Scholarly article on topic 'Insights into the Anaerobic Biodegradation Pathway of n-Alkanes in Oil Reservoirs by Detection of Signature Metabolites'

Insights into the Anaerobic Biodegradation Pathway of n-Alkanes in Oil Reservoirs by Detection of Signature Metabolites Academic research paper on "Biological sciences"

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Academic research paper on topic "Insights into the Anaerobic Biodegradation Pathway of n-Alkanes in Oil Reservoirs by Detection of Signature Metabolites"


OPEN Insights into the Anaerobic Biodegradation Pathway of n-Alkanes in Oil Reservoirs by

Received: 16 April 2014 a m

A epted: 6 Marh 205 Detection of Signature Metabolites

Published: 13 May 2015

Xin-Yu Bian1'2, Serge Maurice Mbadinga1'2, Yi-Fan Liu1'2, Shi-Zhong Yang1'2, Jin-Feng Liu1'2, Ru-Qiang Ye1'2, Ji-Dong Gu3 & Bo-Zhong Mu1'2

Anaerobic degradation of alkanes in hydrocarbon-rich environments has been documented and different degradation strategies proposed, of which the most encountered one is fumarate addition mechanism, generating alkylsuccinates as specific biomarkers. However, little is known about the mechanisms of anaerobic degradation of alkanes in oil reservoirs, due to low concentrations of signature metabolites and lack of mass spectral characteristics to allow identification. In this work, we used a multidisciplinary approach combining metabolite profiling and selective gene assays to establish the biodegradation mechanism of alkanes in oil reservoirs. A total of twelve production fluids from three different oil reservoirs were collected and treated with alkali; organic acids were extracted, derivatized with ethanol to form ethyl esters and determined using GC-MS analysis. Collectively, signature metabolite alkylsuccinates of parent compounds from Ci to C8 together with their (putative) downstream metabolites were detected from these samples. Additionally, metabolites indicative of the anaerobic degradation of mono- and poly-aromatic hydrocarbons (2-benzylsuccinate, naphthoate, 5,6,7,8-tetrahydro-naphthoate) were also observed. The detection of alkylsuccinates and genes encoding for alkylsuccinate synthase shows that anaerobic degradation of alkanes via fumarate addition occurs in oil reservoirs. This work provides strong evidence on the in situ anaerobic biodegradation mechanisms of hydrocarbons by fumarate addition.

Petroleum (crude oil) is a complex mixture containing thousands of chemicals, mainly of hydrocarbons, making up 80% of the oil chemical constituents. Hydrocarbons are relatively less reactive due to strong and localized C-C bond, and lack of reactive functional group1'2. Microorganisms can use hydrocarbons as the sole source of carbon and energy under either aerobic or anaerobic conditions3-7. The current understanding is that the majority of the oils degraded are a result of the activities of anaerobes living in subsurface environments under anaerobic conditions8-10. Indeed, diverse physiological groups of microorganisms (fermentative, iron-, nitrate-, sulfate-reducers, syntrophs, and methanogens, for instance) have been isolated from and/or detected in production fluids of oil reservoirs2,11. Anaerobic enrichment cultures capable of degrading and converting hydrocarbons and crude oils were also established with oil reservoir production fluids12-17. The research provides new insights into the biochemical capabilities of microorganisms originally from oil reservoirs and their involvement in the anaerobic degradation of oil hydrocarbons via detection of genes encoding for specific metabolic pathways (i.e., PCR amplification of genes assA/masD and bssA encoding for the fumarate addition pathways12-16,18) (for review see Refs.

1State Key Laboratory of Bioreactor Engineering and Institute of Applied Chemistry, East China University of

Science and Technology, Shanghai, P.R. China. 2Shanghai Collaborative Innovation Center for Biomanufacturing

Technology, Shanghai 200237, PR. China. 3School of Biological Sciences, The University of Hong Kong, Pokfulam Road, Hong Kong, P.R. China. Correspondence and requests for materials should be addressed to J-D. G. (e-mail: or B-Z. M. (email:

assABC fumarate addition


C- skeleton rearrangement decarboxylation p-oxidation

Figure 1. Proposed fumarate addition mechanism in anaerobic degradation of alkanes.

H1 H2 H3 H4 J1 J2 J3 J4 J5 J6 X1 X2

Temperature (°C) 37~45 80~90 21 32

Na + (mg/L) 1148.3 1173.9 868.1 962.2 9577.2 4235.3 8750.2 3909.2 3801.4 6218.7 4196 5399.0

NH4 (mg/L) 375.0 65.5 1109.7 1040.3 963.9 44.0 n. d.* 1139.9 85.5 61.1 n. d. n. d.

Cl- (mg/L) 306.7 428.3 445.9 423.7 21394.5 4500 13178.7 8418.5 6825.8 12575.17 2000 5336

SO2- (mg/L) 91.0 80.7 n. d. 42.6 2676.2 6513.5 82.3 335.3 2050.6 2392.4 124.8 7.7

Table 1. Physicochemical parameters of the samples collected from the twelve oil reservoir samples. *n. d.: not detectable

19,20). Knowledge about biodegradation of petroleum hydrocarbons in oil reservoir systems will advance our understanding of the biochemical processes catalyzed by specific microorganisms and also enrich the information on the global geobiochemical carbon cycle. Sulfidogenic and denitrifying bacterial strains, such as Desulfatibacillum alkenivorans AK-01 and Aromatoleum HxN1 serve as models to investigate the fumarate addition biochemical pathway and alkylsuccinate synthase (or methylalkylsuccinate synthase) in the metabolism of oil hydrocarbons21-24. The biochemical pathways are well documented and described here in Fig. 24-30. In the fumarate addition pathway, n-alkanes are initially activated by addition to the double bond of fumarate at the subterminal21,25-28,30 or terminal31 (with propane) carbon, producing 2-(1-methylalkyl)succinates (or 2-alkylsuccinates). This process is catalyzed by alkylsuccinate synthase. Further degradation of 2-(1-methylalkyl)succinates involves carbon skeleton re-arrangement, de-carboxylation, and ^-oxidations21. The initial products, 2-(1-methylalkyl)succinates, are commonly considered biochemical markers indicating the occurrence of the fumarate addition mechanism in the initial step19. Also, some cycloalkanes, iso-alkanes, aromatic hydrocarbons and polycyclic aromatic hydrocarbons have been reported being degraded anaerobically via addition to fumarate (such as, the detection of 2-cyclopentylsuccinate and/or 2-benzylsuccinate indicates fumarate addition pathway occurring in the anaerobic degradation of cyclopentane or toluene, respectively)26,29,32-36. Therefore, hydrocarbon-derived succinates can be used as biochemical indicators for hydrocarbon-based metabolic process, which is a powerful piece of evidences for in situ biodegradation mechanisms carried out biochemically through microbial activity19,37.

Metabolite profiling32,36,38,39 and assA/masD gene40-42 analysis of hydrocarbons-contaminated environments showed the occurrence of succinate metabolites, indicative of in situ biological activities. Various hydrocarbon-derived succinate analogues with side chains ranging from C3 to C11 were found in hydrocarbon-contaminated environments according to the summary available19. Non-saturated fatty acids with 2 mass units less than the corresponding alkylsuccinates were also detected36,38,39. Thus, evidence of anaerobic degradation of alkanes in such systems is well recognized. However only a limited number of metabolite profiles has been reported directly from oil reservoir production fluids. Studies on samples from Alaska North Slope oilfield showed that C1-C4 alkylsuccinates together with putative downstream metabolites were detected in production fluids17,43.

In this work, we used a multidisciplinary approach that combines metabolite profiling and functional gene (assA/masD) assays to investigate the specific biochemical mechanism in production fluids of twelve oil reservoirs obtained from three distinct oilfields in China. Our results, combined with the earlier works17,43 illustrate that anaerobic degradation of alkanes via the fumarate addition pathway appears to be a common initial activation strategy in different oil reservoir systems.


Physicochemical characteristics of oil reservoir production fluids. The characteristics of the investigated oil reservoir production fluids are summarized in Table 1. GC analyses of alkanes in the 12 oil samples are also provided in Supplementary Materials, (Figs. S2.1-S2.12). The amount of Na+, NH4+ and Cl- of samples H1 to H4 ranged from 868.1 mg/L to 1148.3 mg/L, 65.5 mg/L to 1040.3 mg/L, and 306.7 mg/L to 445.9 mg/L, respectively. The SO42- was from non-detectable in H3 to as high as 91.0 mg/L in H1. For samples J1 to J6, the concentration of Na+, NH4+, Cl-and SO42- was between 3801.4 mg/L and

H1 H2 H3 H4 J1 J2 J3 J4 J5 J6 X1 X2


C1 +

C2 + + + + + + +

C3 + + + + + + +

C4 + + + + + +

C5 + + + + + +

C6 + + +

C7 + +

C8 + +

B enzylsuccinate +

Products of the carbon skeleton rearrangement

Ethylmalonate +

Butylmalonate + +

2-(methylpentyl)malonate +

Metabolite of naphthalene and/or methylnaphthalene

Naphthoate + + + + +

5,6,7,8-tetrahydronaphthoate + +


Formate +

Acetate + + + + +

Propionate + +

2- methylpropionate +

Butyrate + + + +

1-methylbutyrate + + +

Hydroxycaproate + +

Octanoate +

4-octenoate + + + +

3-nonenoate + + + + +

Nonanoate +

Laurate + + +

9-hexadecenoate + +

Myristate + + + + + + +

3-hydroxytridecanoate +

Palmitate + + + + + + + + + + + +

Oleate + + +

Stearate + + + +

Table 2. Signature metabolites of anaerobic alkanes degradation detected in production fluid samples. "+": detected.

9577.2 mg/L, 44.0 mg/L and 1139.9 mg/L, 4500 mg/L and 21394.5 mg/L and 82.3 mg/L and 6513.5 mg/L, respectively. For samples X1 and X2, the amount of Na+, Cl-and SO|- was between 4196 mg/L and 5399.0 mg/L, 2000 mg/L and 5336 mg/L, and 7.7 mg/L and 124.8 mg/L, respectively.

Signature metabolites of anaerobic degradation of hydrocarbons in oil reservoirs. Twelve production fluids collected from three different oil reservoir systems were analyzed for the presence of specific chemical metabolites showing evidence of in situ anaerobic biodegradation of alkanes and the biochemical mechanism involved. The mass spectra of organic extracts after derivatization with ethanol (for non-volatile organic acids) and n-butanol (for volatile organic acids) were also obtained and the results are given in Table 2.

Sample H4 was randomly chosen as an example to show its diverse alkylsuccinates and the corresponding representative mass spectra (Table 2). Fig. 2a shows the total ion chromatogram of fatty acids extracted from production fluids of H4. According to our previous study44, m/z 128 and 174 were first selected as potential indicators of alkylsuccinates and a cluster of peaks with retention time from 10.00 to 30.00 min was evident in the selected ion chromatograms (Fig. 2b). A detailed examination of the existence of fragments m/z M+-45 and M+-87 was also undertaken. The mass spectra of some detected

Figure 2. (a) The total ion chromatogram of fatty acid ethyl esters from production fluid H4; (b) Selected ion chromatogram of ethyl esters from production fluid H4 containing fragments m/z 174 and 128 (The most abundant peak in (a) is 1-chlorohexadecane that serves as the internal standard).

diethyl alkylsuccinates in sample H4 are shown in Fig. 3. The mass spectra of other fatty acids, including diethyl and dibutyl esters, are provided in Figs. S1.13-S1. 29.

Comparison of retention times and mass spectral characteristics of chemically synthesized alkyl-succinates44, a suite of (putative) alkyl-substituted succinates of parent n-alkanes with chain length from C1 to C8 (Fig. 4 and Table 2) were detected by GC-MS analysis. In addition, 2-benzylsuccinate and 2-naphthoate known as indicators of anaerobic degradation of mono- or poly-aromatic hydrocarbons were also detected in some of the samples in this study (Fig. 4b, c and Table 2).

Putative downstream metabolites of the fumarate addition pathway. Alkylmalonates (C2, C4 and C6), known as downstream degradation intermediates of alkyl-succinates via C-skeleton rearrangement, were also observed in samples H3, H4, J2 and J4. Volatile fatty acids such as formate, acetate, propionate, butyrate, their branched derivatives and long-chain fatty acids (stearate, palmitate and myristate) were also found in some samples. These fatty acids are also produced during the anaerobic

Figure 3. Mass spectra of some detected diethyl alkylsuccinates in sample H4.

degradation of alkanes. Remarkably, 5, 6, 7, 8-tetrahydro-naphthoate as the downstream metabolite of 2-naphthoate was detected in sample H3 and J4 (Table 1). The metabolic pathways that could produce

Further degradation

Figure 4. Putative anaerobic degradation pathway of hydrocarbons: (a) alkane, (b) toluene, (c) naphthalene or 2-methylnaphthalene. Detected organic acids are marked in red, and putative metabolites of anaerobic degradation of alkanes via fumarate addition are listed at four stages, namely fumarate addition, C-skeleton re-arrangement, decarboxylation and further ^-oxidation.

5, 6, 7, 8-tetrahydro-naphthoate and 2-naphthoate from naphthalene and/or 2-methyl naphthalene are given in Fig. 4c9,45.

Investigation of alkylsuccinate/2-(1-methylalkyl)succinate synthase alpha-subunit (assA/ masD) gene. The detection of alkylsuccinates in most of the samples indicates that anaerobic degradation of alkanes occurs in oil reservoir systems via the fumarate addition mechanism. To provide additional evidence on the microbial communities capable of anaerobic degradation of alkanes via fumarate addition in these samples, genomic DNAs extracted from the production fluid samples were PCR amplified for the presence of functional genes (assA/masD) encoding for enzymes that initiate anaerobic degradation of alkanes. Clone libraries were then established from eleven of the twelve production fluid samples and all the cloned sequences were affiliated with assA/masD genes (Fig. 5), but no expected amplification band could be obtained from DNA extract of sample H2. Subsequent analysis was carried out at the protein level of deduced assA/masD gene sequences. OTUs H3-assA27 and H4-assA20 were 91% similar to an assA/masD gene sequence (AEI52403) obtained from a meth-anongenic alkane-degrading enrichment culture. OTUs H3-assA27, H4-assA20 also shared at least 85% identity with assA/masD sequence from Smithella ME-146,47. OTUs J2-assA112, J5-assA9 and X1-assA24 were highly related (96-98% identity) to assA/masD sequence from Smithella ME-146,47. OTUs J1-assA36 and H1-assA123 showed close relationship to assA/masD gene sequence (AGC24806) from River Tyne sediment microcosms amended with crude oil18. OTU X1-assA32 was a close relative (98% identity) to assA/masD gene sequence (ADJ51090) retrieved from a methanogenic paraffin degrading enrichment40. X1-assA22 clustered with 75% identity to assA/masD gene sequences obtained from fuel incubation48. J4-assA93 appeared to be far related to any assA/masD gene sequence available in the GenBank database. OTUs J3-assA77, J4-assA102, J5-assA10, J6-assA21, H1-assA122, H3-assA28 and H4-assA18 clustered together and were moderately related to the known assA/masD gene sequences; which could indicate the existence of a potentially "new clade" of assA/masD based on gene sequences (Fig. 5).

OTUs J4-assA91, X1-assA31, X1-assA34 and X2-assA64 shared at least 72 to 94% identity to assA/masD gene sequences previously detected in samples obtained from suphidogenic anoxic sediments incubated with dodecane49, oil sands tailings (accession number AIB50974) and sediment from a hydrocarbon seep in the Guaymas Basin, Gulf of California41. Finally, OTUs X1-assA41 and X2-assA66 were most closely related (90-96% identity) to assA/masD gene sequences from hydrocarbon-impacted aquifers near Fort Lupton, Colorado40.


Oil reservoirs represent specific environments in which microorganisms, especially anaerobes, have been implicated in the formation of the heavy oil that is found in geographically distinct reservoirs around

AET09978, Shengli oil field, methanogenic hexadecane consortium clone H15 AEI52403, methanongenic alkane degrading enrichment culture clone assA-63 gAET09982, Shengli oil field, methanogenic hexadecane consortium clone H20 H3-assA27 (KM229476) H4-assA20 (KM251744) J2-assA112 (KM251770) J5-assA9 (KM251772)

X1-assA24 (KM229512) AWGX01000974, Smithella sp. ME-1 (alkane-degrading methanogenic community)

-AIB50972, oil sands tailings clone SCADC_assA3

AHI85732, Smithella sp. enrichment culture clone SCADC ADJ51092, Fort Lupton, hydrocarbon-impacted environment Fort_Lupton_OTU2 J1-assA36 (KM251757) H1-assA123 (KM251716) 75' AGC24806, River Tyne sediment, microcosms ammended with crude oil clone 22_OTU3

- X1 -assA32 (KM229496)

- ADJ51090, methanogenic paraffin degrading enrichment clone SDB_0TU1 J4-assA93 (KM229491)

1— CDJ79983,suphidogenic anoxic sediments incubated with dodecane clone GB-LC_824 1— AGC24789, River Tyne sediment, microcosms ammended with crude oil clone 5_OTU3 AFF59639, fuel incubation clone AC42KL3a01 AFF59640, fuel incubation clone AC42KL3a04 X1-assA22 (KM229511) r J4-assA102(KM229447) J6-assA21 (KM251778) H4-assA18 (KM251747) J5-assA10 (KM251773) H3-assA28 (KM229477) - H1-assA122 (KM251722) J3-assA77 (KM229435) lOOj YP_002430896, Desulfatibacillum alkenivorans AK-01 assA1 WP_028315530, Desulfatibacillum aliphaticivorans assA YP_002431360,Desulfatibacillum alkenivorans AK-01 assA2 AGJ00452, marine sediment contaminated with crude oil clone M-OIL002 ADJ51097, Desulfoglaeba alkanexedens ALDC assA J4-assA91 (KM229490) CDJ79964, suphidogenic anoxic sediments incubated with dodecane clone AMV-LC_523 AIB50974, oil sands tailings clone SCADC_assA4 100 r X1 -ass A31 (KM229495) X1 -assA34 (KM229498)

X2-assA64 (KM251788)

AFX62321, Guaymas hydrocarbon seep clone BGM02 CBK27727, Aromatoleum sp. OcN1 masD CA003074, Azoarcus sp. masD 1001KCW91781, oil sands tailings pond Peptococcaceae bacterium SCADC1_2_3

AIB50971, oil sands tailings clone SCADC_assA2 WP_027352796, Desulfosarcina sp. BuS5 X1-assA41 (KM229504)

ADJ51105, Fort Lupton, hydrocarbon-impacted environment Fort_Lupton_OTU1 X2-assA66 (KM251787)

ADJ51106, Fort Lupton, hydrocarbon-impacted environment Fort_Lupton_OTU2 95; CA072219,.delta proteobacterium NaphS2

CA072222, delta proteobacterium NaphS6 IIITISA

- ADB04297, tar-oil contaminated sediment clone N47

(j) (0

V) U) fO

WP_012647530, Geobacter daltonii WP_004511527, Geobacter metallireducens

WP_011236884, Aromatoleum aromaticum

BAD42366, Magnetospirillum sp. TS-6 AAC38454, Thauera aromatica TutD BAC05501, Thauera sp. DNT-1 bssA

YP_002994045,Thermococcus sibiricus MM 739 PFL AAB89800, Archaeoglobus fulgidus DSM 4304 PFLD

Figure 5. Phylogenetic tree of deduced assA/masD gene sequences amplified from DNAs extracted from the oil reservoir production fluid samples and closely related sequences from the GenBank database. Phylogenetic analyses were conducted with MEGA6 software. The topology of the tree shown was obtained with the neighbor-joining method and the Poisson correction. Values below 70% are not shown. The database was accessed on between July and August 2014. Scale bar = 10% amino acid substitution.

the world8. The presence of anaerobes in such systems has led to the assumption that these microorganisms may be used to recover more energy as natural gas via methanogenic conversion of oil hydrocar-bons50-52. Alkanes represent an important fraction of crude oil and in recent years, studies have been undertaken to understand the activation of these relatively less reactive compounds under anaerobic conditions and the addition of alkanes onto the double bond of fumarate with subsequent formation of alkylsuccinates as a prominent biochemical activation mechanism (for review see Ref 5). The detection of alkylsuccinates in engineered settings and/or environmental samples is indicative of the activity of microorganisms using the fumarate addition mechanism19,37. So far, alkylsuccinates have been found in anaerobic enrichment cultures amended with either alkanes or crude oil15,18 and also in environmental samples obtained from oil-contaminated sites (for review see Ref 19,37), but scarcely reported in samples originating from oil reservoirs. In the present study, metabolite profiles of samples collected from three different oil fields were analyzed using GC-MS, and at the same time, alkylsuccinates as well as putative downstream metabolite alkylmalonates were found in eleven of the twelve samples investigated in the three oilfields. Collectively, these identified metabolites are supportive for the anaerobic activation of alkanes in oil reservoirs via the fumarate addition biochemical pathway. The detection of other alkanoic acids suggests a further degradation of alkylsuccinates in the investigated environments; though these alkanoic acids can have multiple sources.

In addition, 2-benzylsuccinate and naphthoate together with 5,6,7,8-tetrahydro-naphthoate (metabolites produced during the anaerobic degradation of toluene, naphthalene and/or methyl naphthalene) were also identified in samples H3, J1, J4, J6, X1 and X29,34,53. This set of metabolites identified indicates that, besides alkanes, mono- and polycyclic aromatic hydrocarbons were also degraded anaerobically in the oil reservoirs. Therefore, the putative biodegradation pathways of these organic acids were illustrated in Fig. 4a. Generally, at least one compound in each biochemical step was detected in the samples analyzed. It is noteworthy that both 2-(1-methylheptyl)succinate and 2-(methylpentyl)malonate are so far the two largest signature metabolites in terms of molecular weight from alkanes degradation observed in oil reservoirs.

Furthermore, to obtain additional evidence on the microbial communities capable of anaerobic degradation of alkanes via fumarate addition, functional genes assA/masD were PCR amplified from the oil reservoirs samples. Expected DNA bands were obtained successfully in eleven of the twelve samples, and further cloned and sequenced. The results indicate the presence in the oil reservoirs of microorganisms harboring assA/masD gene encoding for enzyme(s) that initiates anaerobic alkane degradation via fuma-rate addition mechanism. Combined with the detection of alkylsuccinates as signature metabolites, our data shows that anaerobic degradation of alkanes via the fumarate addition pathway occurred in the oil reservoirs. The combination of the two methods (metabolite profiling and functional gene amplification) should prove a useful and more comprehensive approach to gain insights into the anaerobic degradation of alkanes (via fumarate addition) in oil reservoirs despite of the fact the occurrence of other degradation strategies cannot be excluded3,18. For example, 2-(1-methylethyl)succinate was indeed detected in H2, but no assA gene products could be amplified from the DNA extracted from this sample, probably due to the specificity and coverage of PCR primers (primers sets used are not "all-inclusive" of "universal" templates) used for DNA amplification.

It is difficult, with limited information, to link the formation of alkylsuccinates in the samples analyzed with specific metabolic processes such as nitrate-, iron-, sulfate-reduction, or methanogenesis (i.e., when and how these specific biomarkers were formed). For instance, several samples (H1, H3, H4, J1, J2, J5 and X1) contained assA/masD gene homologues highly related to those from members of the genus Smithella; a genus that was implicated in the anaerobic degradation of crude oil alkanes under methanogenic conditions46,47,54. However, except for sample X1 (21 °C) in which alkylsuccinates were not detectable, all other samples had temperature above 37 °C; temperature beyond the optimum reported for the growth of Smithella propionica, the only known isolate of the genus Smithella55. Therefore, we speculate that microorganisms carrying assA gene similar to those from members of the Smithella may exist in the oil reservoirs production fluids. On the other hand, assuming that these assA gene sequences were truly from members of the genus Smithella and since, to date, relatives to this genus have not been reported to thrive in high temperature reservoirs, we speculate that they could survive in the cooler part of the reservoirs.

Oil reservoirs are large and complex environments, different from laboratory cultures in tubes and/ or bottles, therefore various metabolic processes may occur in different zones (or depth) of the same reservoir. Previous work showed that anaerobic degradation of alkane (methane to butane) also occurred in the pipeline of Alaskan North Slope oil field43. That is, when sampling from production wells, the information from the geological formation and pipeline infrastructure may be mixed and that should also be considered when interpreting phylogenetic data.

Based on current data from the production fluids, it is impossible to obtain direct evidence on the connection between fumarate addition and methanogenic/sulfate-reducing or other relevant biochemical/physiological processes. However, using a combination of approaches involving biochemical and functional gene profiling simultaneously, this study does suggest that anaerobic degradation of alkanes via fumarate addition pathway occurs in oil reservoirs.


The detection of signature biomarkers of anaerobic degradation of n-alkanes in conjunction with the positive detection of associated alkylsuccinate synthase genes in samples from oil reservoirs supports the occurrence of the fumarate addition pathway in oil reservoirs. Our results, in conjunction with other data, support the hypothesis that fumarate addition mechanism plays an important role in anaerobic alkane transformation in oil reservoirs.

Materials and Methods

All chemical reagents including n-hexane, ethanol, cyclohexane, ethyl acetate, n-butanol, dodecane, chlorohexadecane, NaOH, NaHSO4, Na2SO4, and H2SO4 were of analytical grade and purchased from Shanghai Lingfeng Chemical Reagent Co. Ltd. (Shanghai, China).

Site descriptions and sample collection. Production fluid samples were collected from crude-oil-producing wells in Jiangsu, Xinjiang and Huabei Oilfields, of China. The temperature was 80~90 °C (Jiangsu Oilfield, sample J1 to J6), 37~45 °C (Huabei Oilfield, sample H1 to H4) and 21 and 32 °C (Xinjiang Oilfield, sample X1 and X2). The three oil fields have coverage of low-, mesophilic-, and high-temperature reservoirs. Other physicochemical information of each oilfield is given in Table 1. The alkane composition of the samples Hs, Js, and Xs are also provided in Figs. S2.1-S2.12. The samples were collected into sterile 5 L bottles to completely full and then capped after flushing the lines for about 30 min.

Extraction of organic acids. Extraction of long-chain fatty acids. Production fluid (approximately 0.5 L) from each sample was transferred into a 1 L round-bottom flask containing 2.5 g of NaOH. The contents were thoroughly mixed with a mechanical stirrer for 2 h. Then, aqueous phase was collected and extracted three times with 30 mL of n-hexane. The oil phase was mixed with 100 mL of 1% NaOH in 50% ethanol-water solution for 2 h, and the process was repeated once. The aqueous phase was further treated as described above and the extracts were combined. The solution was filtered, concentrated and acidified with HCl to pH < 2 at 0 °C. Ten mL of ethyl acetate were added and, organic acids extracted three times. The solvent was removed by rotary evaporation at 45 °C and the residues were dried by passing through Na2SO4.

Extraction of volatile fatty acids. To extract volatile fatty acids, ammonia was added to 50 mL of production fluid in a tube until pH > 10. The tube was heated at 105 °C in an oven until completely removal of water. The residues were sealed tightly before further use.

Derivatization of long chain fatty acids. The extracted long-chain fatty acids were derivatized via ethyl esterification. In a 100 mL round-bottom flask, a solution of ethanol-cyclohexane (10 mL, 1: 1), and 0.2 g of NaHSO4 was added into the organic acids extracts. The flask equipped with a water separator was then transferred to an oil-bath, and refluxed at 80 °C till no more water was produced. After cooling to room temperature, ethanol and cyclohexane were removed, and deionized water was added. Esters were extracted three times with 10 mL of ethyl acetate, and then combined. The ethyl acetate was removed after drying over anhydrous Na2SO4.

Derivatization of volatile fatty acids. Volatile fatty acids were analyzed after derivatization via n-butyl esterification as previously described in Ref. 56.

Gas chromatography-Mass spectrometry (GC-MS). All GC-MS analyses were performed on an Agilent 6890 GC equipped with an HP-5MS capillary column (30 m x 0.25 mm x 0.25 |im) and a mass detector (MSD 5975). For analyses of long-chain fatty acids, the injection port temperature was held at 280 °C. The oven temperature was initially held at 60 °C for 2 min, then increased at a rate of 10 °C per min to 260 °C and held at this final temperature for 30 min. The MS detector acquired the data in the scan mode, from 30 to 1000 mass units. For volatile fatty acids, the injection port temperature was 250 °C. The oven temperature was initially held at 60 °C for 1 min, and then increased at 15 °C per min to 145 °C. The MS detector acquired data in the scan mode, from 30 to 210 mass units. EI was operated at 70 eV and the ion source temperature was held at 230 °C for both long-chain and volatile fatty acids. Analytical reproducibility for replicate analyses (n = 3) of the alkylsuccinate of parent alkane C1-C8 in the production fluids was 0.12% relative standard deviation.

Identification of alkylsuccinates/2-(1-methylalkyl)succinates. GC-MS was used for the characterization and identification of possible degradation intermediates. GC-MS is well suited for coping with high sample numbers in reasonable measurement times with respect to both technical accuracy and identification and quantification of low-molecular-weight metabolites57.

The identification of diethyl alkylsuccinate was performed as established previously by Bian et al44. The study revealed that diethyl alkylsuccinates have four EI mass spectrum characteristics at m/z 128, 174, M + -45 and M+-87 (Mass spectra of identified alkylsuccinates are shown in Supplementary Materials as

Figs. S1.1-S1.8). Diethyl alkylsuccinates were identified by scanning the above four characteristic ions in the total ion chromatogram and comparison of the retention times with those of standard compounds44.

DNA extraction. Approximately 600 ml of each production fluid were filtered onto membrane filters (0.2-| m-pore-size, 50 mm diameter, Shanghai, China). Genomic DNAs were extracted from the filters using an E.Z.N.A.™ Soil DNA kit (D5625-01, Omega Bio-Tek, Inc., USA), according to the manufacturer's protocol.

Amplification of alkylsuccinate/2-(1-methylalkyl)succinate synthase alpha-subunit (assA/ masD) gene fragments. Portions of gene encoding the alpha subunit of the alkylsuccinate synthase were amplified with three primers sets, viz. assA2F (5'-YATGWACTGGCACGGMCA-3')/ assA2R(5'-GCRTTTTCMACCCAKGTA-3')18, 7757f-1 (5'-TCGGACGCGTGCAACGATCTGA-3')/ 8543R (5'-TCGTCRTTGCCCCAYTTNGG-3'), and 7766f (5'-TGTAACGGCATGACCATTGCGCT-3')/ 8543R (5'-TCGTCRTTGCCCCAYTTNGG-3')41 were used for the amplification in this study. The thermal cycler program for primers assA2F/assA2R was performed as described by Aitken et al18, and primer sets 7757f-1/8543R and 7766f/8543R followed the condition described by von Netzer et al41. Unless otherwise mentioned, all PCR products obtained above were first visualized by agarose gel (1%, w/v) electrophoresis followed by gel staining (DuRed nucleic acid gel stain, Beijing, China) to ensure the correct size fragment was amplified. Subsequently, PCR products resulting from independent five (5) reactions were pooled and visualized by agarose gel (1.8%, w/v) electrophoresis (50 min at 160 V). The appropriately sized fragments were excised and purified with a DNA purification kit (Axygen" Biosciences, Inc., CA, USA) prior to cloning.

Construction of assA/masD genes clone libraries, sequencing and phylogenetic analyses. Purified assA/masD gene-PCR fragments were directly cloned into Escherichia coli DH5a using a pMD19*-T Simple cloning vector (Takara", Japan) following the instructions of the manufacturer. Recombinant cells were spread onto LB agar plates containing ampicillin, IPTG and X-Gal. White clones were randomly selected and cultured overnight at 37 °C in 0.8 ml of Luria Broth (LB) medium in the presence of ampicillin. The clones were screened for the presence of correct insert by PCR using the forward M13F (-47) (5'-CGCCAGGGTTTTCCCAGTCACGAC-3') and the reverse RV-M (5'-GAGCGGATAACAATTTCACACAGG-3') plasmid specific primers, followed by agarose gel elec-trophoresis with subsequent DuRed staining. Sequencing was performed on an ABI 3730 sequencer (Dye-Terminator Cycle Sequencing; Applied Biosystems). The obtained assA/masD gene sequences were first trimmed to remove vector sequences and then compared to GenBank Database using the BLASTX algorithm to identify nearest related ones. assA/masD gene sequences were clustered into OTUs and representative OTUs from clones libraries as well as reference sequences from GenBank were translated and aligned using Clustal Omega58. Phylogenetic tree was constructed based on the Neighbor-Joining method59 and the Poisson correction method using the MEGA6 software60. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) is shown next to the branches. assA/masD gene fragments obtained in this study were deposited in the GenBank database under accession numbers KM229422-KM229513 and KM251716-KM251802.


1. Labinger, J. A. & Bercaw, J. E. Understanding and exploiting C-H bond activation. Nature 417, 507-514 (2002).

2. Mbadinga, S. M. et al. Microbial communities involved in anaerobic degradation of alkanes. Int. Biodeter. Biodegr. 65, 1-13 (2011).

3. Heider, J. & Schuhle, K. [Anaerobic biodegradation of hydrocarbons including methane] The Prokaryotes [Rosenberg E.(ed.)] [1028-1049] (Springer, Berlin Heidelberg, 2013).

4. Singh, S., Kumari, B. & Mishra, S. [Microbial degradation of alkanes] Microbial Degradation of Xenobiotics [Singh S. N. (ed)] [439-469] (Springer, Berlin Heidelberg, 2012).

5. Widdel, F., Knittel, K. & Galushko, A. [Anaerobic hydrocarbon-degrading microorganisms: an overview] Handbook, of hydrocarbon and lipid microbiology [Timmis K. N. (ed.)] [1997-2021] (Springer, Berlin Heidelberg, 2010).

6. Grossi, V., Cravo-Laureau, C., Guyoneaud, R., Ranchou-Peyruse, A. & Hirschler-Rea, A. Metabolism of n-alkanes and n-alkenes by anaerobic bacteria: A summary. Org. Geochem. 39, 1197-1203 (2008).

7. Das, N. & Chandran, P. Microbial degradation of petroleum hydrocarbon contaminants: an overview. Biotech. Res. Int. 2011, Article ID 941810, doi:10.4061/2011/941810 (2010).

8. Head, I. M., Jones, D. M. & Larter, S. R. Biological activity in the deep subsurface and the origin of heavy oil. Nature 426, 344-352 (2003).

9. Aitken, C. M., Jones, D. M. & Larter, S. R. Anaerobic hydrocarbon biodegradation in deep subsurface oil reservoirs. Nature 431, 291-294 (2004).

10. Jones, D. M. et al. Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451, 176-180 (2007).

11. Magot, M., Ollivier, B. & Patel, B. K. Microbiology of petroleum reservoirs. A. Van. Leeuw. 77, 103-116 (2000).

12. Mbadinga, S. M. et al. Analysis of alkane-dependent methanogenic community derived from production water of a high-temperature petroleum reservoir. Appl. Microbiol. Biotechnol. 96, 531-542 (2012).

13. Li, W. et al. Microbial community characteristics of petroleum reservoir production water amended with n-alkanes and incubated under nitrate-, sulfate-reducing and methanogenic conditions. Int. Biodeter. Biodegr. 69, 87-96 (2012).

14. Zhou, L. et al. 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, 1680-1691 (2012).

15. Berdugo-Clavijo, C. & Gieg, L. Conversion of crude oil to methane by a microbial consortium enriched from oil reservoir production waters. Front. Microbiol. 5, 197, doi:10.3389/fmicb.2014.00197 (2014).

16. Hasegawa, R., Toyama, K., Miyanaga, K. & Tanji, Y. Identification of crude-oil components and microorganisms that cause souring under anaerobic conditions. Appl. Microbiol. Biotechnol. 98, 1853-1861 (2014).

17. Gieg, L. M., Davidova, I. A., Duncan, K. E. & Suflita, J. M. Methanogenesis, sulfate reduction and crude oil biodegradation in hot Alaskan oilfields. Environ. Microbiol. 12, 3074-3086 (2010).

18. Aitken, C. M. et al. Evidence that crude oil alkane activation proceeds by different mechanisms under sulfate-reducing and methanogenic conditions. Geochim. Cosmochim. Ac. 109, 162-174 (2013).

19. Agrawal, A. & Gieg, L. M. In situ detection of anaerobic alkane metabolites in subsurface environments. Front. Microbiol. 4, 140, doi:10.3389/fmicb.2013.00140 (2013).

20. Callaghan, A. V. Enzymes involved in the anaerobic oxidation of n-alkanes: from methane to long-chain paraffins. Front. Microbiol. 4, 89, doi:10.3389/fmicb.2013.00089 (2013).

21. Wilkes, H. et al. 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, 235-243 (2002).

22. Grundmann, O. et al. Genes encoding the candidate enzyme for anaerobic activation of n-alkanes in the denitrifying bacterium, strain HxN1. Environ. Microbiol. 10, 376-385 (2008).

23. Callaghan, A. V., Wawrik, B., Ni Chadhain, S. M., Young, L. Y. & Zylstra, G. J. Anaerobic alkane-degrading strain AK-01 contains two alkylsuccinate synthase genes. Biochem. Bioph. Res. Co. 366, 142-148 (2008).

24. Callaghan, A. V., Gieg, L. M., Kropp, K. G., Suflita, J. M. & Young, L. Y. Comparison of mechanisms of alkane metabolism under sulfate-reducing conditions among two bacterial isolates and a bacterial consortium. Appl. Environ. Microb. 72, 4274-4282 (2006).

25. Rabus, R. et al. 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, 1707-1715 (2001).

26. Wilkes, H. et al. Formation of n-alkane- and cycloalkane-derived organic acids during anaerobic growth of a denitrifying bacterium with crude oil. Org. Geochem. 34, 1313-1323 (2003).

27. Cravo-Laureau, C., Grossi, V., Raphel, D., Matheron, R. & Hirschler-Rea, A. Anaerobic n-alkane metabolism by a sulfate-reducing bacterium, Desulfatibacillum aliphaticivorans strain CV2803T. Appl. Environ. Microb. 71, 3458 (2005).

28. Davidova, I. A., Gieg, L. M., Nanny, M., Kropp, K. G. & Suflita, J. M. Stable isotopic studies of n-alkane metabolism by a sulfate-reducing bacterial enrichment culture. Appl. Environ. Microb. 71, 8174-8182 (2005).

29. Musat, F., Wilkes, H., Behrends, A., Woebken, D. & Widdel, F. Microbial nitrate-dependent cyclohexane degradation coupled with anaerobic ammonium oxidation. ISME J. 4, 1290-1301 (2010).

30. Kropp, K. G., Davidova, I. A. & Suflita, J. M. Anaerobic oxidation of n-dodecane by an addition reaction in a sulfate-reducing bacterial enrichment culture. Appl. Environ. Microb. 66, 5393-5398 (2000).

31. Kniemeyer, O. et al. Anaerobic oxidation of short-chain hydrocarbons by marine sulphate-reducing bacteria. Nature 449, 898-901 (2007).

32. Rios-Hernandez, L. A., Gieg, L. M. & Suflita, J. M. Biodegradation of an alicyclic hydrocarbon by a sulfate-reducing enrichment from a gas condensate-contaminated aquifer. Appl. Environ. Microb. 69, 434-443 (2003).

33. Widdel, F. & Rabus, R. Anaerobic biodegradation of saturated and aromatic hydrocarbons Curr. Opin. Biotech. 12, 259-276 (2001).

34. Meckenstock, R. U., Safinowski, M. & Griebler, C. Anaerobic degradation of polycyclic aromatic hydrocarbons. FEMS Microbiol. Ecol. 49, 27-36 (2004).

35. Biegert, T., Fuchs, G. & Heider, J. Evidence that anaerobic oxidation of toluene in the denitrifying bacterium thauera aromatica is initiated by formation of benzylsuccinate from toluene and fumarate. Eur. J. Biochem. 238, 661-668 (1996).

36. Gieg, L. M. et al. Assessing in situ rates of anaerobic hydrocarbon bioremediation. Microb. Biotech. 2, 222-233 (2009).

37. Callaghan, A. V. Metabolomic investigations of anaerobic hydrocarbon-impacted environments. Curr. Opin. Biotech. 24, 506-515 (2013).

38. Gieg, L. M. & Suflita, J. M. Detection of anaerobic metabolites of saturated and aromatic hydrocarbons in petroleum-contaminated aquifers. Environ. Sci. Technol. 36, 3755-3762 (2002).

39. Parisi, V. A. et al. Field metabolomics and laboratory assessments of anaerobic intrinsic bioremediation of hydrocarbons at a petroleum-contaminated site. Microb. Biotech. 2, 202-212 (2009).

40. Callaghan, A. V. et al. Diversity of benzyl-and alkylsuccinate synthase genes in hydrocarbon-impacted environments and enrichment cultures. Environ. Sci. Technol. 44, 7287-7294 (2010).

41. von Netzer, F. et al. Enhanced gene detection assays for fumarate-adding enzymes allow uncovering of anaerobic hydrocarbon degraders in terrestrial and marine systems. Appl. Environ. Microb. 79, 543-552 (2013).

42. Kimes, N. E. et al. Metagenomic analysis and metabolite profiling of deep-sea sediments from the Gulf of Mexico following the Deepwater Horizon oil spill. Front. Microbiol. 4, 50, doi:10.3389/fmicb.2013.00050 (2013).

43. Duncan, K. E. et al. Biocorrosive thermophilic microbial communities in Alaskan North Slope oil facilities. Environ. Sci. Technol. 43, 7977-7984 (2009).

44. Bian, X.-Y. et al. Synthesis of anaerobic degradation biomarkers alkyl-, aryl-and cycloalkylsuccinic acids and their mass spectral characteristics. Eur. J. Mass Spectrom 20, 287-297 (2014).

45. Tierney, M. & Young, L. [Anaerobic degradation of aromatic hydrocarbons] Handbook of Hydrocarbon and Lipid Microbiology [Timmis K. N. (ed.)] [925-934] (Springer, Berlin Heidelberg, 2010).

46. Embree, M., Nagarajan, H., Movahedi, N., Chitsaz, H. & Zengler, K. Single-cell genome and metatranscriptome sequencing reveal metabolic interactions of an alkane-degrading methanogenic community. ISME J. 8, 757-767 (2014).

47. Tan, B., Nesbo, C. & Foght, J. Re-analysis of omics data indicates Smithella may degrade alkanes by addition to fumarate under methanogenic conditions. ISME J. 8, 2353-2356 (2014).

48. Aktas, D. F. et al. Effects of oxygen on biodegradation of fuels in a corroding environment. Int. Biodeter. Biodegr. 81, 114-126 (2013).

49. Kleindienst, S. et al. Diverse sulfate-reducing bacteria of the Desulfosarcina/Desulfococcus clade are the key alkane degraders at marine seeps. ISME J. 8, 2029-2044 (2014).

50. Gray, N. D., Sherry, A., Hubert, C., Dolfing, J. & Head, I. M. Methanogenic degradation of petroleum hydrocarbons in subsurface environments: remediation, heavy oil formation, and energy recovery. Adv. Appl. Microbiol. 72, 137-161 (2010).

51. Jones, D. M. et al. Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451, 176-180 (2008).

52. Sherry, A. et al. Volatile hydrocarbons inhibit methanogenic crude oil degradation. Front. Microbiol. 5, 131, doi: 10.3389/ fmicb.2014.00131 (2014).

53. Boll, M., Fuchs, G. & Heider, J. Anaerobic oxidation of aromatic compounds and hydrocarbons. Curr. Opin. Chem. Biol. 6, 604-611 (2002).

54. Gray, N. et al. The quantitative significance of Syntrophaceae and syntrophic partnerships in methanogenic degradation of crude oil alkanes. Environ. Microbiol. 13, 2957-2975 (2011).

55. Liu, Y., Balkwill, D. L., Aldrich, H. C., Drake, G. R. & Boone, D. R. Characterization of the anaerobic propionate-degrading syntrophs Smithella propionica gen. nov., sp. nov. and Syntrophobacter wolinii. Int. J. Syst. Bacteriol. 49, 545-556 (1999).

56. Wang, L. Y. et al. Methanogenic microbial community composition of oily sludge and its enrichment amended with alkanes incubated for over 500 days. Geomicrobiol. J. 29, 716-726 (2012).

57. Weckwerth, W. Metabolomics in systems biology. Annu. Rev Plant. Biol. 54, 669-689 (2003).

58. Sievers, F. et al. Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol. Syst. Biol. 7, 539-544 (2011).

59. Saitou, N. & Nei, M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol. Biol. Evol. 4, 406-425 (1987).

60. Tamura, K., Stecher, G., Peterson, D., Filipski, A. & Kumar, S. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol. Biol. Evol. 30, 2725-2729 (2013).


This work was supported by the National Natural Science Foundation of China (No. 51174092, 41373070) and the NSFC/RGC Joint Research Fund (No. 41161160560).

Author Contributions

This study was designed by X.-Y.B., S.M.M., S.-Z.Y. and B.-Z.M. X.-Y.B. performed the analyses of metabolites in production fluid samples. X.-Y.B., S.M.M. and Y.-F.L prepared genomic DNAs, amplification and cloning exercises for assA/masD gene assessment. Phylogenetic analyses were performed by S.M.M. assisted by X.-Y.B. and Y.-F.L. X.-Y.B. and S.M.M. wrote the manuscript, assisted by all co-authors. J.-F.L. provided oil reservoirs data. R.-Q.Y. and J.-D.G. provided valuable suggestions in the design of the experiments and the preparation of the manuscript. All authors reviewed the final manuscript.

Additional Information

Supplementary information accompanies this paper at

Competing financial interests: The authors declare no competing financial interests.

How to cite this article: Bian, X.-Y. et al. Insights into the Anaerobic Biodegradation Pathway of n-Alkanes in Oil Reservoirs by Detection of Signature Metabolites. Sci. Rep. 5, 9801; doi: 10.1038/ srep09801 (2015).

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