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Microbial Hydrocarbon Degradation: Efforts to Understand Biodegradation in Petroleum Reservoirs

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Isabel Natalia Sierra-Garcia and Valéria Maia de Oliveira

Submitted: June 28th, 2012 Published: June 14th, 2013

DOI: 10.5772/55920

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1. Introduction

The understanding of the phylogenetic diversity, metabolic capabilities, ecological roles, and community dynamics taking place in oil reservoir microbial communities is far from complete. The interest in studying microbial diversity and metabolism in petroleum reservoirs lies mainly but not only on providing a better comprehension of biodegradation of crude oils, since it represents a worldwide problem for petroleum industry. Generally, biodegradation of oil affects physical and chemical properties of the petroleum, resulting in a decrease of its hydrocarbon content and an increase in oil density, sulphur content, acidity and viscosity, leading to a negative economic consequence for oil production and refining operations [1,2]. Another important point for studying biodegradation lies on its important role in the global carbon cycle and the direct impact on bioremediation of polluted ecosystems. Furthermore, many of the enzymes involved in the degradation pathways are considered key catalysts in industrial biotechnology [3].

Despite these motivations and long recognition of petroleum as a the most important “primary energy” source, at present, microorganisms and factors involved in biodegradation of crude oil hydrocarbons in petroleum reservoirs are still not fully understood. The inaccessibility and complex microbiological sampling of petroleum reservoirs as well as the inherent limitations of the traditional culturing methods conventionally employed can explain this fact. Culture-based techniques have traditionally been the primary tools utilized for studying the microbiology of terrestrial and subsurface environments [4], which allowed the recovery and documentation of a large collection of bacteria capable of hydrocarbon utilization. Studies of numerous aerobic and anaerobic bacterial isolates have revealed mechanisms, which allow them to degrade specific classes of the highly diverse range of hydrocarbon compounds. Therefore, all we know about the degradation of petroleum compounds has come from studying isolated microorganisms. Here, we provide an overview of what is currently known about the mechanisms of aerobic and anaerobic degradation of hydrocarbons, as a result from biochemical and genomic approaches, we give a perspective of the petroleum microbial diversity unraveled so far, and finally we discuss the common oil reservoir characteristics that can be used to predict the most probable mechanism of degradation into deep petroleum reservoirs.

It is well known that microbial diversity in environment is several orders of magnitude higher than the one assumed based on previous cultivation methods [5]. A particularly large number of novel techniques have been developed, which now allow the determination of the in situmicrobial diversity and activity on a particular site, screening for a particular gene or activity of interest, gene quantification, and DNA and mRNA sequencing and analysis from total communities. This book chapter will address how the implementation of such culture-independent molecular methods allow the access to the microbial diversity and metabolic potential of microorganisms and bring novel information about microbial diversity and new pathways involved in biodegradation processes taking place in petroleum reservoirs. This information will certainly contribute to a broader perspective of the biodegradation processes and corroborate with previous findings that degradation of pollutants in many cases is carried out by microbial consortia rather than a single species [6], where key species and catabolic genes are often not identical to those that have been isolated and described in the laboratory [7, 8].

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2. Microbial diversity in oil reservoirs

Recognition of indigenous microbiota harbored by oil reservoirs has been discussed for a long time. Actually, determining the nature of isolated microorganisms from oil reservoirs (indigenous or nonindigenous) is a difficult issue concerning petroleum microbiologists. The reasons for this controversy rely mainly on the difficulty of aseptic sampling in deep oil reservoirs. This means that microorganisms observed in oil field fluids conceivably could be contaminants introduced during drilling operations and/or during sample retrieval, or could be material sloughed from biofilms growing in installed pipes. Another reason for skepticism is the commonplace practice of ‘‘water- flooding’’ (injection of surface waters or re-injection of natural formation waters to maintain reservoir pressure for oil production); since in this case microbes would be introduced during injection and therefore would not necessarily represent indigenous species [9].

In addition to this controversy, there is the fact that petroleum reservoirs are considered extreme environments where in situconditions, like high pressure, temperature, salinity and anaerobic conditions, are considered as inhospitable to microbial activity. In fact, perception of deep subsurface as a sterile environment has only changed during the past two decades with the increasing awareness of the ability of microbes to colonize extreme environments. Actually, with the use of more sophisticated and appropriate sampling and cultivation techniques, as well as the application of molecular biological techniques to oil field fluids, the dogma of the sterile deep subsurface has been dispelled [9]. Rather, it has become clear that many oil reservoirs do harbor indigenous microbes (e.g. the genera Geotogaand Petrotogaisolated only from oil reservoirs) [10]. Nowadays it is clear that worldwide petroleum reserves are dominated by deposits that have been microbially degraded over geological time and biodegraded petroleum reservoirs represent the most dramatic manifestation of the deep biosphere [11]

In spite of the polemics on which micro-organisms would actually be native and which would be contaminants in oil reservoirs, a wide range of microbial taxonomic groups have been identified in oil reservoirs geographically distant using traditional techniques adapted to in situconditions, as described by L'Haridon et al. [12], Grassia et al. [13] and reviewed by Magot et al [14], or combined with cultivation-independent molecular methods, as reported by Orphan et al. [15]. Table 1 summarizes the various physiological and taxonomical groups and species that have been isolated from oil reservoirs.

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3. Aspects from oil reservoir determining microbial degradation

For a long time, the mechanism considered to be prevalent for oil degradation in petroleum reservoirs was the well documented aerobic microbial metabolism and it has long been thought that the flow of oxygen through meteoric waters was necessary for in-reservoir petroleum biodegradation [16]. This mechanism has been widely accepted despite the fact that oxygen would likely be consumed by oxidation of organic matter in near surface sediments and therefore, would be very unlikely for oxygen to reach deep petroleum reservoirs [11].

Recently, the discovery of the ability of microorganisms to degrade anaerobically hydrocarbon oil components and the detection of metabolites characteristic of anaerobic hydrocarbon degradation in oil samples from biodegraded reservoirs, but not in non-degraded reservoirs or aerobically degraded oils [11], have provided valuable information to determine the processes involved in the degradation of oil reservoirs. Nowadays, evidences of such degradation through anaerobic rather than aerobic processes are becoming more substantial and compelling [17].

It is known that microorganisms in anaerobic conditions can use a variety of final electron acceptors, including nitrate, iron, sulfate, manganese and, more recently, chlorate. Anaerobic degradation has also been coupled to methanogenesis, fermentation and phototrophic metabolism but growth of these microorganisms and, therefore, biodegradation rates are significantly lower compared to aerobic degraders. These anaerobic processes have been demonstrated in surface sediments and pure cultures or enrichments in laboratories [18] and all of them potentially play a role in oil biodegradation in anoxic petroleum reservoirs [11]. However, nitrate, like oxygen, is highly reactive and would likely be completely consumed before it could reach the oil reservoir [17]. In deep reservoirs, the supply of large amounts of Fe(III) or manganese(IV) via meteoric water influx are unlikely due to poor solubility and slow water recharge rates in subterranean cycles. Therefore, iron and manganese, which could be used as electro acceptors for oil oxidation, are unlikely to be responsible for significant compositional changes in the oil, considering their limited availability in the reservoir. Accordingly, oil degradation linked to sulfate reduction and methanogenic would therefore explain the consistent hydrocarbon compositional patterns seen in degraded oils worldwide [17]. Sulfate arises from geological sources, such as evaporitic sediments and limestone, or from the injection of seawater for pressure stabilization, and may lead to significant oil degradation and increased residual-oil sulfur content. Methanogenic oil degradation, on the other hand, does not require external electron acceptors and leads to less overall souring of the oil reservoir. Several studies have described in vitromethanogenic degradation of crude oil related compounds [19, 20] Jones et al., 2008), including n-alkanes [21, 20] and aromatic hydrocarbons [17].

OrganismTaxonomical groupMetabolismOriginReference
Thermodesulforhabdus norvegicusDeltaproteobacteriaSulfate-reducerOil field in Norway[22]
Desulfacinum infernumDeltaproteobacteriaSulfate-reducerNorth see petroleum reservoir near Scotland[23]
Desulfomicrobium norvegicumDeltaproteobacteriaSulfate reducerPetroleum reservoir in Canada[24]
Desulfovibriosp.DeltaproteobacteriaSulfate reducerPetroleum reservoir in Canada[24]
Dethiosulfovibrio peptidovoransBacteria, SynergistetesSulfate reducerOil well in the Emeraude oilfield in Congo, Central Africa,[25]
Desulfotomaculum thermocisternumBacteria, FirmicutesSulfate reducerOil reservoir in the North sea[26]
Deferribactersp.Bacteria, DeferribacteresSulfate reducerCalifornia oil fields[15]
Halanaerobium congolenseBacteria, FirmicutesThiosulfate- and sulfur-reducing bacteriumAfrican oil field[27]
Thauera phenylaceticaBetaproteobacteriaNitrate reducerPetroleum reservoir in Canada[24]
Pseudomonas stutzeriGammaproteobacteriaNitrate reducerPetroleum reservoir in Canada[24]
Garciella nitratireducensBacteria, FirmicutesNitrate reducerOil field in Tabasco, Gulf of Mexico[28]
Geobacillus subterraneus, Geobacillus uzenensisBacteria, FirmicutesNitrate reducerPetroleum reservoir in China[29]
Lactosphaera pasteuriiBacteria, FirmicutesFermentativePetroleum reservoir in Canada[24]
Propionicimonas paludicolaBacteria, FirmicutesFermentativePetroleum reservoir in Canada[24]
AnaerobaculumBacteria, SynergistetesFermentativeCalifornia oil fields[15]
Thermococcus sp.Archaea, EuryarchaeotaFermentativeCalifornia oil fields[15]
Thermococcus sibericusArchaea, EuryarchaeotaFermentativePetroleum reservoir in Western Siberia[30]
Petrotoga sp.Bacteria, ThermotogaeFermentativeCalifornia oil fields[15]
Petrotoga olearia; P. sibericaBacteria, ThermotogaeFermentativePetroleum reservoir in Western Siberia[12]
ThermoanaerobacterBacteria, FirmicutesFermentativeCalifornia oil fields[15]
Thermotoga sp.Bacteria, ThermotogaeFermentativeCalifornia oil fields[15]
Thermosipho geoleiBacteria, ThermotogaeFermentativePetroleum reservoir in Western Siberia[12]
Anaerobaculum thermoterrenumBacteria, SynergistetesFermentativeOil well in Utah[23]
Fusibacter paucivoransBacteria, FirmicutesFermentativeOil well in the Emeraude oilfield in Congo, Central Africa[31]
Thermovirga lieniiBacteria, SynergistetesFermentativeOil reservoir in the North sea[32]
MethanococcusArchaea, EuryarchaeotaMethanogenCalifornia oil fields[15]
Methanococcus thermolithotrophicusArchaea, EuryarchaeotaMethanogenNorth sea old field in Norway[33]
MethanoculleusArchaea, EuryarchaeotaMethanogenCalifornia oil fields[15]
MethanobacteriumArchaea, EuryarchaeotaMethanogenCalifornia oil fields[15]

Table 1.

Summary of bacteria isolated from oil reservoirs worldwide.

Deep subsurface environments such as petroleum reservoirs are logistically much more difficult to study than contaminated shallow subsurface environments [17]. Since in many biodegraded petroleum reservoirs most biodegradation occurs close to the oil water transition zone, it has been proposed that the oil–water transition zone (OWTZ) provides suitable physical and chemical conditions for microbial activity [17].

There are other physical and chemical parameters influencing in situbiodegradation. Temperature is one of the main factors which limits oil degradation in reservoir, and, empirically, it has been repeatedly observed that biodegradation does not occur in oil reservoirs with in situtemperatures >80-90°C [34]. Salinity is another factor that affects in-reservoir oil biodegradation, especially in combination with temperature [13]. Typically, reservoirs with highly saline waters show limited oil biodegradation [11]. This is consistent with the observations that it has not been possible to cultivate microorganisms from reservoir waters with salinity greater than 100 g/L [13]. Pressure seems to be a less limiting factor, except that it may select for certain physiological types and influences the pH of pore waters by increasing dissolution of CO2 [9]. The availability of electron donors and acceptors governs the type of bacterial metabolic activities within oil field environments [14]. The potential electron donors include CO2, hydrocarbons, H2 and numerous organic molecules. Availability of fixed nitrogen is unlikely to limit microbial activity in reservoirs. However, the availability of water-soluble nutrients, like phosphorus and/ or oxidants (terminal electron acceptors such as ferrous iron, sulfate or CO2), is more likely to limit in situmicrobial activity [9]. Nonetheless, physiological characteristics of microorganisms indigenous to petroleum reservoirs shed light on the conditions under which petroleum degradation may occur and the potential degradation mechanisms.

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4. Hydrocarbon degradation

Hydrocarbons are understood as the compounds that consist exclusively of carbon and hydrogen. Because of the lack of functional groups, hydrocarbons are largely apolar and exhibit low chemical reactivity at room temperature. Differences in their reactivities are primarily determined by the occurrence, type and arrangement of unsaturated bonds. Therefore, in this chapter, we will use the common way to classify hydrocarbons according to their bonding features: i) aliphatic group, which includes straight-chain (n-alkanes), branched-chain and cyclic compounds and ii) aromatic group which includes mono or polycyclic hydrocarbons an many important compounds which also contain aliphatic hydrocarbon chains (e. g., alkylbenzenes).

Already a century ago, bacterial isolates had been reported to use aliphatic and aromatic hydrocarbons as sole carbon and energy sources [35]. Since then, numerous aerobic, and also anaerobic, bacterial isolates have been studied in order to understand the mechanisms which allow them to degrade specific members of the highly diverse aliphatic and aromatic compounds. Degradation by such isolates has been investigated thoroughly and results have revealed that they can completely degrade most classes of hydrocarbons, including alkanes, alkenes, alkynes and aromatic compounds. Such degradation can occur aerobically, with oxygen, or anaerobically, with nitrate, ferric iron, sulfate or other electron acceptors [36].

Efforts to overview the metabolism of hydrocarbons in microorganisms are confronted with the chemical diversity of such compounds and their reactivities, as well as with various microbial life styles [36]. The study of biodegradation is conventionally treated in separate areas: aliphatic vs. aromatic hydrocarbons, aerobic vs. anaerobic degradation pathways, physiology and overall metabolic pathways vs. enzymatic mechanisms and structures, often with limited knowledge and data exchange. Nonetheless, each of these study areas deals with the same central point that is the ‘‘metabolic challenge’’ to guide an apolar, unreactive compound composed only of carbon and hydrogen into the metabolism [36]. The hydrocarbon must be first functionalized and currently it has been recognized that there is a surprisingly diversity of reactions of activation that had evolved in microorganisms (Table 2).

Mechanisms for hydrocarbon activation
AerobicAnaerobic
Short-Chain non-methane alkanes C2-C10• Non-heme iron monooxygenase similar to sMMO (C2-C9)
• Copper-containing monooxygenase similar to pMMO (C2-C9)
• Heme-iron monooxygenases (also refered as soluble Cytochrome P450 (C5-C12)
• Fumarate addition
Long-Chain alkanes >C10• Heme-Monooxygenase (P450 type)
• [Fe2]-Monooxygenase
• Non-heme iron monooxygenase (AlkB-related) (C3-C13 or C10-C20)
• Flavin-binding monooxygenase (AlmA) (C20- C36)
• Thermophilic flavin-dependent monooxygenase (LadA) (C10-C30)
• Fumarate addition
• Carboxylation
Aromatic hydrocarbons• [Fe]-Dioxygenase
• [Fe2]-Monooxygenase
• [Flavin]-Monooxygenase
• Fumarate addition
• Hydroxylation
• Carboxylation

Table 2.

Overview of aerobic and anaerobic mechanisms for hydrocarbon activation in bacteria.

Mechanisms for hydrocarbon activation are basically different in aerobic and anaerobic microorganisms. Under oxic conditions, hydrocarbon metabolism is always initiated using molecular oxygen as a co-substrate in mono- or dioxygenase reactions that enable the terminal or sub-terminal hydroxylation of aliphatic alkane chains or the mono or dihydroxylation of aromatic rings [37]. In the hydrocarbon activation under anoxic conditions, some proposed reactions comprise: (1) addition to fumarate by glycyl-radical enzymes, (2) methylation of unsubstituted aromatics, (3) hydroxylation with water by molybdenum cofactor containing enzymes of an alkyl substituent via dehydrogenase, and (4) carboxylation catalyzed by yet- uncharacterized enzymes which may actually represent a combination of reaction (2) followed by reaction (1) [38; 37]. Although all these mechanisms of hydrocarbon anaerobic activation have been proposed, the required signature metabolites and enzymes involved have been characterized only for (1) addition to fumarate (demonstrated for toluene, xylene, ethylbenzene, methylnaphthalene, alkanes and alicyclic alkanes); for (3) hydroxylation (demonstrated for ethylbenzene); and for (4) carboxylation (demonstrated for benzene and naphtalene) [39].

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5. Biochemical and genetic pathways of microbial hydrocarbon degradation

The enzymatic reactions involved in the aerobic degradation of hydrocarbons by bacteria have been extensively studied for several decades [37]. Genes encoding enzymes for degradation are relatively well understood for aerobic and easily cultivable microorganisms, particularly for a Pseudomonasstrain, known as P. putidaGPo1, as well as for the strains Acinetobactersp. ADP1 and Mycobacterium tuberculosisH37Rv [39, 40]. On the other hand, the anaerobic hydrocarbon degradation has gained more attention since is supposed to be the predominant mechanism occurring in several polluted environments and oil reservoirs. However, its study is an incipient area because of the peculiarities of the reservoir environment and difficulties that arise from attempts to characterize these communities. Nevertheless, several bacteria from other environments able to use alkanes as carbon source in the absence of oxygen have been described in the last few years [41], but anaerobic bacteria able to degrade hydrocarbons under conditions found in deep petroleum reservoirs have not been isolated so far [2]. Figure 1 represents an overview of the main mechanisms and pathways used by microorganisms to degrade hydrocarbon compounds under aerobic and anaerobic conditions.

5.1. Aerobic degradation

5.1.1. Aliphatic hydrocarbons

In most degradation pathways described, the substrate n-alkane is oxidized to the corresponding alcohol by substrate-specific terminal monooxygenases/hydroxylases. The alcohol is then oxidized to the corresponding aldehyde, and finally converted into a fatty acid. Fatty acids are conjugated to CoA and subsequently processed by β – oxidation to generate acetyl-CoA [42, 40]. Subterminal oxidation has also been described for both short and long-chain alkanes [40]. Both terminal and sub-terminal oxidation can coexist in some microorganisms [41]. Initial terminal hydroxylation of n-alkanes in bacteria can be carried out by enzymes belonging to different classes, named: (1) propane monooxygenase (C3), (2) different classes of butane monooxygenase (C2-C9), (3) CYP153 monooxygenases (C5-C12), (4) AlkB-related non-heme iron monooxigenase (C3-C10 or C10-C20), (5) flavin-binding monooxigenase AlmA (C20-C36), (6) flavin-dependent monooxygenase LadA (C10-C30), (7) copper flavin-dependent dioxygenase (C10-C30) [43].

Figure 1.

Pathways for aerobic and anaerobic bacterial degradation of hydrocarbon compounds. Two arrows represent more than one reaction.

Among all the alkane activating enzymes, the integral membrane non-heme iron monooxygenase (AlkB) is the best characterized one. Microorganisms degrading medium (C5-C11) and long (>C12)-length alkanes have been frequently related to the presence of alkB genes and that is why the presence of such genes have been widely used as functional biomarker for the characterization of aerobic alkane-degrading bacterial populations in several environmental samples [44, 45] and in bioremediation experiments [46, 47]. The degradation pathway of the alksystem was first described in Pseudomonas putidaGPo1 (formerly identified as P. oleovoransGPo1), where it is located on the OCT plasmid. In this model system, OCT plasmid contains two operons: alkBFGHJKL and alkST [48]. The first operon encodes two components of the alksystem, a particulate non-heme integral membrane alkane monooxygenase (AlkB) and the soluble protein rubredoxin (AlkG), as well as other enzymes involved in further steps. The second operon encodes for a rubredoxin reductase (AlkT and AlkS), which regulates the expression of the alkBFGHJKL operon [48, 49]. Since this system was described, AlkB homologous have been found in many alkane-degrading α- β – and γ –Proteobacteria and high G + C content Gram-positive bacteria (Actinobacteria) [39] and an increasing collection of alkane hydroxylase gene sequences has allowed the diversity analysis of hydrocarbon-degrading microbial populations in different ecosystems. However, comparisons of cloned alkB genes or gene fragments have showed that sequence diversity is very high, even among alkB genes within the same species [50].

In despite of the relevance of alkBgenes as a functional biomarker of alkane-degrading bacterial communities, knowledge on the presence and diversity of alkB genes in oil reservoirs is scarce. Tourova et al. [51] analysed alkB diversity in thermophilic bacterial strains of the genus Geobacillusisolated from oil reservoirs or hot springs. They detected, for the first time, sets of alkB gene homologous in thermophilic bacteria, and some strains showed different homologous within the same genome. This fact was explained by the occurrence of horizontal gene transfer among these bacteria. Recently, Li et al. [52] aimed to evaluate alkB gene diversity and distribution in production water from 3 oilfields in China through a specific PCR-DGGE method. Results showed that sequences found in the water samples were similar to alkB genes from other corresponding alkane-degrading strains. But at the same time, they showed the presence of a considerable genetic diversity of alkB genes in the wastewater as evidenced by a total of 13 unique DNA bands detected. Studies on the degradation of alkanes in oil reservoirs are currently in a start point, but in the future they certainly will help to understand the process of degradation in oil reservoir.

In comparison to the few efforts in studying alkB system in oil reservoirs, much less is known about the presence of the other enzymatic systems previously listed, which have been described for aerobic degradation of n-alkanes in isolated bacteria or laboratory microcosms. For the most recent elucidated systems for alkane oxidation, named almAand ladAgenes, nothing is known about the environmental distribution of these type of genes in petroleum contaminated sites [53] or oil fields, although the LadA complete degradation pathway has been characterized through genome and proteome analysis of Geobacillus thermodenitrificansNG80-2, a thermophilic strain isolated from a deep oil reservoir in Northern China [54]. Currently, it is believed that there are enzyme systems for alkane degradation which have still not been characterized and that may include new proteins unrelated to those already known [41]. Moreover, in many alkane degraders more than one alkane oxidation system have been observed, which have been reported exhibiting overlapping substrate ranges [39, 40]. These observations point out that in order to characterize and explore metabolic diversity and functions involved in alkane degradation one should take into consideration the high diversity of enzymes capable of initiating such metabolism.

5.1.2. Aromatic hydrocarbons

The aerobic bacterial catabolism of aromatic compounds involves a wide variety of peripheral pathways that activate structurally diverse substrates into a limited number of common intermediates that are further cleaved and processed by a few central pathways to the central metabolism of the cell [55]. Metabolic pathways and encoding genes responsible for the degradation of specific members of a highly diverse range of aromatic compounds have been characterized for many isolated bacterial strains, predominantly from the Proteobacteria and Actinobacteria phyla [56]. Degradation by such isolates is typically initiated by members of one of the three superfamilies: the Rieske non-heme iron oxygenases (RNHO), the flavoprotein monooxygenases (FPM) and the soluble diiron multicomponent monooxygenases (SDM). Further metabolism is achieved through di- or trihydroxylated aromatic intermediates. Alternatively, activation is mediated by CoA ligases where the formed CoA derivates are subjected to selective hydroxylation [58, 53]. In the case of hydrophobic pollutants, such as benzene, toluene, naphthalene, biphenyl or polycyclic aromatics, aerobic degradation is usually initiated by activation of the aromatic ring through oxygenation reactions catalyzed by RNHO enzymes or, as intensively described for toluene degradation, through members of SDM enzymes [56].

Further intermediates can be catalyzed by two kinds of enzyme, intradiol and extradiol dioxygenases, which represent two classes of phylogenetically unrelated proteins [58]. These enzymes are key enzymes in the degradation of aromatic compounds, and many of such proteins and their encoding sequences have been described, purified and characterized in the last decades [56]. While all intradiol dioxygenases described so far belong to the same superfamily, the extradiol dioxygenases include at least three members of different families. Type I extradiol dioxygenases (e.g. catechol 2,3-dioxygenases and 1,2-dioxygenases) belong to the vicinal oxygen chelate superfamily enzymes. Type II extradiol dioxygenases are related to LigB superfamily (e.g. protocatechuate 4,5-dioxygenases) and the type III enzymes belongs to the cupin superfamily (e.g. gentisate dioxygenases) [53]. However, members of novel superfamilies performing crucial steps in aromatic metabolic pathways are still being discovered [56, 53].

The knowledge of metabolic properties of isolates has allowed the monitoring of the ability of microorganisms to mineralize aromatic hydrocarbons in soils. Typically, these studies have used primers designed based on conserved gene regions and focused on RNHO or SDM as targets for initiating degradation, or on Extradiol dioxygenases (EXDO) cleaving the aromatic ring [59]. These studies range from those searching for a narrow range of genes similar or identical to those observed in type strains using non-degenerated primers to those searching for subfamilies of homologous genes using degenerated primers [59]. However, due to the immense heterogeneity of such enzymes [57], there will never be a pair of primers that will reliably cover the huge diversity of a catabolic gene family in nature [53].

5.2. Anaerobic degradation

5.2.1. Aromatic hydrocarbons

We have already described the main mechanism for degradation of aromatic compounds in aerobic conditions, where oxygen is not only the final electron acceptor but also co-substrate of two key processes: hydroxylation and cleavage of the aromatic ring by oxygenases. In contrast, in the absence of oxygen, microorganisms use a complete different pathway, based in reductive reactions to attack the aromatic ring [61].

The biochemistry of some anaerobic degradation pathways of aromatic compounds has been studied to some extent; however, the genetic determinants of all these processes and the mechanisms involved in their regulation are much less studied [55]. Recent advances in genome sequencing have led to the complete genetic information for six bacterial strains that are able to anaerobically degrade aromatic compounds using different electron acceptors and that belong to different taxonomic groups of bacteria: denitrifying betaproteobacteria, Thauera aromaticaand Azoarcussp. EbN1, two alphaproteobacteria, the phototroph Rhodopseudomonas palustrisstrain CGA009 and the denitrifying Magnetospirillum magneticumstrain AMB-1, and two obligate anaerobic deltaproteobacteria, the iron reducer Geobacillus metallireducensGS-15 and the fermenter Syntrophus aciditrophicusstrain SB [55]. It is worth remembering that, in recent years, important inferences and generalizations have been made about the genetics involved in hydrocarbon metabolism based on these isolated bacteria under conventional laboratory conditions. However, potential novel genes, enzymes and metabolic pathways responsible for degradation processes are probably harbored by yet uncultivated bacteria.

The best understood and apparently the most widespread of these anaerobic mechanisms is the radical-catalyzed addition of fumarate to hydrocarbons, yielding substituted succinate derivatives. This reaction has been recognized for the activation of several alkyl-substituted benzenes as well for n-alkanes [62]. However, understanding of this fumarate-dependent hydrocarbon activation is most advanced in the case of toluene. The key enzyme in this process is the enzyme benzylsuccinate synthase. All enzymes required for β-oxidation of benzylsuccinate are encoded by the bbsoperon. Subsequent degradation of benzoyl-CoA proceeds via reductive dearomatization, hydrolytic ring cleavage, β-oxidation to acetyl-CoA units and terminal oxidation to Co2 [63]. In contrast to the anaerobic metabolism of toluene, degradation of ethylbenzene (and probably other alkylbenzenes with carbon chain of at least 2) is entirely different, despite the chemical and structural similarities between the two compounds, and involves a direct oxidation of the methylene carbon via (S)-1-phenylethanol to acetophenone [55]. Ethylbenzene is anaerobically hydroxylated and dehydrogenated to acetophone, which is then carboxyled and converted to benzoylCoA as the first common intermediate of the two pathways [62].

Genetics of the enzymatic system have been only characterized for these two mechanisms for anaerobic hydrocarbon activation. Genes encoding pathways that involve fumarate addition are typically organized in two operons. One operon includes the three structural genes of the protein catalyzing fumarate addition and the other includes genes required for converting succinate derivates to benzoyl-CoA [64]. Gene sequences and organization are relatively conserved among nitrate-reducing bacteria but differ somewhat from those of the iron reducer G. metallireducens[64] and substantially from those of the hexane-degrading nitrate reducer strain HxN1 [65]. Hydrocarbon dehydrogenation pathway is also organized in two operons. One operon contains the structural genes for the first two reactions (ethylbenzene dehydrogenase and 1-phenylethanol dehydrogenase) and the other contains the structural genes for acetophone carboxylase [64].

Kane et al. [66] developed the first real-time polymerase chain reaction (PCR) method to quantify hydrocarbon utilizers based on bssA genes of nitrate-reducing Betaproteobacteria. Since then, there have been several additional studies investigating the presence and/or distribution of anaerobic hydrocarbon utilizers in anaerobic environments via functional gene surveys of bssA, extending the range of detectable hydrocarbon-degrading microbes to iron and sulfate-reducing Deltaproteobacteria and revealing partially novel, site specific degrader populations [67, 68]. Other bssA-based detection studies in impacted environments, as well as studies that combine field metabolomics and molecular tools, are described by other authors [69, 70, 71]. Despite of the role of benzylsuccinate synthase in aromatic hydrocarbon degradation and its use as a biomarker are well documented, there is no study on the presence of this gene in oil reservoirs.

5.2.2. Aliphatic hydrocarbons

Anaerobic degradation of alkanes has not been extensively studied as for some aromatic compounds. The presumable reasons include the greater attention given to BTEX compounds (benzene, toluene, ethylbenzene and xylenes) because of their classification as priority pollutants [71], also the fact that anaerobic growth with n-alkanes is even slower than that with the alkylbenzenes, and finally the fact that long chain alkanes are poorly soluble and often prevents the cultivation of cells homogeneously in the medium [72]. However, anaerobic degradation of alkanes is equally relevant, since alkanes are quantitatively the most important hydrocarbon components of petroleum, and some are acutely toxic and difficult to remediate [71]. Several anaerobic bacteria capable of degrading n-alkanes with 6 or more carbons in length, particularly hexadecane (C16), using sulfate or nitrate as electron acceptors have been isolated [72, 73].

The two main mechanisms of anaerobic degradation of n-alkanes described involve unprecedented biochemical reactions that differ completely from those employed in aerobic hydrocarbon metabolism [73]. The first involves activation at the subterminal carbon of the alkane by the addition of fumarate, analogously to the formation of benzyl succinate during anaerobic degradation of toluene, however further reactions are completely different involving dehydrogenation and hydration [72]. Studies conducted with established axenic cultures have indicated that anaerobic metabolism of oil allkanes predominantly proceeds via addition of fumarate to the double bound [72]. Although alkylsuccinate metabolites have rarely been detected in oil reservoir fluids [74, 75], they have been reported in oil-contaminated environments as well as in oilfield facilities, where their detection is indicative of in situmicrobial degradation of oil alkanes [71, 75]. Alkylsuccinic acids as intermediates of anaerobic alkane oxidation were first studied by Gieg and Suflita [76] when surveying these metabolites in aquifers contaminated with condensate gas, natural gas liquids, gasoline, diesel, alkanes and BTEX. They found alkylsuccinates originating from C3 to C11 alkanes, as well as putative metabolites originating from compounds with one degree of unsaturation, such as alkenes or alicyclic alkanes. Since this report, other studies have detected alkylsuccinate derivates in petroleum contaminated groundwater systems [76], coal beds [70] and oil fields [74, 77]. The formation of alkylsuccinates is catalyzed by a strictly anaerobic glycyl radical enzyme which has been termed as alkylsuccinate synthase or (1-methyl-alkyl)succinate synthase (Ass or Mas). The genes encoding Ass have recently been identified in the alkane degrading sulfidogenic bacteria D. alkenivorasAK-01 [78] and Desulfoglaeba alkanedexensALDCT [71], as well as in nitrate reducing strains HxN1 [65] and OcN1 [79], all affiliated to the Proteobacteria phylum [80]. Recently, Callaghan et al. [71] detected assA genes in a propane-utilizing mixed culture and in a paraffin-degrading enrichment culture maintained under sulfate-reducing conditions. Despite of no genes for benzyl-and alkylsuccinate synthase were found when environmental metagenome datasets of uncontaminated sites were analyzed in Callaghan et al [71], the authors consider that assA gene could be a useful biomarker for anaerobic alkane metabolism.

The second mechanism for alkane anaerobic degradation is the carboxylation, mainly developed from the growth pattern of the sulfate-reducing strain Hxd3 [81], tentatively named as Desulfococcus oleovorans. This strain differs from other alkane degraders for converting C-even alkanes into C-odd cellular fatty acids whereas growth on C-odd alkanes resulted in C-even cellular fatty acids [81, 72]. More recently, Callaghan et al. [82] suggested that a carboxylation-like mechanism analogous to the activation strategy previously proposed by So et al. [81] was the probable route for the anaerobic biodegradation of hexadecane in an alkane-degrading, nitrate-reducing consortium. However, in both cases, the hypothetical fatty acid intermediate (2-ethylalkanoate) that should result from the incorporation of inorganic carbon at C-3 of the alkane has never been detected. There is an on-going debate about this initial activation mechanism. From an energetic point of view, the carboxylation of alkanes is not feasible under physiological conditions, unless the concentration of the fatty acid (2-ethylalkanoate) is in the micromolar order of magnitude or less [80].

Other alternative activation mechanisms are proposed for the anaerobic degradation of alkanes. For instance, the mechanism referred as “unusual oxygenation” is used by the strain Pseudomonas chloritidismutansAW-1T, that is assumed to produce its own oxygen via chlorate respiration used for subsequent metabolism of alkanes [60]. Other alternative mechanism considers that activation in the anaerobic methanogenic system may be initiated by an anaerobic hydroxylation reaction [83].

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6. Mechanisms involved in oil biodegradation in petroleum reservoirs

From those microorganisms studied in oilfields, methanogens have received particular attention since they have been isolated and molecularly detected in both low- and high-temperature reservoirs [88, 89]. Their physiological characteristics and potential activity possibly involved in methanogenesis occurring in oil reservoirs have been demonstrated [90]. Furthermore, recently, Jones et al. [20] provided evidence that the patterns of hydrocarbon degradation observed in biodegraded petroleum reservoirs were the result of methanogenic processes. Therefore, microbiological and biogeochemical investigations have indicated that methanogenesis is a widely distributed process in petroleum reservoirs, although still poorly understood [90]. Methanogenesis is the terminal process of biomass degradation. Acetate and hydrogen are the most important immediate precursors for methanogenesis, and are converted into methane by acetoclastic and hydrogenotrophic methanogens, respectively [91]. Acetate can also be a precursor for methanogenesis through syntrophic acetate oxidation coupled to hydrogenotrophic methanogenesis, which is mediated by syntrophic bacteria and methanogenic archaea [92, 93, 94, 95]. Interestingly, acetate is generally abundant in many petroleum reservoirs, at concentrations ranging between 0.3 and 20 mM [96] hence, acetate metabolism is considered an important methane production process in those environments [90].

Cultivation-dependent and -independent approaches have shown the presence of acetoclastic and hydrogenotrophic methanogens and putative syntrophic acetate-oxidizing bacteria in reservoirs [88, 89, 102], indicating that there should be two different pathways of acetate metabolism in the environment, namely acetoclastic methanogenesis and syntrophic acetate oxidation coupled with hydrogenotrophic methanogenesis. Some previous studies suggested that syntrophic acetate oxidation was most likely to occur in petroleum reservoirs, based on molecular biological analysis [89] and thermodynamic calculations [98]. In Jones et al. [20], the composition of oil in microcosms exhibiting methanogenic oil degradation is compared to patterns observed in biodegraded oils from the Gullfaks field in the North Sea. Analysis of the methanogenic communities from oil-degrading microcosms revealed a strong selection for CO2-reducing methanogens against acetoclastic methanogens, and gas isotope modeling also revealed that to match the d13C of methane and carbon dioxide from biodegraded petroleum reservoirs 75–92% of methanogenesis should be via the CO2 reduction pathway [20, 11].

The reason why syntrophic acetate oxidation predominates over acetoclastic methanogenesis in oil reservoirs remains unclear. There is evidence from studies of oil contaminated aquifers that crude oil can have a detrimental effect on acetoclastic methanogenesis and, in situations where acetoclastic methanogenesis is inhibited, methanogenic alkane degradation via syntrophic acetate oxidation may be thermodynamically the most favorable alternative pathway [11]. Nonetheless, one recent report suggests that acetoclastic methanogenesis may predominate in some methanogenic oil-degrading systems [19]. Although there is currently great interest in how much each of the two pathways contributes to methane production in petroleum reservoirs, no studies are being conducted to address this question [90].

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7. Metagenomics as a tool for a better comprehension of biodegradation

As stated previously, cultivation-based methods have traditionally been utilized for studying the microbiology in oil fields and have yielded valuable information about microbial interactions and their relations with hydrocarbons [42]. However, nowadays, it is known that only a small fraction of the microbial diversity in nature (1-10%) can be grown in the laboratory [84, 85, 86]. Therefore, it is assumed that the ecological functions of the majority of microorganisms in nature and their potential applications in biotechnology remain obscure [87].

In metagenomics, total DNA is extracted from appropriately chosen environmental samples, propagated in the laboratory by cloning techniques, submitted to sequence or function-based screenings and/or subjected to large-scale sequence analysis (Fig. 2). Functional screening of metagenomic libraries offer the advantage that it does not rely on sequence homology to known genes, and for this reason, has allowed the isolation of different enzyme classes from several environments. The probability (hit rate) of identifying a certain gene depends on multiple factors that are intrinsically linked to each other: the host–vector system, size of the target gene, its abundance in the source metagenome, the assay method, and the efficiency of heterologous gene expression in a surrogate host [99].

Figure 2.

Schematic representation of the different steps for metagenomic analysis.

One of the first studies using metagenomics to study microbial degradation of aromatic compounds was performed by Suenaga and colleagues [100], who constructed a metagenomic library from activated sludge for industrial wastewater. The library was functionally screened for extradiol dioxygenase activities (enzymes for aromatic degradation) and 38 clones were subjected to sequencing analysis [101]. As a result, various types of gene subsets were identified that were not similar to the previously reported pathways performing complete degradation. Moreover, the authors discussed the fact that aromatic compounds in the environment may be degraded through the concerted action of various fragmented pathways. Sierra-Garcia [101] reported the organization of hydrocarbon degradation genes of selected metagenomic fosmid clones derived from a metagenomic library from Brazilian petroleum reservoir and functional screening for hydrocarbon degradation activities. The author found many putative proteins of different aerobic and anaerobic well described catabolic pathways, however the complete catabolic pathways described for hydrocarbon degradation in previous studies were absent in the fosmid clones. Instead, the metagenomic fragments comprised genes belonging to different pathways, showing novel gene arrangements where hydrocarbon compounds were degraded through the concerted actions of these fragmented pathways. These results suggest that there are marked differences between the degradation genes found in microbial communities derived from enrichments of oil reservoir sample and those that have been previously identified in bacteria isolated from contaminated or pristine environments.

However, function-based screening of metagenomic libraries for xenobiotic degradation genes is often considered problematic because of insufficient and biased expression of the heterologous genes in the host Escherichia coli[99]. Only a few efforts have been made to solve these problems. In Uchiyama et al. [103], a novel method for function-driven screening is described, which was termed substrate-induced gene expression screening (SIGEX). This high-throughput screening approach employs an operon trap gfp expression vector in combination with fluorescence-activated cell sorting. The screening is based on the fact that catabolic-gene expression is induced mainly by specific substrates and is often controlled by regulatory elements located close to catabolic genes [103]. Using this approach, Uchiyama et al. [103] isolated aromatic-hydrocarbon-induced genes from a metagenomic library derived from groundwater. In Ono et al. [104] another screening strategy was based on functional complementation of a Pseudomonas putidahost strain containing a naphthalene degrading pathway devoid of the naphthalene dioxygenase (NDO) encoding gene. Two clones were able to restore the ability of the host strain to use naphthalene as a sole carbon source and their genes were similar but no identical to already known operons. The authors refer to the use of other host strains for the construction of metagenomic libraries instead of the well-established E. colias a simpler and economical way to perform function-driven screening in comparison to other reported systems such as SIGEX [103].

In the context of this chapter, several aspects of the hydrocarbon degradation need to be studied to obtain a comprehensive overview of the biodegradation processes that take place in oil reservoirs or petroleum impacted environments. These studies should take into consideration the high diversity of enzymes capable of initiating such metabolism as well as the implementation of integrated studies combining culture and molecular techniques, linking with metabolomics or compound-specific isotope analysis and microcosm studies for a better resolution of in situ microbial activity in petroleum reservoirs.

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8. Conclusions and research needs

The understanding about biodegraded petroleum reservoirs have advanced considerably in recent years, but the organisms responsible for the in situactivity and a quantitative understanding of the factors which control in-reservoir oil biodegradation remain far from complete. The inaccessibility of petroleum reservoirs and inherent difficulties of microbiological sampling from commercially operating oil wells have required a multidisciplinary approach to delineating the study of subsurface petroleum biodegradation, and to date there are still prevailing paradigms relating to hydrocarbon biodegradation processes. This multidisciplinary approach to study in situpetroleum degradation should consider molecular biology, microbiology, and geological and geochemical parameters in order to establish the key organisms, biochemical reactions and mechanisms involved in such complex associations. Indeed, the isolation of anaerobic microorganisms capable of utilizing hydrocarbons is essential for a comprehensive understanding of their role and behavior in anoxic habitats and their complex interactions within methanogenic hydrocarbon-degrading communities. In addition, novel approaches, combining functional metagenomics, transcriptomics, metabolomics and other molecular surveys in microcosms are urgently required to better allow access to a more realistic phylogenetic and metabolic diversity governing oil biodegradation in petroleum reservoirs.

References

  1. 1. RolingW2003The microbiology of hydrocarbon degradation in subsurface petroleum reservoirs: perspectives and prospects.Res Microbiol. 154(5), 321-328.
  2. 2. HeadI. MJonesD. MLarterS. R2003Biological activity in the deep subsurface and the origin of heavy oil.Nature, 426(6964), 344-52.
  3. 3. IsmailWandGescherJ2012Epoxy coenzyme a thioester pathways for degradation of aromatic compounds.Appl Environ Microbiol.7815504351
  4. 4. ChandlerD. PLiS. MSpadoniC. MDrakeG. RBalkwillD. LFredricksonJ. KBrockmanF. J1997A molecular comparison of culturable aerobic heterotrophic bacteria and 16S rDNA clones derived from a deep subsurface sediment.FEMS Microbiol Ecol.23131144
  5. 5. LeighM. BPellizariV. HUhlikOSutkaRRodriguesJOstromN. Eet al2007Biphenyl-utilizing bacteria and their functional genes in a pine root zone contaminated with polychlorinated biphenyls (PCBs).ISME J1134148
  6. 6. De LorenzoV2008Systems biology approaches to bioremediation.Curr Opin Biotechnol19579589
  7. 7. JeonCParkWPadmanabhanPDeritoCSnapeJMadsenE2003Discovery of a bacterium, with distinctive dioxygenase, that is responsible forin situbiodegradation in contaminated sediment.Proc Natl Acad Sci USA1001359113596
  8. 8. WitzigRJuncaHHechtH. JPieperD. H2006Assessment of toluene/biphenyl dioxygenase gene diversity in benzene-polluted soils: links between benzene biodegradation and genes similar to those encoding isopropylbenzene dioxygenases.Appl Environ Microbiol7235043514
  9. 9. FoghtJ2010Microbial comminities in oil shales, biodegraded and heavy oil reservoirs, and bitumen deposits. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  10. 10. BirkelandN. K2004The microbial diversity of deep subsurface oil reservoirs.Stud Surface Sci Catal151385403
  11. 11. HeadI. MAitkenC. MGrayN. DSherryAAdamsJ. JJonesD. MRowanA. Ket al2010Hydrocarbon degradation in petroleum reservoirs. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  12. 12. LHaridonSReysenbachA. LGlenatPPrieurDJeanthonC. (1995Hot subterranean biosphere in a continental oil reservoir.Nature377223224
  13. 13. GrassiaG. SMcleanK. MGlenatPBauldJSheehyA. J1996A systematic survey for thermophilic fermentative bacteria and archaea in high temperature petroleum reservoirs.FEMS MicrobiolEcol214758
  14. 14. MagotMOllivierBPatelB. K. C2000Microbiology of petroleum reservoirs.Antonie van Leeuwenhoek.772103116
  15. 15. OrphanV. JTaylorL. THafenbradlDandDelongE. F2000Culture-dependent and culture-independent characterization of microbial assemblages associated with high-temperature petroleum reservoirs.Appl Environ Microbiol.66270011
  16. 16. AitkenC. MJonesD. MLarterS. R2004Anaerobic hydrocarbon biodegradation in deep subsurface oil reservoirs.Nature,43170062914
  17. 17. GrayN. DSherryAHubertCDolfingJHeadI. M2010Methanogenic degradation of petroleum hydrocarbons in subsurface environments remediation, heavy oil formation, and energy recovery.Adv Appl Microbiol.7213761
  18. 18. WiddelFRabusR2001Anaerobic biodegradation of saturated and aromatic hydrocarbons.Curr Opin Biotechnol12259276
  19. 19. GiegL. MDuncanK. ESuflitaJ. M2008Bioenergy production via microbial conversion of residual oil to natural gas. Appl Environ Microbiol7430223029
  20. 20. JonesDHeadIGrayNAdamsJRowanAAitkenCBennettBet al2007Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs.Nature, 451(7175), 176-180.
  21. 21. ZenglerKRichnowH. HRossello-moraRMichaelisWWiddelF1999Methane formation from long chain alkanes by anaerobic microorganisms.Nature401266269
  22. 22. BeederJTorsvikTandLienT1995Thermodesulforhabdus norvegicusgen. nov., sp. nov., a novel thermophilic sulfate-reducing bacterium from oil field water.Arch. Microbiol164331336
  23. 23. ReesG. NGrassiaG. SSheehyA. JDwivediP. PPatelB. K. C1995Desulfacinum infernumgen. nov., sp. nov., a thermophilic sulfate-reducing bacterium from a petroleum reservoir.Int. J. Syst. Bacteriol458589
  24. 24. [24]GrabowskiANercessianOFayolleFBlanchetDJeanthonC2005Microbial diversity in production waters of a low-temperature biodegraded oil reservoir.FEMS microbiology ecology, 54(3), 427-43.
  25. 25. MagotMRavotGCampaignolleXOllivierBPatelB. KFardeauM. LThomasPCroletJ. LGarciaJ. L1997Dethiosulfovibrio peptidovorans gen. nov., sp. nov., a new anaerobic, slightly halophilic, thiosulfate-reducing bacterium from corroding offshore oil wells.Int. J. Syst. Bacteriol.47818824
  26. 26. NilsenR. KTorsvikTLienT1996Desulfotomaculum thermocisternum sp. nov., a sulfate reducer isolated from a hot North Sea oil reservoir.Int. J. Syst. Bacteriol.46397402
  27. 27. RavotGMagotMOllivierBPatelB. K. CAgeronEGrimontP. A. DThomasPGarciaJ. L1997Haloanaerobium congolensesp. nov., an anaerobic, moderately halophilic, thiosulfate- and sulfur-reducing bacterium from an African oil field.FEMS Microbiol. Lett.1478188
  28. 28. Miranda-telloEFardeauM. LFernandezLRamirezFCayolJ. LThomasPGarciaJ. LOllivierB2003Desulfovibrio capillatussp. nov., a novel sulfatereducing bacterium isolated from an oil field separator located in the Gulf of Mexico.Anaerobe997103
  29. 29. NazinaT. NTourovaT. PPoltarausA. BNovikovaE. VGrigoryanA. AIvanovaA. Eet al2001Taxonomic study of aerobic thermophilic bacilli: Descriptions ofGeobacillus subterraneusgen. nov., sp. nov. andGeobacillus uzenensissp. nov. from petroleum reservoirs and transfer ofBacillus stearothermophilus,Bacillus hermocatenulatus, Bacillus thermoleovorans, Bacillus kaustophilus, Bacillus thermoglucosidasiusandBacillus thermodenitrificanstoGeobacillusas the new combinationsG. stearothermophilus, G. thermocatenulatus, G. thermoleovorans, G. kaustophilus, G. thermoglucosidasiusandG. thermodenitrificans.Int. J. Syst. Evol. Microbiol.51433446
  30. 30. MiroshnichenkoM. LHippeHStackebrandtEKostrikinaN. AChernyhN. AJeanthonCNazinaT. NBelyaevS. SBonch-osmolovskayaE. A2001Isolation and characterization ofThermococcus sibiricussp. nov. from a Western Siberia high-temperature oil reservoir.Extremophiles.58591
  31. 31. RavotGMagotMFardeauM. LPatelB. K. CThomasPGarciaJ. LOllivierB1999Fusibacter paucivoransgen. nov., sp. nov., an anaerobic, thiosulfate-reducing bacterium from an oil-producing well.Int. J. Syst. Bacteriol.4911411147
  32. 32. DahleHandBirkelandN. K2006Thermovirga lieniigen. nov., sp. nov., a novel moderately thermophilic, anaerobic, amino-acid-degrading bacterium isolated from a North Sea oil well.Int. J. Syst. Evol. Microbiol.5615391545
  33. 33. NilsenR. KandTorsvikT1996Methanococcus thermolithotrophicusisolated from North sea oil field reservoir water.Appl. Environ. Microbiol.62728731
  34. 34. MagotM2005Indigenous microbial communities in oil fields. In B. Ollivier and M. Magot, (Eds.) Petroleum microbiology.2134ASM, Washington, DC.
  35. 35. SöhngenN. L1913Benzin, Petroleum, Paraffinöl und Paraffin als Kohlenstoff- und Energiequelle für Mikroben.Zentr Bacteriol Parasitenk Abt II37595609
  36. 36. WiddelFandMusatF2010Diversity and common principles in enzymatic activation of hydrocarbons. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  37. 37. BollMandHeiderJ2010Anaerobic Degradation of Hydrocarbons: Mechanisms of C-H-Bond activation in the absence of oxygen. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  38. 38. FoghtJ2008Anaerobic biodegradation of aromatic hydrocarbons: pathways and prospects.J Mol Microbiol Biotechnol.15(2-3): 93-120.
  39. 39. Van BeilenJ. BandFunhoffE. G2007Alkane hydroxylases involved in microbial alkane degradation.Appl Microbiol Biotechnol.7411321
  40. 40. WentzelAEllingsenT. EKotlarH. KZotchevS. BThrone-holstM2007Bacterial metabolism of long-chain n-alkanes.Appl Microbiol Biotechnol.76612091221
  41. 41. RojoF2009Degradation of alkanes by bacteria.Environmental microbiology.
  42. 42. Van HammeJ. DSinghAWardO. P2003Recent advances in petroleum microbiology.Microbiol Mol Biol Rev.674503549
  43. 43. RojoF2010Enzymes for Aerobic Degradation of Alkanes. In K. N. Timmis (Ed.),Handbook of Hydrocarbon and Lipid Microbiology(781Berlin, Heidelberg: Springer Berlin Heidelberg.
  44. 44. MargesinRLabbeDSchinnerFGreerCWhyteL2003Characterization of hydrocarbon-degrading microbial populations in contaminated and pristine alpine soils.Appl Environ Microbiol.69630853092
  45. 45. KuhnEBellicantaG. SPellizariV. H2009New alk genes detected in Antarctic marine sediments.Environ Microbiol.113669673
  46. 46. SalminenJ. MTuomiP. MJorgensenK. S2008Functional gene abundances (nahAc, alkB, xylE) in the assessment of the efficacy of bioremediation.Appl Biochem Biotechnol151638652
  47. 47. HamamuraNFukuiMWardD. MInskeepW. P2008Assessing soil microbial populations responding to crude-oil amendment at different temperatures using phylogenetic, functional gene (alkB) and physiological analyses.Environ Sci Technol4275807586
  48. 48. Van BeilenJ. BWubboltsM. GWitholtB1994Genetics of alkane oxidation byPseudomonas oleovorans.Biodegradation561174
  49. 49. MarchantRSharkeyF. HBanatI. MRahmanT. JPerfumoA2006The degradation of n-hexadecane in soil by thermophilic geobacilli.FEMS Microbiol Ecol.561444
  50. 50. Van BeilenJ. BLiZDuetzW. ASmitsT. H. MWitholtB2003Diversity of Alkane Hydroxylase Systems in the Environment.Oil Gas Sci Technol.584427440
  51. 51. TourovaT. PNazinaT. NMikhailovaE. MRodionovaT. AEkimovA. NMashukovaA. VPoltarausA. B2008alkB homologs in thermophilic bacteria of the genusGeobacillus.Mol Biol.422217226
  52. 52. LiWWangL. YDuanR. YLiuJ. FGuJ. DMuB. Z2012Microbial community characteristics of petroleum reservoir production water amended with n-alkanes and incubated under nitrate-, sulfate-reducing and methanogenic conditions.Inter Biodeterior Biodegradation.698796
  53. 53. Vilchez-vargasRJuncaHPieperD. H2010Metabolic networks, microbial ecology and “omics” technologies: towards understanding in situ biodegradation processes.Environ Microbiol.1230893104
  54. 54. FengLWangWChengJRenYZhaoGGaoCTangYet al2007Genome and proteome of long-chain alkane degrading Geobacillus thermodenitrificans NG80-2 isolated from a deep-subsurface oil reservoir.Proc Natl Acad Sci U S A.1041356027
  55. 55. CarmonaMZamarroMBlazquezBDurante-rodriguezGJuarezJValderramaJet al2009Anaerobic catabolism of aromatic compounds: a genetic and genomic view.Microbiol Mol Biol Rev.7371133
  56. 56. BrennerovaM. VJosefiovaJBrennerVPieperD. HJuncaH2009Metagenomics reveals diversity and abundance of meta-cleavage pathways in microbial communities from soil highly contaminated with jet fuel under air-sparging bioremediation.Environ Microbiol.119221627
  57. 57. Pérez-pantojaDGonzálezBPieperD. H2010Aerobic degradation of aromatic hydrocarbons. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  58. 58. JouanneauY2010Oxidative inactivation of ring cleavage extradiol dioxigenases: mechanism and ferredoxin mediated reactivation. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  59. 59. JuncaHandPieperD. H2003Functional gene diversity analysis in BTEX contaminated soils by means of PCR-SSCP DNA fingerprinting: comparative diversity assessment against bacterial isolates and PCR-DNA clone libraries.Environ Microbiol.6295110
  60. 60. MehboobFJuncaHSchraaGStamsA. J. M2009Growth of Pseudomonas chloritidismutans AW-1(T) on n-alkanes with chlorate as electron acceptor.Appl Microbiol Biotechnol.83473947
  61. 61. FuchsG2008Anaerobic metabolism of aromatic compounds.Ann N Y Acad Sci.11258299
  62. 62. KubeMHeiderJAmannJHufnagelPKühnerSBeckAReinhardtRet al2004Genes involved in the anaerobic degradation of toluene in a denitrifying bacterium, strain EbN1.Arch Microbiol.181318294
  63. 63. BollMFuchsGHeiderJ2002Anaerobic oxidation of aromatic compounds and hydrocarbons.Curr Opin Chem Biol.6560411
  64. 64. Kaserf. MandCoatesJ. D2010Nitrate, Perchlorate and Metal respirers. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  65. 65. GrundmannOBehrendsARabusRAmannJHalderTHeiderJWiddelF2008Genes encoding the candidate enzyme for anaerobic activation of n-alkanes in the denitrifying bacterium, strain HxN1.Environ Microbiol.10237685
  66. 66. KaneS. RBellerH. RLeglerT. CAndersonR. T2002Biochemical and genetic evidence of benzylsuccinate synthase in toluene-degrading, ferric iron-reducingGeobacter metallireducens.Biodegradation,13214954
  67. 67. WinderlCSchaeferSLuedersT2007Detection of anaerobic toluene and hydrocarbon degraders in contaminated aquifers using benzylsuccinate synthase (bssA) genes as a functional marker.Environ Microbiol910351046
  68. 68. WinderlCAnneserBGrieblerCMeckenstockR. ULuedersT2008Depth resolved quantification of anaerobic toluene degraders and aquifer microbial community patterns in distinct redox zones of a tar oil contaminant plume.Appl Environ Microbiol74792801
  69. 69. StaatsMBrasterMRolingW. F. M2011Molecular diversity and distribution of aromatic hydrocarbon-degrading anaerobes across a landfill leachate plume.Environ Microbiol1312161227
  70. 70. WawrikBMendivelsoMParisiV. ASuflitaJ. MDavidovaI. AMarksC. RVan NostrandJ. DLiangYZhouJHuizingaB. Jet al2012Field and laboratory studies on the bioconversion of coal to methane in the San Juan Basin.FEMS Microbiol Ecol.812642
  71. 71. CallaghanA. VDavidovaI. ASavage-ashlockKParisiV. AGiegL. MSuflitaJ. MKukorJ. Jet al2010Diversity of benzyl- and alkylsuccinate synthase genes in hydrocarbon-impacted environments and enrichment cultures.Environ Sci Technol.4419728794
  72. 72. WiddelFandGrundmannO2010Biochemistry of the anaerobic degradation of non-methane alkanes. In: K. N. Timmis (Ed.)Handbook of Hydrocarbon and Lipid Microbiology. Berlin, Heidelberg: Springer Berlin Heidelberg.
  73. 73. GrossiVCravolaureauCGuyoneaudRRanchoupeyruseAHirschlerreaA2008Metabolism of n-alkanes and n-alkenes by anaerobic bacteria: A summary.Org Geochem.39811971203
  74. 74. GiegL. MDavidovaI. ADuncanK. ESuflitaJ. M2010Methanogenesis, sulfate reduction and crude oil biodegradation in hot Alaskan oilfields.Environ Microbiol.1211307486
  75. 75. MbadingaS. MLiK. PZhouLWangL. YYangSZLiuJ. FGuJ.D.,et al2012Analysis of alkane-dependent methanogenic community derived from production water of a high-temperature petroleum reservoir.Appl Microbiol Biotechnol.96253142
  76. 76. GiegL. MandSuflitaJ. M2002Detection of anaerobic metabolites of saturated and aromatic hydrocarbons in petroleum-contaminated aquifers.Environ. Sci. Technol.361737553762
  77. 77. DuncanK. EGiegL. MParisiV. ATannerR. SSuflitaJ. MGreen Tringe, S., Bristow, J. (2009Biocorrosive thermophilic microbial communities in Alaskan North Slope oil facilities.Environ Sci Technol4379777984
  78. 78. CallaghanA. VWawrikBNıChadhain, S.M., Young, L.Y., Zylstra, G.J. (2008Anaerobic alkane-degrading strain AK-01 contains two alkylsuccinate synthase genes.Biochem Biophys Res Commun.366142148
  79. 79. ZedeliusJRabusRGrundmannOWernerIBrodkorbDSchreiberFEhrenreichPBehrendsAWilkesHKubeMReinhardtRWiddelF2010Alkane degradation under anoxic conditions by a nitrate-reducing bacterium with possible involvement of the electron acceptor in substrate activation.Environ Microbiol Rep.31125135
  80. 80. MbadingaS. MWangL. YZhouLLiuJ. FGuJ. DMuB. Z2011Microbial communities involved in anaerobic degradation of alkanes.Inter Biodeterior Biodegradation.651113
  81. 81. SoCPhelpsCYoungL2003Anaerobic transformation of alkanes to fatty acids by a sulfate-reducing bacterium, strain Hxd3.Appl Environ.69738923900
  82. 82. CallaghanA. VTierneyMPhelpsC. DYoungL. Y2009Anaerobic biodegradation of n-hexadecane by a nitrate-reducing consortium.Appl Environ Microbiol7513391344
  83. 83. Head, I., Gray, N., Aitken, C., Sherry, A., Jones, M., Larter, S. (2010). Hydrocarbon activation under sulfate-reducing and methanogenic conditions proceeds by different mechanisms. Geophysical Research Abstracts 12 (EGU General Assembly 2010
  84. 84. TorsvikVGoksoyrJDaaeF. L1990High diversity in DNA of soil bacteria.Appl Environ Microbiol56782787
  85. 85. AmannR. ILudwigWSchleiferK. H1995Phylogenetic identification and in situ detection of individual microbial cells without cultivation.Microbiol Rev59143169
  86. 86. TorsvikVDaaeF. LSandaaR. AØvreåsL1998Novel techniques for analyzing microbial diversity in natural and perturbed environments.J Biotechnol645362
  87. 87. KellenbergerE2001Exploring the unknown: the silent revolution of microbiology.EMBO reports, 2(1), 2-5.
  88. 88. OrphanV. JGoffrediS. KDelongE. FBolesJ. R2003Geochemical influence on diversity and microbial processes in high temperature oil reservoirs.Geomicrobiol J20295311
  89. 89. NazinaT. NShestakovaN. MGrigor’yan, A.A., Mikhailova, E.M., Tourova, T.P., Poltaraus, A.B.,et al. (2006Phylogenetic diversity and activity of anaerobic microorganisms of high-temperature horizons of the Dagang oil field (P.R. China).Microbiology755565
  90. 90. MayumiDMochimaruHYoshiokaHSakataSMaedaHMiyagawaYIkarashiMet al2011Evidence for syntrophic acetate oxidation coupled to hydrogenotrophic methanogenesis in the high-temperature petroleum reservoir of Yabase oil field (Japan).Environ Microbiol.13819952006
  91. 91. GarciaJ. LPatelB. KOllivierB2000Taxonomic, phylogenetic, and ecological diversity of methanogenic Archaea.Anaerobe6205226
  92. 92. ZinderS. HandKochM1984Non-acetoclastic methanogenesis from acetate: acetate oxidation by a thermophilic syntrophic coculture.Arch Microbiol138263272
  93. 93. SchnurerAHouwenF. PSvenssonB. H1994Mesophilic syntrophic acetate oxidation during methane formation by a triculture at high ammonium concentration.Arch Microbiol1627074
  94. 94. HattoriSKamagataYHanadaSShounH2000Thermacetogenium phaeumgen. nov., sp. nov., a strictly anaerobic, thermophilic, syntrophic acetate-oxidizing bacterium.Int J Syst Evol Microbiol5016011609
  95. 95. BalkMWeijmaJStamsA. J2002Thermotoga lettingaesp. nov., a novel thermophilic, methanoldegrading bacterium isolated from a thermophilic anaerobic reactor.Int J Syst Evol Microbiol5213611368
  96. 96. BarthT1991Organic-acids and inorganic-ions in waters from petroleum reservoirs, Norwegian continental-shelf: a multivariate statistical-analysis and comparison with American reservoir formation waters.Appl Geochem6115
  97. 97. SilvaT. RVerdeL. C. LSantos Neto, E.V., Oliveira, V.M. (2012Diversity analyses of microbial communities in petroleum samples from Brazilian oil fields. Inter Biodeterior Biodegradationdoi:10.1016/j.ibiod.2012.05.005.
  98. 98. DolfingJLarterS. RHeadI. M2008Thermodynamic constraints on methanogenic crude oil biodegradation.ISME J2442452
  99. 99. UchiyamaTandMiyazakiK2009Functional metagenomics for enzyme discovery: challenges to efficient screening.Curr Opin Biotechnol.206616622
  100. 100. SuenagaHOhnukiTMiyazakiK2007Functional screening of a metagenomic library for genes involved in microbial degradation of aromatic compounds.Environ Microbiol.9922892297
  101. 101. SuenagaHKoyamaYMiyakoshiMMiyazakiRYanoHSotaMOhtsuboYet al2009Novel organization of aromatic degradation pathway genes in a microbial community as revealed by metagenomic analysis.ISME J.312133548
  102. 102. Sierra-garciaI. NCaracterização estrutural e funcional de genes de degradação de hidrocarbonetos originados de metagenoma microbiano de reservatório de petróleo. M SC. Thesis. Universidade Estadual de Campinas;2011
  103. 103. UchiyamaTAbeTIkemuraTWatanabeK2005Substrate-induced gene-expression screening of environmental metagenome libraries for isolation of catabolic genes.Nat Biotechnol.2318893
  104. 104. OnoAMiyazakiRSotaMOhtsuboYNagataYTsudaM2007Isolation and characterization of naphthalene-catabolic genes and plasmids from oil-contaminated soil by using two cultivation-independent approaches.Appl Microbiol Biotechnol.74250110

Written By

Isabel Natalia Sierra-Garcia and Valéria Maia de Oliveira

Submitted: June 28th, 2012 Published: June 14th, 2013