et al. 2010, 2012).
n-Alkanes are the major constituents of crude oil and also the significant contaminants in oil-polluted environments. Methanogenic biodegradation of n-alkanes requires the initial activation of these substrates before the further degradation (Zengler et al. 1999). Alkane activation by homolytic cleavage of the C–H bond, followed by addition of the resulting radical to the double bond of fumarate with the formation of alkylsuccinates is the most ubiquitous anaerobic n-alkane activation mechanism (Callaghan 2013), which has been demonstrated under sulfate- (Kniemeyer et al. 2007; Kropp et al. 2000) and nitrate-reducing conditions (Rabus et al. 2001). Only a few studies proved fumarate addition occurred under methanogenic conditions with limited detection of initial metabolites alkylsuccinates. Toth and Gieg detected C1 to C9 alkylsuccinates and assA genes over the incubation time of the methanogenic crude oil-degrading enrichment cultures (Toth and Gieg 2017). Qin et al. identified C15 and C16 alkylsuccinates in methanogenic pentadecane- and hexadecane-degrading enrichment cultures, respectively (Qin et al. 2017). In our recent work, a series of C16 to C20 alkylsuccinates and assA genes were detected in the methanogenic enrichment cultures amended with C16 to C20n-alkanes (Ji et al. 2019).
Although methanogenic biodegradation of crude oil (Aitken et al. 2013; Gieg et al. 2010; Gray et al. 2011; Jones et al. 2008; Toth and Gieg 2017) and longer n-alkanes (≥ C14) (Liang et al. 2016; Siddique et al. 2011; Wawrik et al. 2016; Zengler et al. 1999; Zhou et al. 2012) has been intensively investigated, methanogenic biodegradation of low molecular weight n-alkanes has not been extensively studied (e.g. activation mechanisms and syntrophic degraders). Members of genus Smithella (in the family Syntrophaceae), implicated in syntrophic alkane degradation, were frequently identified in methanogenic crude oil-degrading enrichment cultures (Gray et al. 2011; Jones et al. 2008; Toth and Gieg 2017). Sherry et al. observed that Smithella was significantly enriched in both the weathered and non-weathered oil-amended (containing C5 to C10n-alkanes) methanogenic enrichment cultures, indicating that Smithella can utilize low molecular weight n-alkanes (Sherry et al. 2014). Novel members of the family Peptococcaceae were identified to be the primary degraders in several methanogenic short alkane-degrading (C5 to C10; n-, iso- and cyclo-alkanes) enrichment cultures derived from oil sands tailings ponds (Abu Laban et al. 2015; Mohamad Shahimin et al. 2016; Mohamad Shahimin and Siddique 2017; Siddique et al. 2015; Tan et al. 2014; Tan et al. 2013). By investigating methanogenic biodegradation of C7 to C8iso-alkanes, Abu Laban et al. proposed a novel family Peptococcaceae activated these substrates by addition to fumarate with the detection of high abundance of Peptococcaceae, Peptococcaceae-related assA gene and fumarate addition metabolites of C7 to C8iso-alkans (Abu Laban et al. 2015). Although a positive expression of assA gene and fumarate addition metabolites of 2-methylpentane and methylcyclopentane were detected in the methanogenic short alkane-degrading (C6 to C10; n-, iso- and cyclo-alkanes) cultures, Tan et al. still failed to detect initial activation metabolites of n-alkanes (Tan et al. 2015).
Here, we established a methanogenic enrichment culture growing on C9 to C12n-alkanes inoculated with production water from a low-temperature petroleum reservoir. Methane production was periodically monitored during the incubation. Microbial community compositions, functional genes (assA and mcrA) and metabolite profiles were analyzed at the end of the incubation period.
Materials and methods
Production water from Xinjiang Kelamayi oil field block 6 (about 21 °C) was collected and stored in a serum bottle with headspace filled with N2 (99.99% purity). The production water was stored for over 1 year to consume the residual organics. Sterilized basal medium with no electron acceptor (Wang et al. 2011) was dispensed in 120 mL-serum bottles as 48 mL per each. 2 mL of the production water was transferred to each bottle by syringe. Each alkane-amended enrichment culture contained 0.225 mmol of each n-alkane, including n-nonane (C9; ≥ 99%), n-decane (C10; ≥ 99%), n-undecane (C11; ≥ 99%) and n-dodecane (C12; ≥ 99%). Alkane-free control cultures received no n-alkane. The serum bottles were sealed with butyl rubber stoppers. All the cultures were set up in two replications and stationarily incubated at room temperature (around 21 °C) in the dark.
Methane production was measured using a gas chromatography (GC model 9890B, Shanghai, China) equipped with a flame ionization detection (FID). 200 μL headspace gas taken by a gastight syringe were injected into GC for analysis. Program setting of the GC analysis was: the initial column temperature was set at 50 °C for 2 min, then increased to 130 °C at a rate of 15 °C/min, the temperature of 130 °C sustained for 1 min; the second increase was conducted at a rate of 30 °C/min to 180 °C for 30 min. The temperature of injector and FID was 200 °C. External standard curve of the methane was used for methane concentration calculation (Ma et al. 2018).
To detect acid metabolites in the cultures, about 40 mL of culture aliquots was collected. These culture aliquots were refluxed at 100 °C for 8 h with 50 mL 1 M KOH in a 50% methanol, 50% water mixture for saponification. This was followed by acidification to pH < 2 with HCl. The organic fraction was then extracted with ethyl acetate and derivatized to ethyl esters with 10 mL of ethanol, 10 mL of cyclohexane and 0.2 g of NaHSO4 (refluxed at 80 °C for 8 h). After rotary evaporation, 10 mL deionized water was added. Metabolites were extracted with 10 mL ethyl acetate for three times and concentrated to about 200 μL. 1 μL sample was injected into GC–MS in a splitless mode for analysis. An Agilent 7890A GC coupled to a MSD 5975C mass detector was used. The injector temperature was 280 °C. The program of GC–MS was followed: the initial temperature was held at 60 °C for 2 min, then increased at a rate of 10 °C/min to 280 °C for 20 min. The MS detector was run in the scan mode from 30 to 1000 mass units.
Diethyl (1-methyloctyl)succinate was synthesized according to Bian et al. (2014). The identification of (1-methyloctyl)succinate in the enrichment cultures was compared with the synthesized authentic standard (Additional file 1: Figure S1). (1-methylnonyl)succinate, (1-methyldecyl)succinate and (1-methylundecyl)succinate were identified by their characteristic fragment ions (128, 174, [M–45]+ and [M–87]+) (Bian et al. 2014) and relative retention times. Diethyl succinate, diethyl glutarate, diethyl adipate, diethyl suberate and diethyl azelate were synthesized by ethyl esterification of succinic acid, glutaric acid, adipic acid, suberic acid and azelaic acid, respectively. The reaction mixture contained 2 mg of dicarboxylic acid, 10 mL of ethanol, 10 mL of cyclohexane and 0.2 g of NaHSO4. The reaction mixture was refluxed at 80 °C for 8 h. The ethanol and cyclohexane were removed by rotary evaporation, and the residue was treated with 10 mL water. Diethyl products were extracted with extracted with 10 mL ethyl acetate for three times and analyzed by GC–MS in a same program of culture metabolites analysis. α,ω-Dicarboxylic acids in the enrichment cultures were identified by comparison with these authentic standards (Additional file 1: Figure S2). Fatty acids were identified by matching library spectra NIST (https://webbook.nist.gov/chemistry/).
Microbial community analysis
10 mL of culture aliquot were collected for genomic DNA extraction using the AxyPrep™ Bacterial Genomic DNA Maxiprep Kit (Axygen Biosciences, USA). Archaeal and bacterial 16S rRNA genes were amplified using 344F (5′-ACGGGGYGCAGCAGGCGCGA-3′)/915R (5′-GTGCTCCCCCGCCAATTCCT-3′) (Casamayor et al. 2002) and 515F (5′-GTGCCAGCMGCCGCGG-3′)/907R (5′-CCGTCAATTCMTTTRAGTTT-3′) (Xiong et al. 2012), respectively. 16S rRNA gene polymerase chain reaction (PCR) and Illumina sequencing were performed as previously described (Ma et al. 2018). Operational taxonomic units (OTUs) were classified using Usearch (Edgar 2013) against the SILVA SSU database 128 (Quast et al. 2013) with the 97% similarity.
assA and mcrA genes analysis
Alkylsuccinate synthase gene (assA) and methyl coenzyme-M reductase gene (mcrA) as the key functional genes involved in the methanogenic n-alkane degradation process were investigated. PCR primer sets of assA2F/assA2R (Callaghan et al. 2010) and MLF/MLR (Luton et al. 2002) were used for the PCR amplification of assA and mcrA gene, respectively. PCR cycling conditions for both assA and mcrA gene were conducted as follows: 95 °C for 3 min; 40 cycles of 95 °C for 45 s, 55 °C for 60 s, 72 °C for 2 min; and 72 °C for 10 min. PCR products were purified and cloned, and positive clones were picked for sanger sequencing on ABI 377 automated sequencer (Liang et al. 2015). The valid nucleotide sequences were translated to protein sequences using ORFfinder translation tool (https://www.ncbi.nlm.nih.gov/orffinder/). Protein sequences were classified to OTUs using CD-HIT Suite (Huang et al. 2010) with the 97% similarity. Representative protein sequences were compared with GenBank Database using BLAST to identify similar sequences. Phylogenetic analyses were conducted using MEGA6.0 software with neighbor-joining method and 1000 bootstrap replicates.
The sequences generated in this study were deposited in GenBank under accession numbers SAMN08904491 and SAMN08904496 (bacterial and archaeal 16S rRNA genes), MH192396-MH192461 (assA genes), MH192647-MH192713 (mcrA genes). The sequencing data of alkane-free control cultures were available as previously (Ji et al. 2019).
Methane and intermediate metabolites analysis
Methane production started after 85 days’ incubation in alkane-amended (a mixture of C9 to C12n-alkanes) enrichment cultures and reached about 33 μmol at the end of the incubation (364 days) (Fig. 1). No methane was detected in the alkane-free controls (Ji et al. 2019) (Fig. 1).
Potential anaerobic intermediates of n-alkanes were analyzed by GC–MS. (1-Methyloctyl)succinate (C9 alkylsuccinate), (1-methylnonyl)succinate (C10 alkylsuccinate), (1-methyldecyl)succinate (C11 alkylsuccinate) and (1-methylundecyl)succinate (C12 alkylsuccinate), generated from fumarate addition to the alkanes n-nonane, n-decane, n-undecane and n-dodecane respectively, were detected in the alkane-amended enrichment cultures (Fig. 2). All identified metabolites displayed the signature fragments at m/z 128, 174, [M–45]+ and [M–87]+, which are distinctive for alkylsuccinates (Bian et al. 2014) (Fig. 2). The identity of C9 alkylsuccinate in the alkane-amended enrichment cultures was confirmed by comparing its ion fragmentation patterns and retention time with that of a synthesized standard (Fig. 2, Additional file 1: Figure S1). No alkylsuccinates were identified in the alkane-free controls (Ji et al. 2019).
Long-chain fatty acids included tetradecanoate, pentadecanoate, hexadecanoate, heptadecanoate, octadecanoate, eicosanoate and docosanoate were detected in the alkane-amended enrichment cultures (Additional file 1: Figure S3). Only hexadecanoate and octadecanoate were detected in the alkane-free controls. Several α,ω-dicarboxylic acids were specifically identified in the alkane-amended enrichment cultures by comparing to the authentic standards with mass spectral profiles and retention times. These included butanedioic (succinic) acid, pentanedioic (glutaric) acid, hexanedioic (adipic) acid, octanedioic (suberic) acid and nonanedioic (azelaic) acid (Additional file 1: Figure S2).
Microbial community compositions
Substantial difference of microbial community compositions was observed between alkane-amended enrichment cultures and alkane-free control cultures at the end of the incubation (364 days). Smithella sp. had the highest relative abundance in the alkane-amended enrichment cultures (Fig. 3a). Other abundant bacterial phylotypes affiliated to Anaerolineaceae, Desulfovibrio, Desulfatibacillum, Proteiniphilum, Thermovirga, and unclassified NB1-n (Fig. 3a). In the alkane-free control cultures, members of Geoalkalibacter and Thermacetogenium became the dominant bacteria (Ji et al. 2019) (Fig. 3a). The archaeal community in the alkane-amended enrichment cultures was dominated by hydrogenotrophic methanogens of Methanocalculus (84%) and Methanothermobacter (10%) (Fig. 3b). Methanothrix (Methanosaeta, acetoclastic methanogen) was also detected in the alkane-amended enrichment cultures and comprised about 5% of the total archaeal community (Fig. 3b). The archaeal community in the alkane-free control cultures was essentially comprised by Methanothermobacter (98%) (Ji et al. 2019) (Fig. 3b).
Phylogenetic analysis of assA and mcrA genes
Genes encoding for the alkylsuccinate synthase were only detected in the alkane-amended enrichment cultures. All sequences were clustered into Smithella subclade and were most closely related with assA sequence of Smithella sp. SC_K08D17 (Fig. 4). Both alkane-amended enrichment cultures and control cultures detected mcrA genes (Ji et al. 2019). In the alkane-amended enrichment cultures, most mcrA sequences affiliated with Methanocalculus and only one sequence (a total of 67 valid sequences) belonged to Methanothermobacter (Additional file 1: Figure S4).
Methanogenic biodegradation of C9 to C12n-alkanes initiated by addition to fumarate
The detection of corresponding fumarate addition products (C9 to C12 alkylsuccinates) provides convincing evidence that the oxidation of C9 to C12n-alkanes was initiated by addition to fumarate under methanogenic conditions. It is supported further by the detection of assA genes. Previous studies have reported C1 to C8 alkylsuccinates detected in the production water from oil reservoirs (Agrawal and Gieg 2013; Bian et al. 2015; Duncan et al. 2009; Gieg et al. 2010). And C1 to C9, C15 to C20 alkylsuccinates have been detected under methanogenic conditions associated with microorganisms derived from oil reservoirs (Qin et al. 2017; Toth and Gieg 2017). Here the identification of C9 to C12 alkylsuccinates fills a gap that a series of n-alkanes can be activated by fumarate addition by oilfield-related microorganisms.
Except for fumarate addition products, several dicarboxylic acids were detected in alkane-amended enrichment cultures. These diacids may be cell-associated or secreted from the cells. Oberding and Gieg detected several α,ω-dicarboxylic acids in methanogenic n-octacosane-degrading enrichment cultures (Oberding and Gieg 2018). The authors suggested that these dicarboxylic acids might act as biosurfactants, which could increase substrate accessibility (Oberding and Gieg 2018). Although the origin of these dicarboxylic acids is elusive, the fatty acids as the downstream metabolites involved in fumarate addition pathway (Wilkes et al. 2002), may play a role in alkane emulsification in the current study (Embree et al. 2014).
Key members involved in methanogenic n-alkane degradation
The dominant bacteria in the alkane-amended enrichment cultures were Smithella. Members of Smithella have been detected in numerous methanogenic alkane- and crude oil-degrading enrichment cultures (Cheng et al. 2013; Oberding and Gieg 2018; Siddique et al. 2011; Wawrik et al. 2016; Zengler et al. 1999) and are generally considered as syntrophic n-alkane degraders (Gray et al. 2011). In this study, assA genes closely related to Smithella species was detected, suggesting that Smithella participated in methanogenic n-alkane degradation and initiated alkane activation by fumarate addition reaction.
The abundance of Anaerolineaceae was found to be increased in the alkane-amended enrichment cultures. Microorganisms affiliated to the family Anaerolineaceae have been detected in a vast number of methanogenic alkane-degrading enrichment cultures and were implicated to be responsible for alkane activation in these cultures (Liang et al. 2015; Liang et al. 2016; Mohamad Shahimin et al. 2016; Mohamad Shahimin and Siddique 2017). However, Anaerolineaceae-related assA genes were not detected in the current culture, consistent with previous studies (Liang et al. 2016; Mohamad Shahimin et al. 2016). It has also been suggested that Anaerolineaceae may serve as a secondary degrader in oxidizing fermentative products from primary degraders (Tan et al. 2015). Based on the results of this study, the role of Anaerolineaceae is currently unknown.
Our results suggest that fumarate addition is a key alkane initial activation mechanism under methanogenic conditions. Smithella were identified as primary syntrophic n-alkane degraders, which can activate C9 to C12n-alkanes by addition to fumarate. This work expands our knowledge about the biochemical process involved in the methanogenic hydrocarbon biodegradation in petroleum reservoirs and oil-contaminated environments.
Availability of data and materials
Raw reads from microbial community sequencing are available in the GenBank archive at the National Center for Biotechnological Information (NCBI) as listed in the manuscript.
- assA :
alkylsuccinate synthase gene
- mcrA :
methyl coenzyme-M reductase gene
- C9 alkylsuccinate:
- C10 alkylsuccinate:
- C11 alkylsuccinate:
- C12 alkylsuccinate:
Abu Laban N, Dao A, Semple K, Foght J (2015) Biodegradation of C7 and C8 iso-alkanes under methanogenic conditions. Environ Microbiol 17(12):4898–4915. https://doi.org/10.1111/1462-2920.12643
Agrawal A, Gieg LM (2013) In situ detection of anaerobic alkane metabolites in subsurface environments. Front Microbiol 4:140. https://doi.org/10.3389/fmicb.2013.00140
Aitken CM, Jones DM, Maguire MJ, Gray ND, Sherry A, Bowler BFJ, Ditchfield AK, Larter SR, Head IM (2013) Evidence that crude oil alkane activation proceeds by different mechanisms under sulfate-reducing and methanogenic conditions. Geochim Cosmochim Ac 109:162–174. https://doi.org/10.1016/j.gca.2013.01.031
Amos RT, Mayer KU, Bekins BA, Delin GN, Williams RL (2005) Use of dissolved and vapor-phase gases to investigate methanogenic degradation of petroleum hydrocarbon contamination in the subsurface. Water Resour Res. https://doi.org/10.1029/2004wr003433
Bian XY, Mbadinga SM, Yang SZ, Gu JD, Ye RQ, Mu BZ (2014) Synthesis of anaerobic degradation biomarkers alkyl-, aryl- and cycloalkylsuccinic acids and their mass spectral characteristics. Eur J Mass Spectrom 20(4):287–297. https://doi.org/10.1255/ejms.1280
Bian XY, Mbadinga SM, Liu YF, Yang SZ, Liu JF, Ye RQ, Gu JD, Mu BZ (2015) Insights into the anaerobic biodegradation pathway of n-alkanes in oil reservoirs by detection of signature metabolites. Sci Rep 5:9801. https://doi.org/10.1038/srep09801
Callaghan AV (2013) Enzymes involved in the anaerobic oxidation of n-alkanes: from methane to long-chain paraffins. Front Microbiol 4:89. https://doi.org/10.3389/fmicb.2013.00089
Callaghan AV, Davidova IA, Savage-Ashlock K, Parisi VA, Gieg LM, Suflita JM, Kukor JJ, Wawrik B (2010) Diversity of benzyl- and alkylsuccinate synthase genes in hydrocarbon-impacted environments and enrichment cultures. Environ Sci Technol 44(19):7287–7294. https://doi.org/10.1021/es1002023
Casamayor EO, Massana R, Benlloch S, Øvreås L, Díez B, Goddard VJ, Gasol JM, Joint I, Rodríguez-Valera F, Pedrós-Alió C (2002) Changes in archaeal, bacterial and eukaryal assemblages along a salinity gradient by comparison of genetic fingerprinting methods in a multipond solar saltern. Environmen Microbiol 4(6):338–348. https://doi.org/10.1046/j.1462-2920.2002.00297.x
Cheng L, Ding C, Li Q, He Q, Dai LR, Zhang H (2013) DNA-SIP reveals that Syntrophaceae play an important role in methanogenic hexadecane degradation. PLoS ONE 8(7):e66784. https://doi.org/10.1371/journal.pone.0066784
Duncan KE, Gieg LM, Parisi VA, Tanner RS, Tringe SG, Bristow J, Suflita JM (2009) Biocorrosive thermophilic microbial communities in Alaskan North Slope oil facilities. Environ Sci Technol 43(20):7977–7984
Edgar RC (2013) UPARSE: highly accurate OTU sequences from microbial amplicon reads. Nat Methods 10(10):996–998. https://doi.org/10.1038/nmeth.2604
Embree M, Nagarajan H, Movahedi N, Chitsaz H, Zengler K (2014) Single-cell genome and metatranscriptome sequencing reveal metabolic interactions of an alkane-degrading methanogenic community. ISME J 8(4):757–767. https://doi.org/10.1038/ismej.2013.187
Feisthauer S, Siegert M, Seidel M, Richnow HH, Zengler K, Gründger F, Krüger M (2010) Isotopic fingerprinting of methane and CO2 formation from aliphatic and aromatic hydrocarbons. Org Geochem 41(5):482–490. https://doi.org/10.1016/j.orggeochem.2010.01.003
Feisthauer S, Seidel M, Bombach P, Traube S, Knöller K, Wange M, Fachmann S, Richnow HH (2012) Characterization of the relationship between microbial degradation processes at a hydrocarbon contaminated site using isotopic methods. J Contam Hydrol 133:17–29. https://doi.org/10.1016/j.jconhyd.2012.03.001
Gieg LM, Duncan KE, Suflita JM (2008) Bioenergy production via microbial conversion of residual oil to natural gas. Appl Environ Microbiol 74(10):3022–3029. https://doi.org/10.1128/AEM.00119-08
Gieg LM, Davidova IA, Duncan KE, Suflita JM (2010) Methanogenesis, sulfate reduction and crude oil biodegradation in hot Alaskan oilfields. Environ Microbiol 12(11):3074–3086. https://doi.org/10.1111/j.1462-2920.2010.02282.x
Gray ND, Sherry A, Grant RJ, Rowan AK, Hubert CR, Callbeck CM, Aitken CM, Jones DM, Adams JJ, Larter SR, Head IM (2011) The quantitative significance of Syntrophaceae and syntrophic partnerships in methanogenic degradation of crude oil alkanes. Environ Microbiol 13(11):2957–2975. https://doi.org/10.1111/j.1462-2920.2011.02570.x
Head IM, Jones DM, Larter SR (2003) Biological activity in the deep subsurface and the origin of heavy oil. Nature 426(6964):344–352. https://doi.org/10.1038/nature02134
Head IM, Jones DM, Roling WF (2006) Marine microorganisms make a meal of oil. Nat Rev Microbiol 4(3):173–182. https://doi.org/10.1038/nrmicro1348
Huang Y, Niu B, Gao Y, Fu L, Li W (2010) CD-HIT Suite: a web server for clustering and comparing biological sequences. Bioinformatics 26(5):680–682. https://doi.org/10.1093/bioinformatics/btq003
Ji JH, Liu YF, Zhou L, Mbadinga SM, Pan P, Chen J, Liu JF, Yang SZ, Sand W, Gu JD, Mu BZ (2019) Methanogenic degradation of long n-alkanes requires fumarate-dependent activation. Appl Environ Microbiol. https://doi.org/10.1128/aem.00985-19
Jones DM, Head IM, Gray ND, Adams JJ, Rowan AK, Aitken CM, Bennett B, Huang H, Brown A, Bowler BF, Oldenburg T, Erdmann M, Larter SR (2008) Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451(7175):176–180. https://doi.org/10.1038/nature06484
Kniemeyer O, Musat F, Sievert SM, Knittel K, Wilkes H, Blumenberg M, Michaelis W, Classen A, Bolm C, Joye SB, Widdel F (2007) Anaerobic oxidation of short-chain hydrocarbons by marine sulphate-reducing bacteria. Nature 449(7164):898–901. https://doi.org/10.1038/nature06200
Kropp KG, Davidova IA, Suflita JM (2000) Anaerobic oxidation of n-dodecane by an addition reaction in a sulfate-reducing bacterial enrichment culture. Appl Environ Microbiol 66(12):5393–5398
Liang B, Wang LY, Mbadinga SM, Liu JF, Yang SZ, Gu JD, Mu BZ (2015) Anaerolineaceae and Methanosaeta turned to be the dominant microorganisms in alkanes-dependent methanogenic culture after long-term of incubation. AMB Express 5(1):117. https://doi.org/10.1186/s13568-015-0117-4
Liang B, Wang LY, Zhou Z, Mbadinga SM, Zhou L, Liu JF, Yang SZ, Gu JD, Mu BZ (2016) High frequency of Thermodesulfovibrio spp. and Anaerolineaceae in association with Methanoculleus spp. in a long-term incubation of n-alkanes-degrading methanogenic enrichment culture. Front Microbiol 7:1431. https://doi.org/10.3389/fmicb.2016.01431
Luton PE, Wayne JM, Sharp RJ, Riley PW (2002) The mcrA gene as an alternative to 16S rRNA in the phylogenetic analysis of methanogen populations in landfill. Microbiology 148:3521–3530. https://doi.org/10.1099/00221287-148-11-3521
Ma L, Zhou L, Mbadinga SM, Gu JD, Mu BZ (2018) Accelerated CO2 reduction to methane for energy by zero valent iron in oil reservoir production waters. Energy 147:663–671. https://doi.org/10.1016/j.energy.2018.01.087
Mohamad Shahimin MF, Siddique T (2017) Methanogenic biodegradation of paraffinic solvent hydrocarbons in two different oil sands tailings. Sci Total Environ 583:115–122. https://doi.org/10.1016/j.scitotenv.2017.01.038
Mohamad Shahimin MF, Foght JM, Siddique T (2016) Preferential methanogenic biodegradation of short-chain n-alkanes by microbial communities from two different oil sands tailings ponds. Sci Total Environ 553:250–257. https://doi.org/10.1016/j.scitotenv.2016.02.061
Oberding LK, Gieg LM (2018) Methanogenic paraffin biodegradation: alkylsuccinate synthase gene quantification and dicarboxylic acid production. Appl Environ Microbiol 84(1):e01773. https://doi.org/10.1128/AEM.01773-17
Qin QS, Feng DS, Liu PF, He Q, Li X, Liu AM, Zhang H, Hu GQ, Cheng L (2017) Metagenomic characterization of Candidatus Smithella cisternae Strain M82_1, a syntrophic alkane-degrading bacteria, enriched from the Shengli Oil Field. Microbes Environ 32(3):234–243. https://doi.org/10.1264/jsme2.ME17022
Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, Peplies J, Glockner FO (2013) The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. https://doi.org/10.1093/nar/gks1219
Rabus R, Wilkes H, Behrends A, Armstroff A, Fischer T, Pierik AJ, Widdel F (2001) Anaerobic initial reaction of n-alkanes in a denitrifying bacterium: evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in n-hexane metabolism. J Bacteriol 183(5):1707–1715. https://doi.org/10.1128/JB.183.5.1707-1715.2001
Sherry A, Grant RJ, Aitken CM, Jones DM, Head IM, Gray ND (2014) Volatile hydrocarbons inhibit methanogenic crude oil degradation. Front Microbiol 5:131. https://doi.org/10.3389/fmicb.2014.00131
Siddique T, Penner T, Semple K, Foght JM (2011) Anaerobic biodegradation of longer-chain n-alkanes coupled to methane production in oil sands tailings. Environ Sci Technol 45(13):5892–5899. https://doi.org/10.1021/es200649t
Siddique T, Mohamad Shahimin MF, Zamir S, Semple K, Li C, Foght JM (2015) Long-term incubation reveals methanogenic biodegradation of C5 and C6 iso-alkanes in oil sands tailings. Environ Sci Technol 49(24):14732–14739. https://doi.org/10.1021/acs.est.5b04370
Tan B, Dong X, Sensen CW, Foght J (2013) Metagenomic analysis of an anaerobic alkane-degrading microbial culture: potential hydrocarbon-activating pathways and inferred roles of community members. Genome 56(10):599–611. https://doi.org/10.1139/gen-2013-0069
Tan B, Charchuk R, Li C, Nesbø C, Abu Laban N, Foght J (2014) Draft genome sequence of uncultivated Firmicutes (Peptococcaceae SCADC) single cells sorted from methanogenic alkane-degrading cultures. Genome Announc. https://doi.org/10.1128/genomea.00909-14
Tan B, Semple K, Foght J (2015) Anaerobic alkane biodegradation by cultures enriched from oil sands tailings ponds involves multiple species capable of fumarate addition. FEMS Microbiol Ecol. https://doi.org/10.1093/femsec/fiv042
Toth CRA, Gieg LM (2017) Time course-dependent methanogenic crude oil biodegradation: dynamics of fumarate addition metabolites, biodegradative genes, and microbial community composition. Front Microbiol 8:2610. https://doi.org/10.3389/fmicb.2017.02610
Wang LY, Gao CX, Mbadinga SM, Zhou L, Liu JF, Gu JD, Mu BZ (2011) Characterization of an alkane-degrading methanogenic enrichment culture from production water of an oil reservoir after 274 days of incubation. Int Biodeter Biodegr 65(3):444–450. https://doi.org/10.1016/j.ibiod.2010.12.010
Wawrik B, Marks CR, Davidova IA, McInerney MJ, Pruitt S, Duncan KE, Suflita JM, Callaghan AV (2016) Methanogenic paraffin degradation proceeds via alkane addition to fumarate by ‘Smithella’ spp. mediated by a syntrophic coupling with hydrogenotrophic methanogens. Environ Microbiol 18(8):2604–2619. https://doi.org/10.1111/1462-2920.13374
Wilkes H, Rabus R, Fischer T, Armstroff A, Behrends A, Widdel F (2002) Anaerobic degradation of n-hexane in a denitrifying bacterium: further degradation of the initial intermediate (1-methylpentyl)succinate via C-skeleton rearrangement. Arch Microbiol 177(3):235–243. https://doi.org/10.1007/s00203-001-0381-3
Xiong J, Liu Y, Lin X, Zhang H, Zeng J, Hou J, Yang Y, Yao T, Knight R, Chu H (2012) Geographic distance and pH drive bacterial distribution in alkaline lake sediments across Tibetan Plateau. Environ Microbiol 14(9):2457–2466. https://doi.org/10.1111/j.1462-2920.2012.02799.x
Zengler K, Richnow HH, Rosselló-Mora R, Michaelis W, Widdel F (1999) Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 401(6750):266–269. https://doi.org/10.1038/45777
Zhou L, Li KP, Mbadinga SM, Yang SZ, Gu JD, Mu BZ (2012) Analyses of n-alkanes degrading community dynamics of a high-temperature methanogenic consortium enriched from production water of a petroleum reservoir by a combination of molecular techniques. Ecotoxicology 21(6):1680–1691. https://doi.org/10.1007/s10646-012-0949-5
This work was supported by the National Natural Science Foundation of China (Grants Nos. 41530318, 41403066), NSFC/RGC Joint Research Fund (No. 41161160560), the Fundamental Research Funds for the Central Universities (Nos. 222201817017, 50321101917017, 22221818014) and the Research Program of State Key Laboratory of Bioreactor Engineering.
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Additional file 1: Figure S1. GC–MS analysis of a diethyl 2-(1-methyloctyl)succinate (C9 alkylsuccinate) standard. (a) GC partial ion chromatogram following selection for the m/z 128 ion of a diethyl 2-(1-methyloctyl)succinate standard, (b) Mass spectral profiles of diethyl 2-(1-methyloctyl)succinate (retention time, 17.60 min). Figure S2. Mass spectral profiles of dicarboxylic acids identified in alkane-amended enrichment cultures. Left panel: compound detected in the alkane-amended enrichment cultures. Right panel: ethyl-derivatized authentic standards. Figure S3. Mass spectral profiles of fatty acids (ethyl derivatives) identified in alkane-amended enrichment cultures. Figure S4. Phylogenetic tree of deduced amino acid sequences of methyl coenzyme-M reductase genes (mcrA) from alkane-amended enrichment culture (in red). Topology of the tree was obtained by the neighbor-joining method. Bootstrap values (n = 1000 replicates), values below 75% are not shown.
About this article
Cite this article
Ji, J., Zhou, L., Mbadinga, S.M. et al. Methanogenic biodegradation of C9 to C12n-alkanes initiated by Smithella via fumarate addition mechanism. AMB Expr 10, 23 (2020). https://doi.org/10.1186/s13568-020-0956-5
- Fumarate addition
- Oil reservoirs