FEMS Microbiology Ecology Advance Access published April 6, 2015
Anaerobic alkane biodegradation by cultures enriched from oil sands tailings ponds involves multiple species capable of fumarate addition
BoonFei Tan1, Kathleen Semple and Julia Foght* Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9
*Corresponding author: Julia Foght phone: +1 780.492.3279 fax: +1 780.492.0234 email: [email protected]
1
Current address: Center for Environmental Sensing and Modeling, Singapore-MIT Alliance for Research and Technology, 1 Create Way, #09-03 Create Tower, Singapore 138602.
Running title: Anaerobic alkane biodegradation Keywords: aliphatic hydrocarbon degradation / methanogenesis / sulphidogenesis
1
Abstract A methanogenic short-chain alkane-degrading culture (SCADC) was enriched from oil sands tailings and transferred several times with a mixture of C6, C7, C8 and C10 n-alkanes as the predominant organic carbon source, plus 2-methylpentane, 3-methylpentane and 5
methylcyclopentane as minor components. Cultures produced ~40% of the maximum theoretical methane during 18 months incubation while depleting the n-alkanes, 2-methylpentane and methylcyclopentane. Substrate depletion correlated with detection of metabolites characteristic of fumarate activation of 2-methylpentane and methylcyclopentane but not n-alkane metabolites. During active methanogenesis with the mixed alkanes, reverse-transcription PCR confirmed the
10
expression of functional genes (assA and bssA) associated with hydrocarbon addition to fumarate. Pyrosequencing of 16S rRNA genes amplified during active alkane degradation revealed enrichment of Clostridia (particularly Peptococcaceae) and methanogenic Archaea (Methanosaetaceae and Methanomicrobiaceae). Methanogenic cultures transferred into medium containing sulphate produced sulphide, depleted n-alkanes and produced the corresponding
15
succinylated alkane metabolites, but were slow to degrade 2-methylpentane and methylcyclopentane; these cultures were enriched in Deltaproteobacteria rather than Clostridia. 3-Methylpentane was not degraded by any cultures. Thus, nominally methanogenic oil sands tailings harbour dynamic and versatile hydrocarbon-degrading fermentative syntrophs and sulphate reducers capable of degrading n-, iso- and cyclo-alkanes by addition to fumarate.
20
2
20
Introduction Aliphatic hydrocarbons including n-, iso- and cyclo-alkanes are major constituents of crude oil and its refined products and are significant contaminants in petroleum-impacted environments. Whereas alkane biodegradation under aerobic conditions has been well known for decades (Van 25
Hamme et al., 2003) and metabolism under nitrate- and sulphate-reducing conditions has been described more recently (reviewed by Mbadinga et al., 2011), the key microbes, biodegradation pathways, metabolites, and functional genes involved in methanogenic alkane degradation have been cryptic or controversial (Aitken et al., 2013; Callaghan 2013; Embree et al. 2014). Recent reports (e.g., Tan et al. 2014a, b, c; Abu Laban 2015a) have begun to expand the repertoire of
30
microbes and genes associated with methanogenic alkane-degrading cultures. Enzymatic activation of aliphatic hydrocarbons by addition to fumarate is the best described and most widely documented mechanism for initiating anaerobic alkane degradation under nitrate- and sulphate-reducing conditions (Agrawal & Gieg, 2013, Callaghan, 2013). The glycyl radical enzyme alkylsuccinate synthase (ASS: also called methylalkylsuccinate synthase,
35
MAS) and its counterparts responsible for aromatic hydrocarbon activation [benzylsuccinate synthase (BSS) and naphthyl-2-methylsuccinate synthase (NMS)], have been described and reviewed (Mbadinga et al. 2011; Callaghan 2013; Agrawal and Gieg, 2013). The requisite genes, particularly those encoding the alpha-subunit, e.g., assA, have been detected in contaminated environments (Acosta-González et al., 2013), oil field fluids (Zhou et al., 2012), enrichment
40
cultures (Callaghan et al., 2010; Aitken et al., 2013) and model n-alkane-degrading organisms such as nitrate-reducing ‘Aromatoleum’ sp. HxN1 (Grundmann et al., 2008) and sulphatereducing Desulfatibacillum alkenivorans AK-01 (Callaghan et al., 2008). This has led to the use
3
of assA sequences as diagnostic markers for anaerobic n-alkane degradation (Acosta-González et al., 2013) and detection of succinylated ‘signature metabolites’ from ASS activity as a priori 45
evidence for n-alkane degradation under nitrate-reducing or sulfidogenic conditions (e.g., Gieg et al., 2010). However, rigorous examination of methanogenic n-alkane-degrading cultures usually fails to detect these characteristic metabolites (Callaghan et al., 2010; Aitken et al. 2013) even though they may be found at low concentrations in the environment (Gieg et al., 2010). This has fuelled speculation that alternative mechanisms (and genes) are used under methanogenic
50
conditions to activate n-alkanes (Aitken et al., 2013) and/or that the intermediates are neither excreted to the environment nor accumulate, but rather are metabolized intracellularly and syntrophically. Even less is known about anaerobic degradation of iso- and cyclo-alkanes, with few reports of sulphidogenic and/or methanogenic degradation (Rios-Hernandez et al., 2003; Townsend et al., 2004; Prince & Suflita, 2007; Tan et al., 2013) including detection of putative
55
iso-alkane metabolites analogous to n-alkane metabolites and the presence of assA homologues in methanogenic cultures (Abu Laban et al., 2015a). We are particularly interested in anaerobic degradation of short-chain n-, iso- and cycloalkanes because oil sands (historically, “tar sands”) tailings ponds (OSTP) in northern Alberta (Canada) represent engineered environments heavily impacted by such hydrocarbons. These
60
enormous impoundments store aqueous colloidal suspensions of clays, bitumen and light hydrocarbon solvents. Diverse indigenous microbiota (Penner and Foght, 2010; Siddique et al., 2012; Ramos-Padrón et al. 2011) degrade certain components of the solvents to produce methane (Siddique et al., 2006; 2007) or sulphides (Ramos-Padrón et al. 2011; Stasik et al., 2014) depending on electron acceptor availability in situ. Thus, anaerobic hydrocarbon degradation is
65
an important component of OSTP management and long-term reclamation strategies.
4
Mildred Lake Settling Basin, the largest and oldest OSTP operated by Syncrude Canada Ltd. contains small proportions of solvent (naphtha, comprising n-, iso- and cyclo-alkanes of ~C6-C10, plus monoaromatic hydrocarbons); cultures enriched from this OSTP degrade shortchain C6-C10 n-alkanes within weeks (Siddique et al., 2006) and C14-C18 n-alkanes within months 70
(Siddique et al., 2011), whereas the C5-C8 iso-alkanes and cycloalkanes are more recalcitrant (Tan et al., 2013; Abu Laban et al., 2015a) and some components persist for years (T. Siddique et al., in preparation). Thus, resistant aliphatic hydrocarbons may accumulate and contribute to toxicity as tailings are deposited in OSTP, or may slowly biodegrade over years or decades. In the latter case such substrates may sustain ‘legacy’ activity in situ, producing mobile water-
75
soluble metabolites and/or greenhouse gas or hydrogen sulphide emissions. Therefore it is important to understand biodegradation of such alkanes for long term environmental management of OSTP and prediction of toxicity and greenhouse gas emissions (Siddique et al., 2008) during tailings storage and eventual site reclamation. Some OSTP including Mildred Lake Settling Basin historically have been or currently
80
are sulphidogenic (Stasik et al., 2014; Ramos-Padrón et al. 2011). Therefore, the ability of the indigenous tailings microbiota to degrade diluent alkanes under both conditions is of interest: the rapidity of the response to a new electron acceptor and any changes in community structure would provide insight into the mechanism(s) of anaerobic alkane degradation in situ. Thus, methanogenic cultures derived from Mildred Lake Settling Basin were enriched on a mixture of
85
n-, iso- and cyclo-alkanes then amended with sulphate while monitoring substrate depletion, metabolite formation and end product accumulation. Community structure was monitored using 16S rRNA gene pyrosequencing, and expression of functional genes (assA and bssA) was determined by generating and sequencing cDNA. The results reflect the presence of a diverse,
5
metabolically dynamic and versatile microbial community. 90
Materials and methods Enrichment cultures Mature fine tailings from Mildred Lake Settling Basin were used to establish a methanogenic short-chain alkane-degrading enrichment culture (SCADC; Tan et al. 2013), as described in 95
Supporting Information (Fig. S1). Briefly, 37 mL of SCADC was used to inoculate each of five sealed 158-mL serum bottles containing 37 mL methanogenic medium (Widdel and Bak 1992) with a headspace of 30% CO2, balance N2. A filter-sterilized C6-C10 alkane mixture comprising equal volumes of ‘hexanes’ (CAS 110-54-3, Fisher Scientific), n-C7 (CAS 142-82-5, >97% purity, Fisher Scientific), n-C8 (CAS 111-65-9, >98% purity, Sigma-Aldrich) and n-C10 (CAS
100
124-18-5, >99% purity, Sigma-Aldrich) was added to three of the cultures to 0.1 vol% final concentration in the medium. The ‘hexanes’ comprised n-C6 (62%) plus the C6 isomers 2methylpentane (3%), 3-methylpentane (16%) and methylcyclopentane (19%), as determined by relative peak area using gas chromatography with mass spectrometry (GC-MS) (Tan et al., 2013). The remaining two cultures did not receive alkanes to control for metabolism of
105
endogenous substrates in tailings. Using the same inoculum, seven single cultures were also prepared with individual carbon sources: ‘hexanes’, n-C7, n-C8, n-C10, 2-methylpentane (CAS 107-83-5, ≥95% purity, Fluka, USA) or methylcyclopentane (CAS 96-37-7, 97% purity, SigmaAldrich), or without amendment. Duplicate 400-mL methanogenic cultures inoculated with SCADC parent culture were amended with 0.1 vol% alkane mixture and incubated for 4 months;
110
one was used for RNA isolation and cDNA synthesis and the other for metabolite detection. Methane production in both 400-mL cultures was monitored to verify activity. After ~18 months incubation the 75-mL alkane-degrading cultures were pooled and used
6
to inoculate (10 vol%) new triplicate 75-mL cultures in methanogenic medium supplemented with 15 mM sodium sulphate plus 0.1 vol% alkane mixture, to determine the composition and 115
activity of the community under sulfidogenic conditions. All cultures were prepared under 30% O2-free CO2–balance N2 headspace with positive pressure to preclude introduction of atmospheric O 2, and were incubated stationary in the dark at ~28 ºC with gentle manual mixing by inversion once per week prior to sample collection.
120
Analysis of methane, sulphide and volatile hydrocarbons in culture bottles Methane was measured by analyzing 50 µL of culture bottle headspace using a gas chromatograph with a flame-ionization detector (Siddique et al., 2006). The maximum theoretical methane yield was calculated using the Symons and Buswell equation (Roberts, 2002), based on the calculated mass of hydrocarbons added to the cultures (Supporting
125
Information, Table S1). Soluble sulphide production in sulphate-amended cultures was determined using a colourimetric methylene blue method (Cline, 1969). For residual volatile hydrocarbon detection by GC-MS, 50 µL-samples of culture headspace were analyzed as described by Abu Laban et al. (2015a), except that carrier gas flow was 0.8 mL min-1 and the temperature ramp was 5 ºC min-1. Headspace hydrocarbon depletion
130
was calculated as previously described (Tan et al., 2013) based on the method of Prince & Suflita (2007). Briefly, the percentage of target compounds remaining in the headspace was calculated based on the following equation: % of headspace hydrocarbon = [(Asample/Csample)/(Auninoculated sterile medium/Cuninoculated sterile medium)] x 100, where A and C represent the headspace abundance of target analytes and an internal standard, respectively.
135
Detection of putative metabolites
7
Culture fluid (2–10 mL from the triplicate 75-mL cultures or 50-200 mL from the 400-mL methanogenic culture; Fig. S1) was removed from sealed bottles by using a sterile needle and syringe, and 4-fluoro-1-naphthoic acid (Sigma-Aldrich) dissolved in ethyl acetate was added as a 140
surrogate extraction and derivatization standard. Samples plus parallel abiotic controls containing only doubly-distilled water were either immediately acidified to pH
Anaerobic alkane biodegradation by cultures enriched from oil sands tailings ponds involves multiple species capable of fumarate addition
BoonFei Tan1, Kathleen Semple and Julia Foght* Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9
*Corresponding author: Julia Foght phone: +1 780.492.3279 fax: +1 780.492.0234 email: [email protected]
1
Current address: Center for Environmental Sensing and Modeling, Singapore-MIT Alliance for Research and Technology, 1 Create Way, #09-03 Create Tower, Singapore 138602.
Running title: Anaerobic alkane biodegradation Keywords: aliphatic hydrocarbon degradation / methanogenesis / sulphidogenesis
1
Abstract A methanogenic short-chain alkane-degrading culture (SCADC) was enriched from oil sands tailings and transferred several times with a mixture of C6, C7, C8 and C10 n-alkanes as the predominant organic carbon source, plus 2-methylpentane, 3-methylpentane and 5
methylcyclopentane as minor components. Cultures produced ~40% of the maximum theoretical methane during 18 months incubation while depleting the n-alkanes, 2-methylpentane and methylcyclopentane. Substrate depletion correlated with detection of metabolites characteristic of fumarate activation of 2-methylpentane and methylcyclopentane but not n-alkane metabolites. During active methanogenesis with the mixed alkanes, reverse-transcription PCR confirmed the
10
expression of functional genes (assA and bssA) associated with hydrocarbon addition to fumarate. Pyrosequencing of 16S rRNA genes amplified during active alkane degradation revealed enrichment of Clostridia (particularly Peptococcaceae) and methanogenic Archaea (Methanosaetaceae and Methanomicrobiaceae). Methanogenic cultures transferred into medium containing sulphate produced sulphide, depleted n-alkanes and produced the corresponding
15
succinylated alkane metabolites, but were slow to degrade 2-methylpentane and methylcyclopentane; these cultures were enriched in Deltaproteobacteria rather than Clostridia. 3-Methylpentane was not degraded by any cultures. Thus, nominally methanogenic oil sands tailings harbour dynamic and versatile hydrocarbon-degrading fermentative syntrophs and sulphate reducers capable of degrading n-, iso- and cyclo-alkanes by addition to fumarate.
20
2
20
Introduction Aliphatic hydrocarbons including n-, iso- and cyclo-alkanes are major constituents of crude oil and its refined products and are significant contaminants in petroleum-impacted environments. Whereas alkane biodegradation under aerobic conditions has been well known for decades (Van 25
Hamme et al., 2003) and metabolism under nitrate- and sulphate-reducing conditions has been described more recently (reviewed by Mbadinga et al., 2011), the key microbes, biodegradation pathways, metabolites, and functional genes involved in methanogenic alkane degradation have been cryptic or controversial (Aitken et al., 2013; Callaghan 2013; Embree et al. 2014). Recent reports (e.g., Tan et al. 2014a, b, c; Abu Laban 2015a) have begun to expand the repertoire of
30
microbes and genes associated with methanogenic alkane-degrading cultures. Enzymatic activation of aliphatic hydrocarbons by addition to fumarate is the best described and most widely documented mechanism for initiating anaerobic alkane degradation under nitrate- and sulphate-reducing conditions (Agrawal & Gieg, 2013, Callaghan, 2013). The glycyl radical enzyme alkylsuccinate synthase (ASS: also called methylalkylsuccinate synthase,
35
MAS) and its counterparts responsible for aromatic hydrocarbon activation [benzylsuccinate synthase (BSS) and naphthyl-2-methylsuccinate synthase (NMS)], have been described and reviewed (Mbadinga et al. 2011; Callaghan 2013; Agrawal and Gieg, 2013). The requisite genes, particularly those encoding the alpha-subunit, e.g., assA, have been detected in contaminated environments (Acosta-González et al., 2013), oil field fluids (Zhou et al., 2012), enrichment
40
cultures (Callaghan et al., 2010; Aitken et al., 2013) and model n-alkane-degrading organisms such as nitrate-reducing ‘Aromatoleum’ sp. HxN1 (Grundmann et al., 2008) and sulphatereducing Desulfatibacillum alkenivorans AK-01 (Callaghan et al., 2008). This has led to the use
3
of assA sequences as diagnostic markers for anaerobic n-alkane degradation (Acosta-González et al., 2013) and detection of succinylated ‘signature metabolites’ from ASS activity as a priori 45
evidence for n-alkane degradation under nitrate-reducing or sulfidogenic conditions (e.g., Gieg et al., 2010). However, rigorous examination of methanogenic n-alkane-degrading cultures usually fails to detect these characteristic metabolites (Callaghan et al., 2010; Aitken et al. 2013) even though they may be found at low concentrations in the environment (Gieg et al., 2010). This has fuelled speculation that alternative mechanisms (and genes) are used under methanogenic
50
conditions to activate n-alkanes (Aitken et al., 2013) and/or that the intermediates are neither excreted to the environment nor accumulate, but rather are metabolized intracellularly and syntrophically. Even less is known about anaerobic degradation of iso- and cyclo-alkanes, with few reports of sulphidogenic and/or methanogenic degradation (Rios-Hernandez et al., 2003; Townsend et al., 2004; Prince & Suflita, 2007; Tan et al., 2013) including detection of putative
55
iso-alkane metabolites analogous to n-alkane metabolites and the presence of assA homologues in methanogenic cultures (Abu Laban et al., 2015a). We are particularly interested in anaerobic degradation of short-chain n-, iso- and cycloalkanes because oil sands (historically, “tar sands”) tailings ponds (OSTP) in northern Alberta (Canada) represent engineered environments heavily impacted by such hydrocarbons. These
60
enormous impoundments store aqueous colloidal suspensions of clays, bitumen and light hydrocarbon solvents. Diverse indigenous microbiota (Penner and Foght, 2010; Siddique et al., 2012; Ramos-Padrón et al. 2011) degrade certain components of the solvents to produce methane (Siddique et al., 2006; 2007) or sulphides (Ramos-Padrón et al. 2011; Stasik et al., 2014) depending on electron acceptor availability in situ. Thus, anaerobic hydrocarbon degradation is
65
an important component of OSTP management and long-term reclamation strategies.
4
Mildred Lake Settling Basin, the largest and oldest OSTP operated by Syncrude Canada Ltd. contains small proportions of solvent (naphtha, comprising n-, iso- and cyclo-alkanes of ~C6-C10, plus monoaromatic hydrocarbons); cultures enriched from this OSTP degrade shortchain C6-C10 n-alkanes within weeks (Siddique et al., 2006) and C14-C18 n-alkanes within months 70
(Siddique et al., 2011), whereas the C5-C8 iso-alkanes and cycloalkanes are more recalcitrant (Tan et al., 2013; Abu Laban et al., 2015a) and some components persist for years (T. Siddique et al., in preparation). Thus, resistant aliphatic hydrocarbons may accumulate and contribute to toxicity as tailings are deposited in OSTP, or may slowly biodegrade over years or decades. In the latter case such substrates may sustain ‘legacy’ activity in situ, producing mobile water-
75
soluble metabolites and/or greenhouse gas or hydrogen sulphide emissions. Therefore it is important to understand biodegradation of such alkanes for long term environmental management of OSTP and prediction of toxicity and greenhouse gas emissions (Siddique et al., 2008) during tailings storage and eventual site reclamation. Some OSTP including Mildred Lake Settling Basin historically have been or currently
80
are sulphidogenic (Stasik et al., 2014; Ramos-Padrón et al. 2011). Therefore, the ability of the indigenous tailings microbiota to degrade diluent alkanes under both conditions is of interest: the rapidity of the response to a new electron acceptor and any changes in community structure would provide insight into the mechanism(s) of anaerobic alkane degradation in situ. Thus, methanogenic cultures derived from Mildred Lake Settling Basin were enriched on a mixture of
85
n-, iso- and cyclo-alkanes then amended with sulphate while monitoring substrate depletion, metabolite formation and end product accumulation. Community structure was monitored using 16S rRNA gene pyrosequencing, and expression of functional genes (assA and bssA) was determined by generating and sequencing cDNA. The results reflect the presence of a diverse,
5
metabolically dynamic and versatile microbial community. 90
Materials and methods Enrichment cultures Mature fine tailings from Mildred Lake Settling Basin were used to establish a methanogenic short-chain alkane-degrading enrichment culture (SCADC; Tan et al. 2013), as described in 95
Supporting Information (Fig. S1). Briefly, 37 mL of SCADC was used to inoculate each of five sealed 158-mL serum bottles containing 37 mL methanogenic medium (Widdel and Bak 1992) with a headspace of 30% CO2, balance N2. A filter-sterilized C6-C10 alkane mixture comprising equal volumes of ‘hexanes’ (CAS 110-54-3, Fisher Scientific), n-C7 (CAS 142-82-5, >97% purity, Fisher Scientific), n-C8 (CAS 111-65-9, >98% purity, Sigma-Aldrich) and n-C10 (CAS
100
124-18-5, >99% purity, Sigma-Aldrich) was added to three of the cultures to 0.1 vol% final concentration in the medium. The ‘hexanes’ comprised n-C6 (62%) plus the C6 isomers 2methylpentane (3%), 3-methylpentane (16%) and methylcyclopentane (19%), as determined by relative peak area using gas chromatography with mass spectrometry (GC-MS) (Tan et al., 2013). The remaining two cultures did not receive alkanes to control for metabolism of
105
endogenous substrates in tailings. Using the same inoculum, seven single cultures were also prepared with individual carbon sources: ‘hexanes’, n-C7, n-C8, n-C10, 2-methylpentane (CAS 107-83-5, ≥95% purity, Fluka, USA) or methylcyclopentane (CAS 96-37-7, 97% purity, SigmaAldrich), or without amendment. Duplicate 400-mL methanogenic cultures inoculated with SCADC parent culture were amended with 0.1 vol% alkane mixture and incubated for 4 months;
110
one was used for RNA isolation and cDNA synthesis and the other for metabolite detection. Methane production in both 400-mL cultures was monitored to verify activity. After ~18 months incubation the 75-mL alkane-degrading cultures were pooled and used
6
to inoculate (10 vol%) new triplicate 75-mL cultures in methanogenic medium supplemented with 15 mM sodium sulphate plus 0.1 vol% alkane mixture, to determine the composition and 115
activity of the community under sulfidogenic conditions. All cultures were prepared under 30% O2-free CO2–balance N2 headspace with positive pressure to preclude introduction of atmospheric O 2, and were incubated stationary in the dark at ~28 ºC with gentle manual mixing by inversion once per week prior to sample collection.
120
Analysis of methane, sulphide and volatile hydrocarbons in culture bottles Methane was measured by analyzing 50 µL of culture bottle headspace using a gas chromatograph with a flame-ionization detector (Siddique et al., 2006). The maximum theoretical methane yield was calculated using the Symons and Buswell equation (Roberts, 2002), based on the calculated mass of hydrocarbons added to the cultures (Supporting
125
Information, Table S1). Soluble sulphide production in sulphate-amended cultures was determined using a colourimetric methylene blue method (Cline, 1969). For residual volatile hydrocarbon detection by GC-MS, 50 µL-samples of culture headspace were analyzed as described by Abu Laban et al. (2015a), except that carrier gas flow was 0.8 mL min-1 and the temperature ramp was 5 ºC min-1. Headspace hydrocarbon depletion
130
was calculated as previously described (Tan et al., 2013) based on the method of Prince & Suflita (2007). Briefly, the percentage of target compounds remaining in the headspace was calculated based on the following equation: % of headspace hydrocarbon = [(Asample/Csample)/(Auninoculated sterile medium/Cuninoculated sterile medium)] x 100, where A and C represent the headspace abundance of target analytes and an internal standard, respectively.
135
Detection of putative metabolites
7
Culture fluid (2–10 mL from the triplicate 75-mL cultures or 50-200 mL from the 400-mL methanogenic culture; Fig. S1) was removed from sealed bottles by using a sterile needle and syringe, and 4-fluoro-1-naphthoic acid (Sigma-Aldrich) dissolved in ethyl acetate was added as a 140
surrogate extraction and derivatization standard. Samples plus parallel abiotic controls containing only doubly-distilled water were either immediately acidified to pH
