Evidence for in situ methanogenic oil degradation in the Dagang oil field Núria Jiménez a,⇑ , Brandon E.L. Morris a,1 , Minmin Cai a,b , Friederike Gründger c , Jun Yao b , Hans H. Richnow a , Martin Krüger c a Department of Isotope Biogeochemistry, Helmholtz Centre for Environmental Research – UFZ, Permoserstraße 15, 04318 Leipzig, Germany b School of Civil & Environment Engineering, University of Science and Technology Beijing, 30 Xueyuan Road, Haidian District, Beijing, PR China c Department of Resources Geochemistry, Federal Institute for Geosciences and Natural Resources – BGR, Stilleweg 2, 30655 Hannover, Germany article info Article history: Received 23 April 2012 Received in revised form 19 July 2012 Accepted 26 August 2012 Available online 1 September 2012 abstract In situ biotransformation of oil to methane was investigated in a reservoir in Dagang, China using chem- ical fingerprinting, isotopic analyzes and molecular and biological methods. The reservoir is highly meth- anogenic despite chemical indications of advanced oil degradation, such as depletion of n-alkanes, alkylbenzenes and light polycyclic aromatic hydrocarbon (PAH) fractions or changes in the distribution of several alkylated polycyclic aromatic hydrocarbons. The degree of degradation strongly varied between different parts of the reservoir, ranging from severely degraded to nearly undegraded oil com- positions. Geochemical data from oil, water and gas samples taken from the reservoir are consistent with in situ biogenic methane production linked to aliphatic and aromatic hydrocarbon degradation. Micro- cosms were inoculated with production and injection waters in order to characterize these processes in vitro. Subsequent degradation experiments revealed that autochthonous microbiota are capable of producing methane from 13 C labelled n-hexadecane or 2-methylnaphthalene and suggest that further methanogenesis may occur from the aromatic and polyaromatic fractions of Dagang reservoir fluids. The microbial communities from produced oil–water samples were composed of high numbers of micro- organisms (on the order to 10 7 ), including methane producing Archaea within the same order of magni- tude. In summary, the investigated sections of the Dagang reservoir may have significant potential for testing the viability of in situ conversion of oil to methane as an enhanced recovery method and biodeg- radation of the aromatic fractions of the oil may be an important methane source. Ó 2012 Elsevier Ltd. All rights reserved. 1. Introduction Microbial activity in oil reservoirs affects the quality and eco- nomic value of recovered petroleum products, not only by decreas- ing the n-alkane content of the oil, but also by increasing oil density, sulfur content, acidity and viscosity (Connan, 1984; Röling et al., 2003). Unlike shallow subsurface reservoirs, deeper petro- leum reservoirs are not commonly connected to meteoric water cycles, resulting in low nitrate and oxygen availability. Conse- quently, oil degradation by aerobic or facultative anaerobic organ- isms is limited. In addition, the supply of large amounts of Fe(III) or manganese(IV) via the water cycle is also unlikely due to poor sol- ubility and slow water recharge rates in subterranean cycles. Moreover, although iron and manganese oxides from sandstone could be used as electron acceptors for oil oxidation, they are un- likely to be responsible for significant compositional changes in the oil, due to their limited availability in the reservoir. However, oil degradation linked to sulfate reduction is possible when sulfate arises from geological sources, such as evaporitic sediments and limestone, or from the injection of seawater for pressure stabiliza- tion and may lead to significant oil degradation and increased residual-oil sulfur content. Methanogenic oil degradation, on the other hand, does not re- quire external electron acceptors and leads to less overall souring of the oil reservoir. Several studies have described methanogenic degradation of crude oil related compounds in vitro (Gieg et al., 2008; Jones et al., 2008) including n-alkanes (Zengler et al., 1999; Jones et al., 2008) and mono- and polyaromatic hydrocarbons (for a review see Gray et al., 2010). To sustain the process in an oil bearing reservoir, the only requirements would be adequate amounts of N and P for biomass production, trace metals and vita- mins for enzymes and a sufficient water supply delivered over geo- logical time scales for biodegradation to occur. This water and nutrient supply also may be provided during secondary production involving waterflood. Methane may be recovered relatively easily using extant production infrastructure and used as a downstream energy source. Therefore, the transformation of residual oil to methane is being considered as a tertiary recovery method for abandoned reservoirs with high water cuts and low oil recovery. 0146-6380/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.orggeochem.2012.08.009 ⇑ Corresponding author. Tel.: +49 341 235 1418; fax: +49 341 235 1443. E-mail address: [email protected](N. Jiménez). 1 Present address: University of Freiburg, Institute for Biology II – Microbiology, Schänzlestraße 1, 79104 Freiburg, Germany. Organic Geochemistry 52 (2012) 44–54 Contents lists available at SciVerse ScienceDirect Organic Geochemistry journal homepage: www.elsevier.com/locate/orggeochem
Transcript
Organic Geochemistry 52 (2012) 44–54
Contents lists available at SciVerse ScienceDirect
Evidence for in situ methanogenic oil degradation in the Dagang oil field
Núria Jiménez a,⇑, Brandon E.L. Morris a,1, Minmin Cai a,b, Friederike Gründger c, Jun Yao b,Hans H. Richnow a, Martin Krüger c
a Department of Isotope Biogeochemistry, Helmholtz Centre for Environmental Research – UFZ, Permoserstraße 15, 04318 Leipzig, Germanyb School of Civil & Environment Engineering, University of Science and Technology Beijing, 30 Xueyuan Road, Haidian District, Beijing, PR Chinac Department of Resources Geochemistry, Federal Institute for Geosciences and Natural Resources – BGR, Stilleweg 2, 30655 Hannover, Germany
a r t i c l e i n f o a b s t r a c t
Article history:Received 23 April 2012Received in revised form 19 July 2012Accepted 26 August 2012Available online 1 September 2012
0146-6380/$ - see front matter � 2012 Elsevier Ltd. Ahttp://dx.doi.org/10.1016/j.orggeochem.2012.08.009
1 Present address: University of Freiburg, InstituteSchänzlestraße 1, 79104 Freiburg, Germany.
In situ biotransformation of oil to methane was investigated in a reservoir in Dagang, China using chem-ical fingerprinting, isotopic analyzes and molecular and biological methods. The reservoir is highly meth-anogenic despite chemical indications of advanced oil degradation, such as depletion of n-alkanes,alkylbenzenes and light polycyclic aromatic hydrocarbon (PAH) fractions or changes in the distributionof several alkylated polycyclic aromatic hydrocarbons. The degree of degradation strongly variedbetween different parts of the reservoir, ranging from severely degraded to nearly undegraded oil com-positions. Geochemical data from oil, water and gas samples taken from the reservoir are consistent within situ biogenic methane production linked to aliphatic and aromatic hydrocarbon degradation. Micro-cosms were inoculated with production and injection waters in order to characterize these processesin vitro. Subsequent degradation experiments revealed that autochthonous microbiota are capable ofproducing methane from 13C labelled n-hexadecane or 2-methylnaphthalene and suggest that furthermethanogenesis may occur from the aromatic and polyaromatic fractions of Dagang reservoir fluids.The microbial communities from produced oil–water samples were composed of high numbers of micro-organisms (on the order to 107), including methane producing Archaea within the same order of magni-tude. In summary, the investigated sections of the Dagang reservoir may have significant potential fortesting the viability of in situ conversion of oil to methane as an enhanced recovery method and biodeg-radation of the aromatic fractions of the oil may be an important methane source.
� 2012 Elsevier Ltd. All rights reserved.
1. Introduction
Microbial activity in oil reservoirs affects the quality and eco-nomic value of recovered petroleum products, not only by decreas-ing the n-alkane content of the oil, but also by increasing oildensity, sulfur content, acidity and viscosity (Connan, 1984; Rölinget al., 2003). Unlike shallow subsurface reservoirs, deeper petro-leum reservoirs are not commonly connected to meteoric watercycles, resulting in low nitrate and oxygen availability. Conse-quently, oil degradation by aerobic or facultative anaerobic organ-isms is limited. In addition, the supply of large amounts of Fe(III) ormanganese(IV) via the water cycle is also unlikely due to poor sol-ubility and slow water recharge rates in subterranean cycles.Moreover, although iron and manganese oxides from sandstonecould be used as electron acceptors for oil oxidation, they are un-likely to be responsible for significant compositional changes inthe oil, due to their limited availability in the reservoir. However,
ll rights reserved.
x: +49 341 235 1443.Jiménez).for Biology II – Microbiology,
oil degradation linked to sulfate reduction is possible when sulfatearises from geological sources, such as evaporitic sediments andlimestone, or from the injection of seawater for pressure stabiliza-tion and may lead to significant oil degradation and increasedresidual-oil sulfur content.
Methanogenic oil degradation, on the other hand, does not re-quire external electron acceptors and leads to less overall souringof the oil reservoir. Several studies have described methanogenicdegradation of crude oil related compounds in vitro (Gieg et al.,2008; Jones et al., 2008) including n-alkanes (Zengler et al., 1999;Jones et al., 2008) and mono- and polyaromatic hydrocarbons(for a review see Gray et al., 2010). To sustain the process in anoil bearing reservoir, the only requirements would be adequateamounts of N and P for biomass production, trace metals and vita-mins for enzymes and a sufficient water supply delivered over geo-logical time scales for biodegradation to occur. This water andnutrient supply also may be provided during secondary productioninvolving waterflood. Methane may be recovered relatively easilyusing extant production infrastructure and used as a downstreamenergy source. Therefore, the transformation of residual oil tomethane is being considered as a tertiary recovery method forabandoned reservoirs with high water cuts and low oil recovery.
N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54 45
According to estimates more than 50% of the initial oil is unrecov-erable by conventional technology (Youssef et al., 2009) and willremain entrenched in the reservoir matrix. Therefore, methano-genesis could be an interesting strategy for microbially enhancedrecovery of carbon from exploited reservoirs.
Previous experiments performed in the area of microbial en-hanced oil recovery have been intent on increasing oil degradationby stimulating in situ production of biosurfactants after addition ofan oxidant (O2 or H2O2; e.g., Nazina et al., 2008). However, theseexperiments resulted in a slight enhancement in oil recovery uponformation of surfactants by aerobic, thermophilic microbiota, orbiomass that helped to indirectly solubilize the oil. However, thisstrategy requires constant addition of electron acceptors to sustainthe metabolic activity of aerobic microbial communities and maybe limited over longer periods of time. Consequently, recent atten-tion has focused on the potential of anaerobic microbial processesfor microbially enhanced oil recovery (MEOR) applications.
In light of these situations, the aim of this study was to assess theability of indigenous microbial communities from a water-floodedthermophilic oil reservoir to produce methane under reservoir con-ditions using laboratory microcosms and molecular biological ana-lyzes. As an incremental step towards this goal, we have conducteda geochemical study employing isotopic analyzes of reservoir fluidsto characterize microbial methanogenesis. In addition, the oil wasanalyzed using GC–MS fingerprinting techniques to assess the ef-fects of biodegradation on the distribution of oil constituents.
2. Materials and methods/experimental details
2.1. Site description
The Dagang oil field complex is located in the Huanghua depres-sion of the Bohai Bay Basin. It comprises a total area of 24 km2 inCang County, roughly 200 km SE of Beijing and consists of seriesof sandstone oil-bearing strata of the Paleogene and Neogene (Liand Wu, 1991; Vincent et al., 2009). The sampling campaign forthis study included four trips in 2010 and 2011 to the Hebei Prov-ince in NE China. The sampled oil complex contains different pro-duction blocks ranging in age from approximately 3 to more than40 years and well depths from 800 m to 2600 m. Production meth-ods for sampled wells range from primary production to tertiaryenhanced recovery methods (i.e. polymer flooding) (Nazina et al.,2007b). The average reservoir temperature ranged from 35 to80 �C at the time of sampling. Within the oil field complex, severalproduction wells are serviced by one injection well during second-ary recovery/waterflood and range in distances from 50–300 m tothe injection well. Hydraulic residence times for injected watersare typically on the order of 40–80 days (Nazina et al., 2007b). Adescription of the sampled blocks and wells is in Table 1. Three dif-ferent production blocks were sampled during the campaigns andare labelled as Block I, II and III.
2.2. Sampling and establishment of microcosms
Samples were collected directly from the wellhead of produc-tion and injection wells (Table 1) in sterile glass bottles, flushedwith formation gases and transported back to the laboratory inLeipzig, Germany. All samples were cooled to around 4 �C until fur-ther analysis. Anaerobic incubations were established in an anaer-obic chamber, using oil field fluids as inoculum. Glass serumbottles containing 10–30 ml of a sulfate free mineral medium(Widdel and Bak, 1992) were inoculated with 5–10 ml of formationwater and sealed with butyl rubber stoppers and aluminium crimpseals. Controls included autoclaved fluids to assess residual degas-sing of methane from the fluids, as well as replicates without any
added hydrocarbon substrates to determine potential methano-genesis from organic matter present in the fluids. Sulfate wasadded from a sterile anoxic stock solution to several replicates toa final concentration of 2 mM sulfate. The salinity of the microbialmedium was adjusted to mimic the conditions in Dagang (Appen-dix A). Sample headspace was flushed with nitrogen to removeresidual hydrogen (from the anaerobic chamber) and replicateswere incubated statically at 30 �C and 60 �C. The enrichments wereamended individually with non-labelled components to comparewith the labelled ones, [13C16] hexadecane, ethylbenzene, [13C7]toluene, 2-[13C]-methylnaphthalene, or 2-carboxynaphthalene asa sole carbon source. The labelled hexadecane was synthesizedfrom uniformly 13C labelled palmitic acid, (Campro Scientific, Ber-lin, Germany) by reduction of the carboxyl group to an alcohol(with LiAlH4), conversion to the p-tosylate ester and reduction tothe hydrocarbon (with LiAlH4). 2-[13C]-methylnaphthalene wassynthesized by the Institute of Organic Chemistry at the Universityof Leipzig, using a two step methylation of naphthalene: acetyla-tion to 2-[13C]-naphthol (analogous to Coombs et al., 2000) andreduction with Pd/C/H2 (analogous to Ofosu-Asante and Stock,1987). Purity was confirmed via gas chromatography mass spec-trometry (GC–MS).
Methane and CO2 production rates were calculated by perform-ing a linear regression of the methane increase with incubationtime and the values were expressed in lmol CH4 or CO2/day/mlsample, as described previously in Krüger et al. (2001).
2.3. Analytical methods (isotopes and headspace gases)
Methane production was analyzed by measuring microcosmheadspace isocratically at 60 �C using a GC-FID equipped with a 60
Hayesep D column (SRI 8610C, SRI Instruments, USA). Carbon diox-ide concentrations were determined using a methanizer-equippedFID, after reduction of the CO2 to methane. The stable isotopic com-position of methane and CO2 was measured using a gas chromatog-raphy–combustion–isotope ratio monitoring mass spectrometrysystem (GC–C–IRM–MS). The system consisted of a gas chromato-graph (6890 series; Agilent Technology), fitted with a CP-poraBOND Q column coupled to a combustion or high temperaturepyrolysis interface (GC-combustion III or GC/C-III/TC; Thermo Finn-igan, Bremen, Germany) and a MAT 252 IRMS for the carbon analy-sis or a MAT 253 IRMS for hydrogen analysis (both from ThermoFinnigan, Bremen, Germany). The carbon and hydrogen isotopiccompositions (R) are reported as delta notation (d13C and d2H) inparts per thousand (‰) relative to the Vienna Pee Dee Belemnite(VPDB) and Vienna Standard Mean Ocean Water (VSMOW), respec-tively (Richnow et al., 2003; Feisthauer et al., 2011).
Headspace samples were injected directly into the GC using asplit mode for the analysis of d13C and d2H for methane (for details,see Feisthauer et al., 2011). Water subsamples were used for theisotopic analysis of carbonates and deuterium in the H2O as out-lined below. For the carbonate analysis, an aliquot of each samplewas collected with a syringe, transferred to a crimped vial andacidified to pH < 2 using pure HCl. The gas phase was then injectedinto the GC–IRMS for isotope analysis. The error associated withthe system (accuracy and reproducibility) was around 0.5‰ and4‰, for carbon and hydrogen, respectively. The standard deviationof at least three independent measurements is reported. For theH2O analysis, an aliquot (5 ml) of each sample was cleaned withactivated carbon to remove any possible organic contaminationprior to the determination of isotope ratios.
2.4. Gas-chromatography mass-spectrometry of oil
Samples from the reservoir and from degradation experimentswere extracted with dichloromethane and dehydrated through a
5/18/2011 NA NA + NA +12 Prod 12/6/2010 1579–1616 �50 Yes �3 NA NA NA NA +
5/18/2011 NA NA + + +13 Prod 12/6/2010 1436–1443 �50 Yes From 2007 to 2012
(now closed)NA NA NA NA +
5/18/2011 + + + + +14 Prod 5/18/2011 1546–1791 30–40 Yes �2 + + + + +15 Prod 6/1/2010 NA �50 Yes NA NA + NA NA NA
a Prod: production well. Inj: injection well.b Lithology (and age) from Li and Wu, (1991).c Measured when sampling.d + Shows which analyzes have been performed for each sample. NA means not analyzed.
46N
.Jiménez
etal./O
rganicG
eochemistry
52(2012)
44–54
Table 2Diagnostic ratios for source and weathering assessment for the oil samples.
a The indexes %29ab, %32ab, %27Ts, D2/P2 and D3/P3 have been selected for source identification, whereas changes in the other indexes may indicate different degrees ofdegradation.
N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54 47
2 g anhydrous Na2SO4 column. The oil content in the sample wasdetermined by gravimetry in 1.0 ml of the eluate and carefullyevaporated until dryness. An aliquot (10 mg) of the eluate wascleaned by passing it through 2 g of Al2O3 (5% w/w deactivated),concentrated and exchanged to hexane (1.0 ml) by a gentle solventevaporation under a stream of nitrogen gas. GC–MS analysis wasperformed using a 7890A gas chromatograph (Agilent Technolo-gies), fitted with a capillary column (J&W Scientific, Folsom, CA,USA) HP-5 MS (30 m � 0.25 mm i.d., 0.25 lm film), coupled to a5975C MS spectrometer equipped with a triple axis detector (Agi-lent Technologies) as reported elsewhere (Jiménez et al., 2006).
2.5. Assessment of oil degradation
To assess the extent of microbiological oil degradation, the dis-tribution of n-alkanes or polycyclic aromatic hydrocarbons wascalculated with relation to 17a(H),21b(H)-hopane (m/z 191), usedas internal conservative molecular marker (Prince et al., 1994).Sample 30 was used as a reference to calculate the relative degrada-tion for the rest of the samples. The quantification of n-alkanes wasdetermined by using the m/z 85 fragment ion. Linear alkylcyclo-hexanes, alkylbenzenes and alkyltoluenes were quantified by mon-itoring their characteristic ions (m/z 82, 92 and 106, respectively).Quantification of individual aromatic compounds was based on themolecular ion for each: N–N4, naphthalenes (m/z 128, 142, 156,170, 184); F–F3, fluorenes (m/z 166, 180, 194, 208); P–P3, phenan-threnes (m/z 178, 192, 206, 220); D–D3, dibenzothiophenes (m/z184, 198, 212, 226); Py–Py3, fluoranthenes and pyrenes (m/z 202,216, 230, 244); C–C3, chrysenes (m/z 228, 242, 256, 270). In addi-tion, several molecular markers were used to calculate indexes forfingerprinting and weathering assessment: triaromatic steroids(m/z 231), steranes and diasteranes (m/z 217 and 218) and triter-panes (m/z 191). The different diagnostic ratios are specified inTable 2.
2.6. Molecular methods
For molecular characterization and cloning experiments, DNAfrom the microcosms was extracted after incubation with individ-ual hydrocarbons using a modified protocol from Lueders et al.(2004). For further purification of crude DNA, ethidium bromidewas added to 0.6 mg/ml and ammonium acetate to 2.6 M final con-centration (Lovell and Piceno, 1994). Genes of interest were quan-tified by real time PCR using an ABI Prism 7000 (Applied
Biosystems, Life Technologies Corporation, Carlsbad, California,USA). The 16S rRNA gene copy numbers for Archaea and Bacteriawere determined as described previously (Takai and Horikoshi,2000; Nadkarni et al., 2002). Methanogen abundance was assessedusing the methyl-CoM reductase gene A (mcrA) (Steinberg and Re-gan, 2009). Microorganisms capable of dissimilatory sulfate reduc-tion were quantified using dsrA gene copy number. This gene codesfor the alpha subunit of the dissimilatory (bi)sulfite reductase(Kondo et al., 2004; Schippers and Neretin, 2006). All PCR reactionswere run in triplicate at three dilutions. Copy numbers are ex-pressed as DNA copies/ml sample.
3. Results
3.1. Oil chemical analysis
The majority of the oil–water samples taken from the reservoirpresent an elevated content of heavy and polar oil fractions (resinsand asphaltenes, up to 29%), whereas the saturated fraction repre-sents 43 to less than 50% of the bulk oil. These values are slightlyhigher and lower, respectively, than values previously reportedby Nazina and coworkers (around 20% of resins and asphaltenesand >53% saturated hydrocarbons) and might suggest higher bio-degradation levels than previously described (Nazina et al.,2007b). In addition, most samples are completely depleted of n-al-kanes, as revealed by GC–MS analysis (Fig. 1). The degree of degra-dation compared to a relatively non-degraded sample from thesame reservoir was higher than 90% (considering C10–C35). Oil pro-filing revealed that alkylbenzenes and alkyltoluenes are severelydegraded. However, branched alkanes and alkylcyclohexanes arestill present to a varying extent in most of the samples.
Polycyclic aromatic hydrocarbon (PAH) degradation varied sig-nificantly among samples and within compound families, rangingfrom around 10% to nearly 100% for naphthalenes, fluorenes, dib-enzothiophenes and phenanthrenes (Table 3). Compound removaldecreased with increasing molecular weight, as generally ex-pected. Tetra-aromatic hydrocarbons (e.g., pyrenes and chrysenes)were slightly affected, as reflected by the relative decrease of C2-and C3-dibenzothiophenes with respect to C2- and C3-pyrenes(D2/C2 and D3/C3) (Fig. 2). This observation is consistent with hea-vy to severe biodegradation (4–7 using the scale of Peters andMoldowan, 1993) (Head et al., 2003).
For alkylated PAHs, we have detected changes in the isomericdistributions of tetramethylnaphthalenes (TMN), methylphenanth-
Fig. 1. (A) GC–MS total ion current (TIC) profiles of representative samples with different degrees of biodegradation. Each sample corresponds to a different sampling well.The non-degraded profile corresponds to sample Bannan 3 from block I. The numbers 10, 20 and 30 refer to n-alkane chain lengths. (B) Ion chromatograms of the C4-naphthalenes (m/z 184), C1-phenanthrenes (m/z 206) and C1-pyrenes and fluoranthenes (m/z 216) for the same samples. Each profile is presented on a scale relative to thelargest peak. The identification of the different isomers is given below.
48 N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54
Table 3Degradation percentages of different hydrocarbon families with respect to the non-biodegraded sample (Block I, sample 3). Samples have been ordered according to the extent ofbiodegradation.
3 7 11 12 8 10 5 13 1 14
C17/Pr 65 15 NA NA NA 15 NA NA NA NAC18/Ph 63 11 NA NA NA 8 NA NA NA NA
Fig. 2. Molecular marker indices for source and biodegradation assessment. Stars represent sample 3, Block I, which was used as a control. The other samples are representedas shaded circles, going from dark grey (less degraded) to white (more degraded).
N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54 49
renes (MP), methyldibenzothiophenes (MD) and higher molecularweight PAHs such as methylpyrenes (MPy). This observation sug-gests that biodegradation of these oil components in this reservoiroccurs via stereospecific biological processes. Specifically, 1,3,6,7-and 1,2,3,6-TMN are the most easily degraded isomers and, like1,2,6,7-TMN, their relative abundance decreases with increasingbiodegradation. On the other hand, 1,3,5,7- and 1,2,5,7-TMN seemto be more recalcitrant (Fig. 1); 3- and 1-methylphenanthrene ex-hibit higher levels of biodegradation compared to the other iso-mers, whereas the 4/9-MP relative abundance increased over awide range (Fig. 1). However, this pattern was not consistentamong the most degraded samples, as reported in other studies(Williams et al., 1986). The abundance of 2- and 3-MD decreasedin relation to 1-MD (Fig. 2). Persistence of 4-methylpyrene (4-MPy) was noted, while all other methylpyrenes were significantlydegraded (Fig. 1).
Aliphatic and aromatic biomarker distributions (hopanes, ster-anes and triaromatic steroid hydrocarbons) varied with the extentof degradation (Fig. 2). Steranes and triaromatic steroid hydrocar-bons followed a similar trend, whereby degradation decreasedwhen increasing side chain length, as reflected in the ratios of%27bb and %26TA. In addition, a preferential removal of severalC29 steranes was observed: The 14a(H),a(H) and 20R isomers weremore readily degraded compared to the 14b(H),b (H) and 20S iso-mers, respectively (%29aaS and %29bb(R + S)). On the other hand,diagnostic ratios, such as the relative abundance of C2- and C3-phenanthrenes compared to C2- and C3-dibenzothiophenes (D2/P2 and D3/P3) or the distribution of bishomohopane S and R iso-mers (%32abS), suggest that all oils from this study belong to the
same source. In addition, the lack of significant differences in theTm/Ts index (%27Ts) among the samples indicates that they havea similar thermal maturity and imply a similar genesis, with thelone exception of sample 14.
3.2. Gas analysis
Gas phases recovered from the reservoir consisted mainly ofCH4, while C2 and C3 were less abundant and other light n-alkanes(C4–C6) were rarely detected, as reflected by the ratio of C1/R(C1–C5) (Table 4). However, samples from Block I (wells 2 and 3) wereparticularly wet, with values below 0.6. The C1/(C2 + C3) ratios var-ied from 2 to nearly 600 and indicate either thermogenic origin,according to the Bernard diagram (Bernard et al., 1976; AppendixB) or a mixture of biodegraded and secondary microbial gas (Mil-kov, 2011). The ratio of CH4/CO2 ranged from 1 to around 200.Methanogenic alkane degradation produces methane to CO2 ratiosof about 3.27:1. However, our values are generally much higher,which could be related to the diversity of carbon sources, a furtherdegradation of CO2 or the solubility of CO2 in formation water orprecipitation of carbonates in the reservoir.
The isotopic composition of methane carbon (d13C) from differ-ent wells ranged from �39‰ to �68‰ relative to VPDB and aver-aged around �47‰ (Table 4). These values are consistent withbiogenic methane production (Pallaser, 2000; Larter et al., 2005)and similar to values from other degraded reservoirs in China(Dai et al., 2004). Two of the samples (1 and 14) have a lighter va-lue (below �55‰) and, according to Milkov (2011), are typical forprimary microbial gas. Carbon dioxide was highly enriched in 13C
Table 4C and H isotopic composition of the fluids (�) from the reservoir, together with their dryness. (�) relative to the VPDB and VSMOW standards.
Fig. 3. Hydrogen isotopic discrimination between methane and water (Dd2H) vs.carbon isotopic discrimination between methane and CO2 (Dd13C). Error barsrepresent the standard deviation of three independent analyzes.
50 N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54
for most of the samples, with values observed up to +17.2‰. Thelightest value corresponded to sample 14 from Block III(�12.8‰), followed by samples from Blocks I and II (up to�7.6‰). The bulk isotopic discrimination (Dd13C) between meth-ane and CO2 was between 32‰ and 65‰ (Fig. 3), in accordancewith previously reported results for methane formation duringhydrocarbon degradation (Feisthauer et al., 2010; Morris et al.,2012). They also are typical of values found in highly biodegradedsubsurface oils from marine systems (Larter and di Primio, 2005)and may indicate that extensive reduction of CO2 to methane hasoccurred (Jones et al., 2008; Milkov, 2011). However, no clear cor-relation can be established between the isotopic discrimination ofmethane/carbon dioxide and the degree of biodegradation at thistime.
The d2H values for methane and water ranged from �220‰ to�246‰ and �69‰ to �73‰ relative to the VSMOW, respectively.These values are on the lower range of previously reported statis-tics from marine sediments (Whiticar et al., 1986) (�250‰ to�170‰). The d2H discrimination between methane and water
(Dd2H(CH4–H2O)) varied between �145‰ and �170‰ and werefound to be significantly lower than values from enrichment cul-tures studied by Feisthauer et al. (2010) (Dd2H(CH4–H2O) �336‰
to �257‰) (Fig. 3).
3.3. Microbial abundance
Thirteen production wells and three injection wells were sam-pled intermittently over a period of 2 years throughout the Dagangfield complex. A thorough microbiological survey using qPCR wasconducted to characterize the bacterial and Archaeal numbers atthe site via amplification of the 16S rRNA gene. Further insight intobiogeochemical processes was achieved by determining mean copynumber of the dissimilatory sulfate reductase (dsrA) and methyl-CoM reductase (mcrA) genes. Reservoir temperatures in Dagangusually range from 35–60 �C and can reach 80 �C in the deeper sec-tion (Table 1). Average bacterial numbers for injection and produc-tion wells, based on 16S rRNA, ranged from 105 to 108 copies/mlsample. Archaeal numbers were between 103 and 107 copies/mlsample. Generally, bacterial and Archaeal copy numbers were onthe same order of magnitude for individual wells. The exceptionto this rule occurred when wells with high water cuts were sam-pled (above 90%, Well 13). In this case, bacteria outnumbered Ar-chaea by two orders of magnitude. Clone libraries of 16S rRNAgenes were constructed to assess the dominant Archaeal generawithin nine Dagang production/injection wells (Appendix C). Pro-duction water samples from wells 10 and 15 were dominated byMethanosarcina (100%) and Methanobacterium (47%) sequences,respectively. The dominant Archaea in well 14 were the crenar-chaeotal genus Hyperthermus (80%) and the euryarchaeotal genusArchaeoglobus (15%). Interestingly, no methanogenic genera weredetected in this sample. Methanosphaera, Methanosarcina and Met-hanobacterium were the dominant genera in samples from theother wells.
The dsrA numbers ranged from 104 to 107 copies/ml sample andseem to indicate that the Dagang field could quickly become sulfid-ogenic if sulfate-containing injection waters were added to the for-mation. In addition, if sulfide oxidation is promoted by adding ofO2 or H2O2 to the injection water, biogenic sulfate reduction mayoccur (Nazina et al., 2008). Copy numbers for mcrA were compara-ble to dsrA numbers. In several wells, the mcrA copy number was
Well10i 10p 13 8i 8p 9 10i 10p 13 14 1 2 3 4 6
DN
A co
pies
/mL
sam
ple
1e+2
1e+3
1e+4
1e+5
1e+6
1e+7
1e+8
1e+9
BacteriaArchaeadsrAmcrA
Fig. 4. Abundance of bacteria, Archaea, sulfate-reducing prokaryotes, and methanogens in samples as determined by qPCR. The error bars represent the standard deviation ofthree replicates.
N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54 51
higher than the 16S bacterial number (Fig. 4), but this pattern wasnot correlated with higher rates of methane generation. Severalwells were sampled during both the November 2010 and April2011 campaigns. Overall, bacterial gene copy numbers were gener-ally higher in April 2011, especially for injection well 10 (Fig. 4).This may indicate seasonal variation in reservoir bacterial num-bers, most likely associated with the water management of the res-ervoir, but longer term study would be needed to validate thesetrends.
3.4. In vitro methane production from labelled hydrocarbons
Microbial enrichment cultures were established from Dagangproduction fluids and amended with 13C labelled aliphatic and aro-matic hydrocarbons to provide unequivocal evidence of methano-genic hydrocarbon degradation by in vitro microbial populationsfrom this reservoir. The isotopic composition (d13C) of methaneand CO2 from enrichment cultures containing 13C labelled n-hexa-decane and 13C-methylnaphthalene (provided as sole carbonsources) became continuously enriched during the incubation.After 90 days of incubation, samples amended with 13C labelled2-methylnaphthalene produced methane with d13C signaturesabove 30‰ (vs. VPDB, Fig. 5B). Labelled methane continued to beproduced over the investigation time of 150 days (Fig. 5B). Isotopi-cally heavy methane also was observed in the enrichments con-taining 13C labelled n-hexadecane and methane d13C values wereapproximately 50‰ PDB after 40 days (Fig. 5A). Although a higherrate of n-alkane degradation is expected compared to 2-MN, the n-alkanes used in this study were uniformly labelled while the 2-MNwas labelled at the methyl position. This in and of itself can ac-count for the earlier emergence of labelled methane in the n-al-kane amended samples. Carbon dioxide however became muchmore enriched in the n-alkane samples, with 13C values observedat 40 days of over 60‰ (Fig. 5). After 400 days, methane and CO2
were both highly labelled (Fig. 5). Methane production rates werequantified for microcosms inoculated with production fluids fromwell 15 and were found to be between 0.83 and 1.02 lmol/ml/d.
4. Discussion
The majority of worldwide oil reservoirs show evidence for bio-degradation (Jones et al., 2008). However, we still understand very
little about the underlying reservoir microbiology, especially undermethanogenic conditions. Understanding these processes mightfacilitate the development of methods for either reducing biodeg-radation, or stimulating metabolic processes for microbially en-hanced oil recovery (MEOR).
4.1. The microbiological potential of the Dagang oilfield for in situmethane production
As a way to increase or sustain oil production, secondary recov-ery by waterflood has been used since its development in the1930s and is still a common production method in many countries.This method entails water injection into the oil bearing strata usinga network of injection wells to maintain reservoir pressure(Belyaev et al., 2004). In the Dagang oilfield complex, injectionwater is separated from production fluids and continuously rein-jected into the formation in a cyclical fashion. The process of oiland water separation before injection takes, on average, 2–3 h.Consequently, the chemical composition of the formation andinjection waters is very similar at this location (Nazina et al.,2007b). This demonstrates that the water cycle is closed and envi-ronmental water injection is minimal. Overall geochemical condi-tions in the Dagang oil field complex represent a favorableenvironment for the recruitment and sustainability of microbialpopulations. The sulfate concentrations are relatively low and sul-fate is probably not a relevant electron acceptor for biodegradation,as sulfide production was not observed in the field during samplingcampaigns. Elevated HCO3 concentrations indicate higher microbialactivities in some of the production blocks (Nazina et al., 2007a).
We hypothesize that secondary recovery methods includingwater recycling and reinjection select for, and maintain, microbialpopulations that can degrade crude oil constituents to methaneunder in situ temperature and pressure. Tracer studies using 13C la-belled aliphatic and polyaromatic hydrocarbons demonstrated thatmicroorganisms in the Dagang formation waters have the intrinsicability to produce methane from these compounds (Fig. 5). Theseresults are consistent with those obtained by Wang et al. (2011)using a mixture of n-alkanes (C15–C20) where high amounts ofmethane were detected in samples collected from the oil complex.Large methane accumulations are commonly associated with bio-degraded oil (Horstad and Larter, 1997; Milkov, 2010, 2011) andattest to our conclusion of biogenic methane production concomi-
Fig. 5. Isotopic values for the methane and CO2 produced from (A) n-hexadecane and (B) methylnaphthalene in vitro. 13C–CH4 and 13C–CO2 correspond to the gases obtainedin the labelled microcosms, whereas 12C–CH4 and 12C–CO2 represent the gases obtained in the unlabelled ones. Error bars represent the standard deviation of threeindependent injections.
52 N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54
tant with anaerobic hydrocarbon degradation. In this respect, thed13C signatures for formation methane are consistent with the mix-ing of secondary microbial and thermogenic gases (Whiticar et al.,1986; Fuhua, 1987; Pallaser, 2000; Larter et al., 2005; Milkov,2011). The high d13C values for CO2 found in most of the samplesfrom the reservoir are consistent with the model proposed by Joneset al. (2008) for hydrogenotrophic methanogenesis, although thismodel was established for methanogenesis under mesophilic con-ditions (<35 �C). The Dagang reservoir has prevailing mesophilic tothermophilic conditions of around 30–60 �C, except in the deepersections, in which the temperature is much higher. Nevertheless,thermophilic hydrogenotrophic methanogens predominate in hightemperature oil reservoirs, for example the Kondiang oil field (Naz-ina et al., 2006) and our clone libraries suggest that thermophilic,hydrogenotrophic methanogens related to Methanobacterium(Smith et al., 1997) are dominant in the Dagang fields. DetectedMethanosarcina related organisms also have the potential to pro-duce methane using electrons from hydrogen. Nazina et al.(2007b) observed methane production from acetate from a hydro-gen and carbon dioxide mixture by cultures from the same reser-voir, paralleling the results obtained by Gray et al. (2009) usingsamples from a gas field in the North Sea. However Gray et al.(2009) did not detect acetoclastic methanogenesis in their cultures.Thermodynamic considerations also suggest that acetoclasticmethanogenesis will become less favored under higher tempera-tures (Larter et al., 2005; Dolfing et al., 2008).
The blocks sampled during this study have a production historybetween 3 and about 40 years and clear differences were observedbetween the total ion chromatogram (TIC) profiles and specific oilbiomarkers in the well under primary production (Block I, well 3)and the wells under production by water flood (Figs. 1 and 2).There are differences in the degradation extent among the wellsunder secondary production. These results may suggest thatchanges in oil composition could be observable within the sameformation on a human timescale and can be related at least partlyto oil production processes. Nazina et al. (1995) identified anaero-bically active microbial biodegradation processes in water floodedmesophilic and thermophilic oil reservoirs from Kazakhstan andwestern Siberia and concluded that the injection of surface watershelped to facilitate biogenic oil degradation. In addition, Whelanet al. (2001) reported significant decreases in the C11–C19 n-alkanesand to a lesser extent heavier components (up to n-C32), in <8 yearswithin oil and gas reservoirs along the Gulf of Mexico coastline.However, it cannot be fully excluded that oil mixing and geological
complexity in the Dagang region (Fajing and Shulin, 1991) affectthese oil profiles to an unknown extent.
Injection and productions waters from Dagang were filtered tocollect microbial biomass for DNA extraction and analyzed usingqPCR. Large numbers of bacterial and Archaeal 16S rRNA geneswere detected in addition to dsrA and mcrA genes (Fig. 4). Anaero-bic microcosms incubated under temperature regimes of 55–60 �Calso produced labelled methane within 40 days after inoculation(Fig. 5). Taken together, this information suggests that a specializedcommunity exists that is capable of completely mineralising ali-phatic, aromatic and polyaromatic hydrocarbons to methane.Methanogenic hydrocarbon degradation has recently become anarea of intense interest from both biological and industrial pointsof view (Gieg et al., 2008; Gray et al., 2010). From a biochemicalperspective, little is understood about the enzymatic pathways in-volved in hydrocarbon activation for polyaromatic degradation un-der reduced conditions, while some progress has been madetowards our understanding of n-alkane and substituted-aromaticactivation (i.e. addition of the hydrocarbon to fumarate, Bellerand Edwards, 2000; Heider, 2007). Development of enrichmentcultures able to degrade PAHs under methanogenic conditions willencourage further study and a primary understanding of themicrobial ecology and diversity involved in oil field systems willhelp to provide groundwork for further biochemical study.
4.2. Chemical evidence for in situ oil biodegradation
An extensive fingerprinting of Dagang oil was carried out usingGC–MS analyzes. These results suggest that the crude oil is heavilydegraded within this complex, like in most of the reservoirs at thistemperature range (Pepper and Santiago, 2001) and that the degra-dation of several compound classes is especially severe. For exam-ple, n-alkanes are almost completely depleted, as normallyobserved in heavily degraded reservoir. However, we also observedsubstantial losses of aromatic fractions in the oil, specifically alkyl-benzenes, alkyltoluenes and low molecular weight polyaromatichydrocarbons. Preferential losses of specific alkylated PAH isomerssuggest that these compounds also are being degraded biologically.Changes in the distribution of trimethylnaphthalenes or methylph-enanthrenes are consistent with previously reported results of oilbiodegradation in a reservoir (Huang et al., 2004). The relativeabundance of these compounds can be associated with their ther-mal stability. However, in samples with a common source and sim-ilar thermal maturity (reflected by the Ts/Tm indices), as in this
N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54 53
case, differences in the chemical distributions are most likely dueto biodegradation (Huang et al., 2004). Isomeric specificity for aer-obic biodegradation of PAHs has been extensively studied and iswell known. For instance, isomers with b-substituents (like 2- or3-methyl-) are more readily degraded than others. However,trends for PAH degradation under reduced reservoir conditionshave not been clearly established. Huang et al. (2004) reported that2,3,6- and 1,3,6-trimethylnaphthalenes, 1,7- and 2,6- + 3,5-di-methylphenanthrenes, or 1,2,8-trimethylphenanthrene were pref-erentially degraded. Nevertheless, these trends are likely affectedby oil chemistry, formation conditions or oil charging/migration(Larter, 2003), along with microbial variability among wells.
Selective methylphenanthrene and methyldibenzothiophenebiodegradation was observed. For example, 4/9-MP and 1-MD,usually the most conserved in biodegraded samples under aerobicconditions (Bayona et al., 1986), were retained in several heavilydegraded samples from our study. This parallels the results ob-tained by Huang et al. (2004) for samples from other oil reservoirs.Similarly, preferential degradation of several sterane epimers hasbeen observed here, consistent with results from the previouslymentioned study. Our GC–MS results, taken together with themicrocosm study, indicate that, although n-alkane degradation oc-curs before any significant degradation of aromatics takes place(Elias et al., 2007; Jones et al., 2008), PAH degradation is responsi-ble for a significant fraction of the methane produced in this oilfield complex. Further experiments will contribute to the assess-ment of metabolic pathways for methanogenic PAH degradationand identify the microorganisms involved, with hope of establish-ing new indices for the assessment of oil biodegradation undermethanogenic conditions.
Acknowledgements
We thank the Robert-Bosch-Foundation for financial support:Grant 32.5.8003.0083.0 ‘‘In Situ Microbial Degradation of Hydrocar-bons in Oil Reservoirs’’. This work was supported in part by theNational Outstanding Youth Research Foundation of China(40925010), two International Joint Key Projects from the NationalNatural Science Foundation of China (40920134003), and theChinese Ministry of Science and Technology (2010DFA12780), andthe National Natural Science Foundation of China (41273092). Fur-thermore, we thank Daniela Zoch, Holger Probst, Ursula Günther,Matthias Gehre and Falk Bratfisch for help with experiments andMarcel Sickert for synthesis of 13C labelled 2-methylnaphthalene.Financial support for F. Gründger was provided by DFG PriorityProgram 1319, Grants KR3311/6-1 and 6-2. N. Jiménez was sup-ported by a Marie Curie Intra-European Fellowship (IEF) withinthe Marie Curie Mobility Actions of the European Commission’sSeventh Framework Program (PIEF-GA-2010-272569). We grate-fully acknowledge the contribution of one anonymous review andAlexei Milkov for critiques and improvement of the manuscript.
Appendix A. Supplementary material
Supplementary data associated with this article can be found,in the online version, at http://dx.doi.org/10.1016/j.orggeochem.2012.08.009.
Associate Editor—Cliff Walters
References
Bayona, J.M., Albaigés, J., Solanas, A.M., Pares, R., Garrigues, P., Ewald, M., 1986.Selective aerobic degradation of methyl-substituted polycyclic aromatichydrocarbons in petroleum by pure microbial culturest. International Journalof Environmental Analytical Chemistry 23, 289–303.
Beller, H.R., Edwards, E.A., 2000. Anaerobic toluene activation by benzylsuccinatesynthase in a highly enriched methanogenic culture. Applied andEnvironmental Microbiology 66, 5503–5505.
Belyaev, S.S., Borzenkov, I.A., Nazina, T.N., Rozanova, E.P., Glumov, I.F., Ibatullin, R.R.,Ivanov, M.V., 2004. Use of microorganisms in the biotechnology for theenhancement of oil recovery. Microbiology 73, 590–598.
Bernard, B.B., Brooks, J.M., Sackett, W.M., 1976. Natural gas seepage in the Gulf ofMexico. Earth and Planetary Science Letters 31, 48–54.
Elias, R., Vieth, A., Riva, A., Horsfield, B., Wilkes, H., 2007. Improved assessment ofbiodegradation extent and prediction of petroleum quality. OrganicGeochemistry 38, 2111–2130.
Fajing, C., Shulin, Z., 1991. Tectonic evolution and hydrocarbon migratino in theHuanghua Basin, NE China. Journal of Petroleum Geology 14, 197–210.
Feisthauer, S., Siegert, M., Seidel, M., Richnow, H.H., Zengler, K., Gründger, F., Krüger,M., 2010. Isotopic fingerprinting of methane and CO2 formation from aliphaticand aromatic hydrocarbons. Organic Geochemistry 41, 482–490.
Feisthauer, S., Vogt, C., Modrzynski, J., Szlenkier, M., Krüger, M., Siegert, M.,Richnow, H.-H., 2011. Different types of methane monooxygenases producesimilar carbon and hydrogen isotope fractionation patterns during methaneoxidation. Geochimica et Cosmochimica Acta 75, 1173–1184.
Fuhua, L., 1987. Zonation of oil and gas in the Huanghua Basin, China. Journal ofPetroleum Geology 10, 327–334.
Gieg, L.M., Duncan, K.E., Suflita, J.M., 2008. Bioenergy production via microbialconversion of residual oil to natural gas. Applied and EnvironmentalMicrobiology 74, 3022–3029.
Gray, N.D., Sherry, A., Hubert, C., Dolfing, J., Head, I.M., 2010. Methanogenicdegradation of petroleum hydrocarbons in subsurface environments:Remediation, heavy oil formation, and energy recovery. Advances in AppliedMicrobiology 72, 137–161.
Gray, N.D., Sherry, A., Larter, S.R., Erdmann, M., Leyris, J., Liengen, T., Beeder, J., Head,I.M., 2009. Biogenic methane production in formation waters from a large gasfield in the North Sea. Extremophiles 13, 511–519.
Head, I.M., Jones, D.M., Larter, S.R., 2003. Biological activity in the deep subsurfaceand the origin of heavy oil. Nature 426, 344–352.
Heider, J., 2007. Adding handles to unhandy substrates: anaerobic hydrocarbonactivation mechanisms. Current Opinion in Chemical Biology 11, 188–194.
Horstad, I., Larter, S.R., 1997. Petroleum migration, alteration, and remigrationwithin Troll Field, Norwegian North Sea. American Association of PetroleumGeologists Bulletin 81, 222–248.
Huang, H., Bowler, B.F.J., Oldenburg, T.B.P., Larter, S.R., 2004. The effect ofbiodegradation on polycyclic aromatic hydrocarbons in reservoired oils fromthe Liaohe Basin, NE China. Organic Geochemistry 35, 1619–1634.
Jiménez, N., Viñas, M., Sabaté, J., Díez, S., Bayona, J.M., Solanas, A.M., Albaiges, J.,2006. The Prestige oil spill. 2. Enhanced biodegradation of a heavy fuel oil underfield conditions by the use of an oleophilic fertilizer. Environmental Science &Technology 40, 2578–2585.
Jones, D.M., Head, I.M., Gray, N.D., Adams, J.J., Rowan, A.K., Aitken, C.M., Bennett, B.,Huang, H., Brown, A., Bowler, B.F.J., Oldenburg, T.B.P., Erdmann, M., Larter, S.R.,2008. Crude-oil biodegradation via methanogenesis in subsurface petroleumreservoirs. Nature 451, 176–180.
Kondo, R., Nedwell, D.B., Purdy, K.J., Silva, S.Q., 2004. Detection and enumeration ofsulphate-reducing bacteria in estuarine sediments by competitive PCR.Geomicrobiology Journal 21, 145–157.
Krüger, M., Frenzel, P., Conrad, R., 2001. Microbial processes influencing methaneemission from rice fields. Global Change Biology 7, 49–63.
Larter, S.R., 2003. The controls on the composition of biodegraded oils. Part 1.Biodegradation rates in petroleum reservoirs in the deep subsurface. OrganicGeochemistry 34, 601–613.
Larter, S.R., di Primio, R., 2005. Effects of biodegradation on oil and gas field PVTproperties and the origin of oil rimmed gas accumulations. OrganicGeochemistry 36, 299–310.
Larter, S.R., Head, I.M., Huang, H., Bennet, B., Jones, M., Aplin, A.C., Murray, A.,Erdmann, M., Wilhelms, A., di Primio, R., 2005. Biodegradation, gas destructionand methane generation in deep subsurface petroleum reservoirs: an overview.Geological Society, London, Petroleum Geology Conference Series 6, 633–639.
Li, S., Wu, T. (Eds.), 1991. Petroleum Geology of China, vol. 4. Petroleum IndustryPress, Dagang Oil Field, Beijing, pp. 74–113.
Lovell, C.R., Piceno, Y., 1994. Purification of DNA from estuarine sediments. Journalof Microbiological Methods 20, 161–174.
Lueders, T., Manefield, M., Friedrich, M.W., 2004. Enhanced sensitivity of DNA- andrRNA-based stable isotope probing by fractionation and quantitative analysis ofisopycnic centrifugation gradients. Environmental Microbiology 6, 73–78.
Milkov, A.V., 2011. Worldwide distribution and significance of secondary microbialmethane formed during petroleum biodegradation in conventional reservoirs.Organic Geochemistry 42, 184–207.
54 N. Jiménez et al. / Organic Geochemistry 52 (2012) 44–54
Milkov, A.V., 2010. Methanogenic biodegradation of petroleum in the West SiberianBasin (Russia): significance for formation of giant Cenomanian gas pools.American Association of Petroluem Geologists Bulletin 94, 1485–1541.
Morris, B.E.L., Herbst, F.-A., Bastida, F., Seifert, J., von Bergen, M., Richnow, H.-H.,Suflita, J.M., 2012. Microbial interactions during residual oil and n-fatty acidmetabolism by a methanogenic consortium. Environmental MicrobiologyReports 4, 297–306.
Nadkarni, M.A., Martin, F.E., Jacques, N.A., Hunter, N., 2002. Determination ofbacterial load by real-time PCR using a broad-range (universal) probe andprimers set. Microbiology 148, 257–266.
Nazina, T., Grigor’yan, A., Feng, Q., Shestakova, N., Babich, T., Pavlova, N.K., Ivoilov,V., Ni, F., Wang, J., She, Y., Xiang, T., Mei, B., Luo, Z., Belyaev, S.S., Ivanov, M.V.,2007a. Microbiological and production characteristics of the high-temperatureKondian petroleum reservoir revealed during field trial of biotehcnology for hteenhancement of oil recovery. Microbiology 76, 340–353.
Nazina, T., Grigor’yan, A., Shestakova, N., Babich, T., Ivoilov, V., Feng, Q., Ni, F., Wang,J., She, Y., Xiang, T., Luo, Z., Belyaev, S., Ivanov, M., 2007b. Microbiologicalinvestigations of high-temperature horizons of the Kongdian petroleumreservoir in connection with field trial of a biotechnology for enhancement ofoil recovery. Microbiology 76, 287–296.
Nazina, T., Pavlova, N., Ni, F., Shestakova, N., Ivoilov, V., Feng, Q., Dongyun, Z.,Prusakova, T., Belyaev, S., Ivanov, M., 2008. Regulation of geochemical activity ofmicroorganisms in a petroleum reservoir by injection of H2O2 or water–airmixture. Microbiology 77, 324–333.
Nazina, T., Shestakova, N., Grigor’yan, A., Mikhailova, E., Tourova, T., Poltaraus, A.,Feng, C., Ni, F., Belyaev, S., 2006. Phylogenetic diversity and activity of anaerobicmicroorganisms of high-temperature horizons of the Dagang oil field (P. R.China). Microbiology 75, 55–65.
Nazina, T.N., Ivanova, A.E., Borzenkov, I.A., Belyaev, S.S., Ivanov, M.V., 1995.Occurrence and geochemical activity of microorganisms in high-temperature,water-flooded oil fields of Kazakhstan and Western Siberia. GeomicrobiologyJournal 13, 181–192.
Ofosu-Asante, K., Stock, L.M., 1987. A convenient and selective method for reductivedeuteration of aryl carbonyl compounds. Journal of Organic Chemistry 52,2938–2939.
Pallaser, R.J., 2000. Recognising biodegradation in gas/oil accumulations throughthe d13C composition of gas components. Organic Geochemistry 31, 1363–1373.
Pepper, A., Santiago, C., 2001. Impact of biodegradation on petroleum explorationand production, Observations and outstanding problems. In: Abstracts of EarthSystems Processes Conference, Geological Society of London, Geological Societyof America, Edinburgh, 24–28 June.
Douglas, G.S., Butler, E.L., 1994. 17a(H), 21b(H)-hopane as a conserved internalmarker for estimating the biodegradation of crude-oil. Environmental Science &Technology 28, 142–145.
Richnow, H.H., Annweiler, E., Michaelis, W., Meckenstock, R.U., 2003. Microbialin situ degradation of aromatic hydrocarbons in a contaminated aquifer
monitored by carbon isotope fractionation. Journal of Contaminant Hydrology65, 101–120.
Röling, W.F.M., Head, I.M., Larter, S.R., 2003. The microbiology of hydrocarbondegradation in subsurface petroleum reservoirs: perspectives and prospects.Research in Microbiology 154, 321–328.
Schippers, A., Neretin, L.N., 2006. Quantification of microbial communities in near-surface and deeply buried marine sediments on the Peru continental marginusing real-time PCR. Environmental Microbiology 8, 1251–1260.
Smith, D.R., Doucette-Stamm, L.A., Deloughery, C., Lee, H., Dubois, J., Aldredge, T.,Bashirzadeh, R., Blakely, D., Cook, R., Gilbert, K., Harrison, D., Hoang, L., Keagle,P., Lumm, W., Pothier, B., Qiu, D., Spadafora, R., Vicaire, R., Wang, Y.,Wierzbowski, J., Gibson, R., Jiwani, N., Caruso, A., Bush, D., Reeve, J.N., 1997.Complete genome sequence of Methanobacterium thermoautotrophicum DH:functional analysis and comparative genomics. Journal of Bacteriology 179,7135–7155.
Takai, K., Horikoshi, K., 2000. Rapid detection and quantification of members of thearchaeal community by quantitative pcr using fluorogenic probes. Applied andEnvironmental Microbiology 66, 5066–5072.
Vincent, C., Dai, S., Wenying, C., Rongshu, Z., Guosheng, D., Xu, R., Vangkilde-Pedersen, T., Dalhoff, F., 2009. Carbon dioxide storage options for the COACHproject in the Bohai Basin, China. Energy Procedia 1, 2785–2792.
Wang, L.-Y., Gao, C.-X., Mbadingaa, S.M., Zhoua, L., Liua, j.-F., Gub, J.-D., Mua, B.-Z.,2011. Characterization of an alkane-degrading methanogenic enrichmentculture from production water of an oil reservoir after 274 days of incubation.International Biodeterioration and Biodegradation 65, 444–450.
Whelan, J.K., Eglinton, L., Kennicutt II, M.C., Qian, Y., 2001. Short-time-scale (year)variations of petroleum fluids from the U.S. Gulf Coast. Geochimica etCosmochimica Acta 65, 3529–3555.
Whiticar, M.J., Faber, E., Schoell, M., 1986. Biogenic methane formation in marineand freshwater environments: CO2 reduction vs. acetate fermentation – isotopeevidence. Geochimica et Cosmochimica Acta 50, 693–709.
Widdel, F., Bak, W., 1992. The Prokaryotes. Nature Publishing Group, pp. 3352–3379.
Williams, J.A., Bjorøy, M., Dolcater, D.L., Winters, J.C., 1986. Biodegradation in southTexas Eocene oils—effects on aromatics and biomarkers. Organic Geochemistry10, 451–461.
Zengler, K., Richnow, H.H., Rossello-Mora, R., Michaelis, W., Widdel, F., 1999.Methane formation from long-chain alkanes by anaerobic microorganisms.Nature 401, 266–269.
