Abstract
Hydrocarbon-oxidizing bacteria have been isolated from a variety of terrestrial and aquatic environments, using both enrichment and direct plating techniques. Although bacteria able to grow on aliphatic and aromatic hydrocarbons are found in many genera, the genera Alcanivorax appear to be special because these bacteria are specialized for growth on hydrocarbons. The initial step in the bacterial degradation of hydrocarbons is the introduction of oxygen into the molecules by group-specific oxygenases. Since these oxygenases are membrane bound, the cell must come into direct contact with their water-insoluble substrate. Hydrocarbon-oxidizing bacteria have potential applications in bioremediation of oil pollution, enhanced oil recovery, production of surface-active agents, and in the use of hydrocarbons as substrates for industrial fermentation processes.
-
1.
Microbial spoilage of petroleum products
-
2.
Treatment of oil spills and disposal of petroleum wastes
-
3.
Enhanced oil recovery
-
4.
Production of surface-active agents
-
5.
Hydrocarbons as substrates in industrial fermentation processes
Access provided by Autonomous University of Puebla. Download reference work entry PDF
Similar content being viewed by others
Keywords
- Enrichment Culture
- Mineral Salt Medium
- Minimal Salt Medium
- Hydrocarbon Degradation
- Hydrocarbon Substrate
These keywords were added by machine and not by the authors. This process is experimental and the keywords may be updated as the learning algorithm improves.
Introduction
Periodic ecological disasters caused by large oil spills call attention, in a dramatic manner, to the toxicity of petroleum. The fact that hydrocarbons persist for months and even years following major oil spills indicates that hydrocarbon biodegradation is slow in most natural environments. To the microbiologist, the fundamental questions are the following: What are the biochemical mechanisms of hydrocarbon degradation? Which microorganisms are involved? What are their special properties? What limits the rate of hydrocarbon degradation in the environment? And from an applied point of view, what (if anything) can be done to accelerate this rate? Several decades of research on hydrocarbon-oxidizing bacteria have provided considerable data relevant to these questions. This chapter will discuss the distribution, nutritional requirements, enumeration, isolation, identification, special physiologic characteristics, and potential applications of hydrocarbon-degrading bacteria. The specific class of methane oxidizers will be presented in a separate chapter.
Habitats
Hydrocarbons are a ubiquitous class of natural compounds. Not only are they found in petroleum-polluted areas, but chemical analyses have revealed the presence of significant quantities of aliphatic and aromatic hydrocarbons in most soils and sediments (Giger and Blumer 1974; Stevenson 1966). The most probable origin of the low concentrations of widely distributed hydrocarbons is ongoing biosynthesis by certain plants and microorganism (Fehler and Light 1970; Hardwood and Russel 1984; Hunt et al. 1980; Juttner 1976; Kolattukudy et al. 1972; Mikkelson and von Wettstein-Knowles 1978; Winters et al. 1969). Hydrocarbons are produced by reduction of fatty acyl-CoA by enzymes which utilize NADH or NADPH. Other sources of hydrocarbons are natural seeps on the ocean floor and unburned fuel from oil-burning engines (Floodgate 1984). Since hydrocarbons are natural products as well as pollutants, it is not surprising that hydrocarbon-oxidizing bacteria are widely distributed in nature. A sample of ecological studies on hydrocarbon-degrading bacteria is shown in Table. 5.1 .
It can be seen that hydrocarbon oxidizers are located in virtually all natural areas, although with large variations in cell concentration. As would be expected, the ratio of hydrocarbon-oxidizing bacteria to the total population of heterotrophic bacteria, as well as the variety of hydrocarbon-degrading microorganisms found in a particular ecosystem, may change according to the time of sampling or the extent of oil pollution (Geiselbrecht et al. 1996). Atlas (1981) has discussed many of the factors that limit the growth of hydrocarbon-oxidizing bacteria in nature. These include physical constraints, such as temperature, availability of oxygen, salinity, pH, and the extent to which the particular habitat is an open or closed ecosystem. Nutritional factors are also important and include the availability of utilizable sources of nitrogen, phosphorus, and other elements; the nature of the hydrocarbon substrate and its effective concentration; and the possible presence of toxic substances either in the petroleum product or in the environment itself.
Effect of Oil Pollution
The localization of hydrocarbon-oxidizing bacteria in natural environments has received considerable attention because of the possibility of utilizing their biodegradation potential in the treatment of oil spills. Because of the enormous quantities of crude and refined oils that are transported over long distances and consumed in large amounts, the hydrocarbons have now become a very important class of potential substrates for microbial oxidation. It is not surprising, therefore, that hydrocarbon-oxidizing microorganisms have recently been isolated in large numbers from a wide variety of natural aquatic and terrestrial environments. Several investigators have demonstrated an increase in the number of hydrocarbon-oxidizing bacteria in areas that suffer from oil pollution (Table. 5.1 ). Walker and Colwell (1976a, b) observed a positive correlation between the percentage of petroleum-degrading bacteria in the total population of heterotrophic microorganisms and the amount of heptane-extractable material in sediments of Colgate Creek, a polluted area of Chesapeake Bay. In contrast, no correlation was found when total numbers of hydrocarbon oxidizers (rather than percentages) were compared to hydrocarbon levels. Horowitz and Atlas (1977b) observed shifts in microbial populations in an Arctic freshwater lake after the accidental spillage of 55,000 gal of leaded gasoline. The ratio of hydrocarbon-utilizing to total heterotrophic bacteria was reported to be an indicator of the gasoline contamination. These investigators also studied shifts in microbial populations in Arctic coastal water using a continuous flow-through system, following the introduction of an artificial oil slick (Horowitz and Atlas 1977a). The addition of the oil appeared to cause a shift to a greater percentage of petroleum-degrading bacteria. Atlas and Bartha (1973b) found similar results in an oil-polluted area in Raritan Bay off the coast of New Jersey. Hood et al. (1975) compared microbial populations in sediments of a pristine salt marsh with those of an oil-rich marsh in southeastern Louisiana. These investigators also found a high correlation between the percentage of hydrocarbon oxidizers and the level of hydrocarbons in the sediments. Significant increases in the number of hydrocarbon-utilizing microorganisms were found in field soils following the addition of several different oil samples (Raymond et al. 1976). No estimate of the ratio of hydrocarbon oxidizers to the total heterotrophic population was presented.
From the studies discussed above, it is clear that the presence of hydrocarbons in the environment frequently brings about a selective enrichment in situ for hydrocarbon-utilizing microorganisms. Evidence also has been presented suggesting that the supplementation of certain ecosystems, particularly oil-polluted marine environments with nitrogen and phosphorus, may increase the relative number of hydrocarbon oxidizers (Atlas and Bartha 1973a; Gutnick and Rosenberg 1977; Reisfeld et al. 1972; Song and Bartha 1990).
Isolation and Enumeration
The use of hydrocarbons as substrates for bacterial growth presents special problems both to the microorganism using them as a source of carbon and energy and to the investigators in the field of hydrocarbon microbiology. Depending on the solubility of the particular hydrocarbon in water, its physical state (solid, liquid, or gas), and toxicity, different isolation methods must be employed. In all cases, the heterogeneity of the system complicates sampling, enumeration, and growth measurement procedures. After a discussion of general nutritional requirements for hydrocarbon-degrading bacteria, several specific procedures for the selective enrichment and isolation of the different hydrocarbon degraders will be presented.
General Nutritional Requirements
In addition to the requirements for suitable cell-hydrocarbon interactions and the specific genetic potential of the organism for hydrocarbon oxidation, a number of general nutritional conditions must be fulfilled for bacteria to utilize hydrocarbons. These nutritional requirements depend on the fact that hydrocarbons, as the name denotes, are compounds composed solely of carbon and hydrogen atoms. Thus, all other elements essential for cell growth must be available in the growth medium. These include molecular oxygen, utilizable forms of nitrogen, phosphorus, sulfur, metals, and trace components. The requirement for molecular oxygen has been given much attention, particularly with respect to maximum production of single-cell protein by hydrocarbon-degrading microorganisms (Mimura et al. 1973; Schocken and Gibson 1984). The limitation for oxygen is easily overcome in small-scale laboratory studies or in open aqueous systems where the oil–water interface is in direct contact with air at all times. The possibility of anaerobic decomposition of hydrocarbons has received considerable attention (Hollinger and Zehnder 1996). Although hydrocarbon utilization by strictly anaerobic sulfate-reducing bacteria (e.g., Rosenfeld 1947) has been reported, evidence that pure cultures of sulfate-reducing bacteria can attack hydrocarbon in the absence of additional sources of organic carbon is not definite. However, a few microbial species appear to be able to grow on pure alkane in the absence of molecular oxygen, if provided with nitrate as an electron acceptor (Senez and Azoulay 1961; Mihelcic and Luthy 1988) or sulfate (Rueter et al. 1994; Rabus et al. 1999).
The nitrogen and phosphorus requirements for maximum growth of hydrocarbon oxidizers can generally be satisfied by ammonium phosphate. Alternatively, these requirements can be met with a mixture of other salts, such as ammonium sulfate, ammonium nitrate, ammonium chloride, potassium phosphate, sodium phosphate, and calcium phosphate. When ammonium salts of strong acids are used, the pH of the medium generally decreases with growth. This problem can often be overcome by using urea as the nitrogen source. In theory, approximately 150 mg of nitrogen and 30 mg of phosphorus are consumed in the conversion of 1 g of hydrocarbon to cell material. In open systems, the high water solubility of most utilizable sources of nitrogen and phosphorus reduces their effectiveness because of rapid dilution. In principle, this problem can be solved by using oleophilic nitrogen and phosphorus compounds with low C/N and C/P ratios. It was found that a combination of paraffinized urea and octyl phosphate was able to replace nitrate and inorganic phosphate, respectively (Atlas and Bartha 1973a). A more economical way may be to add water-insoluble controlled-release nitrogen and phosphorus fertilizers. This technology has been successfully demonstrated in laboratory and field experiments (Rosenberg et al. 1996). Another practical source of hydrophobic N and P is guano. One intriguing possibility to obviate the need for addition of nitrogen compounds to the medium is to use a bacterium that is capable of both hydrocarbon degradation and nitrogen fixation. Such microorganisms were reported following enrichment on hydrocarbon media lacking nitrogen salts (Coty 1967).
In addition to utilizable sources of nitrogen and phosphorus, the mineral requirements of hydrocarbon-degrading bacteria can be met by the addition of K+, Mg2+, Fe2+, and SO 2−4 to purified media. All other inorganic ions required by bacteria to obtain optimum growth are commonly present in sufficient concentration as contaminants in these salts. For most marine hydrocarbon degraders, artificial seawater (or filtered seawater), supplemented simply with phosphate, a nitrogen source, and the hydrocarbon, serves as an adequate medium for enrichment culture studies. In certain aquatic environments under conditions in which the water was supplemented with nitrogen and phosphorus, a high concentration of iron may limit oil biodegradation (Dibble and Bartha 1976). Under these conditions, an encapsulated oleophilic iron compound, ferric octoate, was found to be as effective in stimulating biodegradation as various water-soluble iron derivatives, such as ferric ammonium citrate.
Enumeration of Hydrocarbon-Degrading Bacteria
The determination of the concentration of hydrocarbon-degrading bacteria is one of the methods commonly used for monitoring oil pollution in the environment. Theoretical difficulties associated with the interpretation of these data have been discussed elsewhere (Floodgate 1973). The enumeration of hydrocarbon-degrading bacteria presents two special technical problems, sampling and choice of carbon source. Petroleum-degrading bacteria tend to adhere to hydrophobic materials (Fig. 5.1 ). Thus, unless the bacteria are removed from the material and dispersed prior to enumeration, only minimum cell numbers can be obtained. The choice of a carbon source is an even more serious problem. Petroleum is an extremely complex mixture of hydrocarbons. Because certain bacteria may grow only on minor components in oil, it would be necessary to incorporate large quantities of petroleum into the growth medium to ensure sufficient substrate for these bacteria to grow well. However, high concentrations of petroleum and mixtures of hydrocarbons cannot be used because they are toxic to bacteria (Vestal et al. 1984). Thus, the enumeration of hydrocarbon-degrading bacteria using petroleum as the carbon source selects primarily for bacteria that can degrade major components of the oil mixture. Often, pure hydrocarbons and mixtures of pure hydrocarbons and fractions of crude oil can be used to advantage in replacing petroleum as the carbon source in the isolation medium. The following four methods have been used to enumerate hydrocarbon-degrading bacteria in the marine, estuarine, and freshwater environments.
Enumeration of Hydrocarbon-Degrading Bacteria in Marine Material Not Miscible with Water
-
1.
Approximately 2 g of the material to be examined is placed in a sterile bottle containing 100 ml sterile seawater or salts medium (Gunkel and Trekel 1967).
-
2.
After 1 ml of a sterile, nontoxic, nonionic emulsifier and 1 drop of an antifoam agent are added to the sample, the mixture is homogenized to disperse and break up the bacterial aggregates (e.g., an Ultra Turrax homogenizer run at 24,000 rpm for 30 s).
-
3.
The homogenized sample is then diluted serially in steps of 1:10 in sterile seawater or salts medium.
-
4.
One-milliliter samples of the appropriate dilutions are then inoculated into bottles or tubes containing the following sterile medium:
-
5.
After addition of one drop of sterile hydrocarbon, the samples are incubated for 2–6 weeks, depending on the temperature of incubation.
-
6.
Bottles remaining turbid after addition of 1 ml HCl to dissolve inorganic salts are scored, and the most probable number is calculated from tables published in Standard Methods (American Public Health Association 1995).
Aged seawater | 750 g |
Distilled water | 250 ml |
NH4Cl | 0.5 g |
K2HPO4 | 0.5 g |
NaH2PO4 | 1.0 g |
Enumeration of Hydrocarbon-Utilizing Bacteria by Direct Plating of Estuarine Water and Sediment Samples
Estuarine salts solution (Colwell et al. 1973):
Distilled water | 1 l |
NaCl | 10 g |
MgCl2 | 2.3 g |
KCl | 0.3 g |
Oil powder: 10 g of hydrocarbon dissolved in 30 ml of diethyl ether is mixed with 10 g of silica gel, allowed to evaporate, and then added to the following basal medium prior to autoclaving.
Oil agar medium:
Distilled water | 1 l |
NaCl | 10 g |
MgSO4 | 0.5 g |
NH4NO3 | 1.0 g |
FeCl3 (25 g · ml−1) | 1 drop |
Purified agar (Difco) | 20 g |
Oil powder | 10 g |
KH2PO4 (10 g · 100 ml−1)a | 3.0 ml |
K2HPO4 (10 g · 100 ml−1)a | 7.0 ml |
Fungizonea | 10 mg |
Enumeration of Hydrocarbon-Degrading Bacteria in Freshwater
Basal medium (Ward and Brock 1976):
Distilled water | 1 l |
NaCl | 0.4 g |
NH4Cl | 0.5 g |
MgSO4 · 7H2O | 0.5 g |
NaHPO4 · 7H2Oa | 0.05 g |
KH2PO a4 | 0.05 g |
Serial dilutions are made in the basal medium. One drop of sterile hydrocarbon is added to 10 ml of basal medium. After incubation at the appropriate temperature, growth is detected by pellicle formation at the surface of the oil droplet. Most probable number is determined from the tables published in Standard Methods (American Public Health Association 1995).
Enumeration of Hydrocarbon-Degrading Bacteria by a 96-Well Plate Procedure
The 96-well plates were processed with a Beckman Biomek 1000 laboratory robot (Beckman Instruments, Fullerton, CA, USA) which filled the wells with medium, performed tenfold serial dilutions of the sample, and added oil to the inoculated wells (Haines et al. 1996). The robot added 180 ml of BH (Bushnell-Haas medium, Difco) to each well in 11 of the 12 rows, leaving the first row empty. It transferred 200 ml of undiluted sample to the wells in the first row, mixed their contents, and then transferred 20 ml to each well in the second row. The contents of the second row were mixed, and 20 ml was transferred to each well in the third row. This procedure of mixing and transfer was carried out for all except the last row, which served as a sterile control. Sterile pipet tips were used for each transfer. After the dilutions were completed, 2 ml of oil was added to each well as the growth substrate. The plates were sealed in plastic bags and incubated for 14 days at 20 °C. Positive wells were scored in one of two ways. When F2 (number 2 fuel oil) was the carbon source, 50 ml of a sterile solution (3 g · l−1) of INT (iodonitrotetrazolium violet; Research Organics, Cleveland, OH, USA) was added to each well. INT competes with O2 for electrons from the respiratory electron transport chain, and it is reduced to an insoluble formazan that deposits as a red precipitate in the presence of active respiring microorganisms. Red or pink wells were scored as positive. When a crude oil was used as the carbon source, a smooth oil slick developed in each well. Positive wells were scored by emulsification or dispersion of this oil slick. INT cannot be used effectively with crude-oil substrates because their dark color interferes with detection of formazan deposition.
Enrichment Culture for Hydrocarbon-Degrading Bacteria
Since hydrocarbons are natural products that are widely distributed in nature, it is not surprising that bacteria able to degrade hydrocarbons can easily be isolated by standard enrichment culture procedures. By varying parameters, such as temperature, pH, hydrocarbon concentration, and basal medium, a wide variety of different hydrocarbon-degrading and emulsifying bacteria can be obtained from either aquatic or terrestrial environments. In most studies, crude oi1 or a petroleum distillate was used as the sole carbon and energy source in the enrichment culture procedure. Under those conditions, bacteria that specialize in the oxidation of low-molecular-weight n-alkanes are generally obtained. Bacteria that grow more slowly or oxidize minor components of crude oil never increase much in batch enrichments, although the activity of these microorganisms may be of special significance in natural environments. To overcome this difficulty, enrichment culture procedures have to be employed using different carbon sources. The following examples represent only a few of the possible variations.
Enrichment of Crude Oil-Degrading Bacteria in Supplemented Seawater
To 20 ml of unsterilized seawater in a 125-ml flask was added 155 mg unsterilized crude oil, 0.056 mM KH2PO4, and 7.6 mM (NH4)2SO4 (Reisfeld et al. 1972). After inoculation with about 1 g beach tar or oily sand, the flask was incubated at 30 °C with shaking. After about 1 week, the oil became evenly dispersed throughout the liquid. One milliliter of this culture was then transferred to 20 ml sterile seawater supplemented with 0.056 mM KH2PO4, 7.6 mM (NH4)2SO4, and 1 drop of sterile crude oil. (The crude oil was sterilized by filtration through a Millipore 0.45 mm membrane filter.) After one passage, the oil became emulsified in 2–4 days. Such mixed cultures were maintained by serial transfers to fresh media at 3–4-day intervals. Pure cultures were obtained by streaking the enrichment culture either onto the above medium solidified with 1.5 % agar (Difco) or nutrient agar (Difco) prepared with filtered seawater. Isolated colony types were found to grow both on nutrient- and oil-containing media.
Enrichment culture procedures used to isolate crude oil-degrading bacteria, such as that described in the preceding paragraph, yield a mixture of several different strains even after several transfers. One reason for this is the heterogeneity of the carbon source. Low-molecular-weight paraffin oxidizers (C10 to C25) generally dominate the cultures because of their more rapid growth rate. To isolate bacteria that could utilize other fractions of crude oil, the following sequential enrichment culture can be employed:
Sequential Enrichment of Hydrocarbon-Degrading Bacteria on Crude Oil in Supplemented Seawater
Inoculate the following sterile medium with a pure culture of an n-paraffin-oxidizing bacterium (which can be obtained by standard enrichment culture procedures): 1 l filtered water, 10 mg K2HPO4 · 3H2O, 450 mg urea, and 0.7 ml crude oil (Horowitz et al. 1975). After 3 days incubation with shaking at 30 °C, the residual oil is extracted with 1 l of benzene-pentane-ether (3:1:1, v/v/v). The oil remaining after evaporation of the organic solvent in vacuo is referred to as “bacteria-depleted oil.” An enrichment culture is now carried out using the “bacteria-depleted oil” in place of crude oil as the sole source of carbon and energy.
The general failure of investigators to isolate microorganism on highly water-insoluble, solid hydrocarbons, such as anthracene, may be due to the fact that the cells remain firmly bound to the substrate. Thus, a standard enrichment culture procedure in which a portion of the bulk water phase is transferred would select against rather than for these specific microorganisms. It may be that for successful enrichments, the solid phase should be used as the inoculum during the sequential transfers.
Enrichment of Hydrocarbon-Degrading Bacteria on Bunker C Fuel Oil in Minimal Salts Medium
One gram of beach sand sample or 1 ml of a water sample was added to the following minimal medium containing 0.125 % Bunker C oil (steam-sterilized at 121 °C and 15 psi for 15 min in tightly capped flasks to prevent evaporation) (Mulkins-Phillips and Stewart 1974a):
Minimal salts medium:
Distilled water | 1 l |
NaCl | 28.4 g |
K2HPO4 | 4.74 g |
KH2PO4 | 0.56 g |
MgSO4 | 0.50 g |
CaCl2 | 0.1 g |
NH4NO3 | 2.5 g |
Trace element stock (pH 7.1) | 1 ml |
Flasks were incubated at 20 °C for 14 days and 120 rpm on a refrigerated gyratory shaker bath. Pure cultures of hydrocarbon-utilizing bacteria were isolated from the enrichment culture by streaking onto minimal salts medium to which 2 % washed Ionagar no. 2 (Oxoid) was added. The carbon source consisted of 0.5 ml of the following hydrocarbon mixture added to sterile filter paper secured in the lids of the petri dishes. The dishes were then inverted and incubated at the appropriate temperature for 1–3 weeks.
Hydrocarbon mixture:
Naphthalene | 0.1 g |
Anthracene | 0.1 g |
Dibenzothiophene | 0.1 g |
Decalin | 5 ml |
Hexadecene-1 | 5 ml |
Hexadecane | 5 ml |
Octadecane | 0.1 g |
Dodecane | 5 ml |
Isooctane | 5 ml |
Enrichment of Polyaromatic Hydrocarbon-Degrading Bacteria (PAHs)
Small amounts of fresh sediment known to be contaminated with PAHs were inoculated into the following mineral salts medium (g/l) (Churchill 1999):
(NH4)2SO4 | 10 |
KH2PO4 | 5.0 |
MgSO4 · 7H2O | 0.1 |
Fe(NH)2(SO4)2 | 0.005 |
Pyrene | 40 |
Trace metals (Beauchop and Elsden 1960)
After adjusting the pH to 7.0 with NaOH, the flasks were shaken for 1 week. Pyrene-degrading bacteria were detected on pyrene-coated mineral medium (as above) agar plates. Zones of clearing around colonies indicated pyrene degradation. The same procedure can be used with other PAHs replacing the pyrene.
Enrichment on Liquid Aromatic Hydrocarbons
Liquid aromatic hydrocarbons, such as benzene, toluene, and ethylbenzene, are toxic to bacteria when present in the liquid phase (Gibson 1971). However, if these carbon sources are introduced in the vapor phase, good growth can be obtained. Figure. 5.2 illustrates two methods that can be used for growing bacteria on volatile toxic hydrocarbons. Since the liquid hydrocarbons do not come in direct contact with the salts medium, they need not be sterilized. When the reservoir of volatile hydrocarbons is exhausted, it can easily be refilled with a Pasteur pipette.
Enrichment for Nitrogen-Fixing Hydrocarbon Oxidizers
Hydrocarbon-oxidizing bacteria able to grow in the absence of added nitrogen compounds were isolated by addition of 0.1 g soil to 25 ml of mineral salts medium of the following composition (g/l) (Coty 1967):
Na2HPO4 | 0.3 |
KH2PO4 | 0.2 |
MgSO4 · 7H2O | 0.1 |
FeSO4 · 7H2O | 0.005 |
Na2MoO4 · 2H2O | 0.002 |
The containers were incubated in an atmosphere of air and hydrocarbon vapors. After turbidity developed, the cultures were streaked and reincubated on the above mineral salts medium containing 1.5 % washed agar. Purification was achieved after several restreakings and culturing on nitrogen-free mineral salts agar medium. Bacteria able to utilize atmospheric nitrogen on addition of naphthenic acid, n-butane, n-tetradecane, or sodium cyclohexane carboxylate were reported to be isolated by this procedure.
Enrichment for Solid Hydrocarbon-Degrading Bacteria
A slurry of soil in 0.1 M phosphate buffer, pH 7.0, with 0.1 % octadecane was incubated for 1 week, with shaking (Miller and Bartha 1989). This enrichment culture was transferred (1:100 ratio) to the following medium (g/l):
Na2HPO4 | 0.4 |
KH2PO4 | 0.15 |
NH4Cl | 0.1 |
MgSO4 · 7H2O | 0.02 |
Iron ammonium citrate | 0.005 |
CaCl2 | 0.001 |
Octadecane | 0.1 |
To obtain pure cultures, the enrichment was streaked on the above medium solidified with 2 % agar. The Pseudomonas sp. that was isolated grew on solid alkanes such as hexatriacontane (C36).
Identification
The variation in bacterial populations isolated by enrichment culture depends largely on the hydrocarbon substrate used in the enrichment, the culture conditions, and the source of the inoculum. Many species capable of hydrocarbon degradation have been isolated (Table. 5.2 ). The most frequently isolated bacterial genera are Pseudomonas, Acinetobacter, Flavobacterium, Corynebacterium, Alcanivorax, and Arthrobacter. Most of the investigations on the degradation of aromatic hydrocarbons have been carried out using Pseudomonas putida and species of Beijerinckia and Nocardia (Gibson 1971). Westlake et al. (1974) studied the effect of oil quality and incubation temperature on the genetic composition of hydrocarbon-decomposing populations isolated from an area in British Columbia that had been exposed to chronic pollution with diesel fuel. All of the populations consisted predominantly of Gram-negative rods, including species of Pseudomonas, Acinetobacter, Xanthomonas, Arthrobacter, and Alcaligenes.
An extensive study of petroleum-degrading bacteria isolated from Chesapeake Bay waters and sediments was carried out by Austin et al. (1977a, b). A total of 99 strains were examined for 48 biochemical, cultural, morphological, and physiological characteristics. A statistical analysis revealed 14 phonetic groups, comprising about 85 % of the hydrocarbon-degrading bacteria. These groups were characterized as actinomycetes; coryneforms; Enterobacteriaceae; Klebsiella aerogenes; species of Micrococcus, Nocardia, and Pseudomonas; and Sphaerotilus natans.
Special mention should be made of the genus Alcanivorax (Cappello and Yakimov 2010) because these bacteria seem to play a particularly pivotal role in the oil-spill bioremediation (Schneiker et al. 2006). This obligate hydrocarbonoclastic bacterium (petroleum hydrocarbons serve almost exclusively as its source of carbon and energy) is cosmopolitan and found in various marine environments.
Physiological Properties
There are two essential characteristics that define hydrocarbon-oxidizing bacteria: (1) hydrocarbon-group-specific oxygenases and (2) mechanisms for optimizing contact between the bacterium and the hydrocarbon.
Group-Specific Oxygenases
Several reviews have appeared on the microbial metabolism of straight-chain and branched alkanes (Asperger and Kleber 1991; Singer and Finnerty 1984), cyclic alkanes (Perry 1984), and aromatic hydrocarbons (Gibson 1977; Cerniglia 1984; Pérez-Pantoja et al. 2010). It has been established that the first step in the degradation of hydrocarbons by bacteria is the introduction of both atoms of molecular oxygen into the hydrocarbon. In the case of aromatic hydrocarbons, ring fission requires a dihydroxylation reaction, the introduction of two atoms of oxygen, and the subsequent formation of a cis-dihydrodiol (Gibson 1968; Simon et al. 1993). This reaction is catalyzed by a dioxygenase which is a multicomponent, membrane-bound enzyme system (Cerniglia 1992). Further oxidation of the cis-dihydrodiol leads to the formation of catechols that are substrates for another dioxygenase that catalyzes ring fission (Evans et al. 1965).
It is important to emphasize that the biochemical mechanism of aromatic hydrocarbon oxidation in prokaryotes is fundamentally different from that of eukaryotes. Fungi and mammalian cells metabolize aromatics using the cytochrome P-450 monooxygenase system, which leads to the formation of arene oxides. These active epoxides can form covalent bonds with nucleophilic sites in DNA, leading to mutations and carcinogenesis. Aromatic hydrocarbons that have been shown to serve as substrates for bacterial oxygenases include benzene, toluene, xylene, naphthalene, phenanthrene, anthracene, benz(a)anthracene, biphenyl, and several of their methylated derivatives. The enzymes necessary for aromatic hydrocarbon degradation are specified, in part, by degradative catabolic plasmids. Enzymes capable of monooxygenating benzene/toluene to phenol/methylphenol and phenols to catechols belong to an evolutionary related family of soluble di-iron monooxygenases (Leahy et al. 2003), which are enzyme complexes consisting of an electron transport system comprising a reductase (and in some cases a ferredoxin), a catalytic effector protein which contains neither organic cofactors nor metal ions and is assumed to play a role in assembly of an active oxygenase (Powlowski et al. 1997), and a terminal hydroxylase with a (αβγ)2 quaternary structure and a di-iron center contained in each α subunit. These monooxygenases are classified according to their α-subunits, which are assumed to be the site of substrate hydroxylation, into four different phylogenetic groups: the soluble methane monooxygenases, the alkene monooxygenase of Rhodococcus corallinus B-276, the phenol hydroxylases, and the four-component alkene/aromatic monooxygenases (Leahy et al. 2003).
In general, alkanes are terminally oxidized to the corresponding alcohol, aldehyde, and fatty acid (Asperger and Kleber 1991). The hydroperoxides may serve as unstable intermediates in the formation of the alcohol (Singer and Finnerty 1984). Fatty acids derived from alkanes are then further oxidized to acetate and propionate (odd-chain alkanes) by inducible β-oxidation systems. The group specificity of the alkane oxygenase system is different in various bacterial species. For example, Pseudomonas putida PpG6 (oct) grows on alkanes of 6–10 carbons in chain length (Nieder and Shapiro 1975), whereas Acinetobacter sp. HOI-N is capable of growth on long-chain alkanes (Singer and Finnerty 1984). The ability of P. putida to grow on C6–C10 alkanes was shown to be plasmid encoded (Chakrabarty et al. 1973). In contrast, all activities necessary for growth of Acinetobacter sp. HOI-N and A. calcoaceticus BD413 appear to be coded by chromosomal genes (Singer and Finnerty 1984).
Subterminal alkane oxidation apparently occurs in some bacterial species (Markovetz 1971). This type of oxidation is probably responsible for the formation of long-chain secondary alcohols and ketones. Pirnik (1977) and Perry (1984) have reviewed the microbial oxidation of branched and cyclic alkanes, respectively.
Physical Interactions Between Bacteria and Hydrocarbons: Adhesion, Desorption, and Emulsification
The low solubility of hydrocarbons in water, coupled to the fact that the first step in hydrocarbon degradation involves a membrane-bound oxygenase, makes it essential for bacteria to come into direct contact with their hydrocarbon substrates. Two general biological strategies have been suggested for enhancing contact between bacteria and water-insoluble hydrocarbons: (1) specific adhesion mechanisms and (2) emulsification of the hydrocarbon.
To understand the special cell-surface properties of bacteria that allow them to grow on hydrocarbons, it is necessary to consider the dynamics of petroleum degradation in natural environments (Rosenberg et al. 1992). Following an oil spill in the sea, the hydrocarbons rise to the surface and come into contact with air. Some of the low-molecular-weight hydrocarbons volatilize; the remainder are metabolized relatively rapidly by microorganisms, such as Pseudomonas sp., which take up soluble hydrocarbons. These bacteria do not adhere to oil and do not have a high cell-surface hydrophobicity (Rosenberg and Rosenberg 1985). The next stage of degradation involves microorganisms with high cell-surface hydrophobicity, which can adhere to the residual high-molecular-weight hydrocarbons. In the case of A. calcoaceticus RAG-1, this adherence is due to thin hydrophobic fimbriae (Rosenberg et al. 1982). Mutants lacking these fimbriae failed to adhere to hydrocarbons and were unable to grow on hexadecane. Other bacteria exhibit high cell-surface hydrophobicity as a result of a variety of fimbriae and fibrils, outer-membrane and other surface proteins and lipids, and certain small cell-surface molecules, such as gramicidin S (Rosenberg et al. 1985) and prodigiosin (Rosenberg et al. 1989). Bacterial capsules and other anionic exopolysaccharides appear to inhibit adhesion to hydrocarbons (Rosenberg et al. 1983).
Desorption from the hydrocarbon is a critical part of the growth cycle of petroleum-degrading bacteria. Petroleum is a mixture of thousands of different hydrocarbon molecules. Any particular bacterium is only able to use a part of the petroleum. As the bacteria multiply at the hydrocarbon/water interface of a droplet, the relative amount of nonutilizable hydrocarbon within the droplet continually increases until the cells can no longer grow. For bacteria to continue to multiply, they must be able to move from the depleted droplet to a fresh oil droplet. A. calcoaceticus RAG-1 has an interesting mechanism for desorption and for ensuring that it only reattaches to a droplet of fresh oil. When cells become starved on the “used” hydrocarbon drop or tar ball, they release their capsule. The capsule is composed of an anionic heteropolysaccharide, with fatty acid side chains, referred to as emulsan (Rosenberg 1986). The extracellular, amphipathic emulsan attaches avidly to the hydrocarbon/water interface, thereby displacing the cells to the aqueous phase. Each “used” oil droplet or tar ball is then covered with a monomolecular film of emulsan. The hydrophilic outer surface of the emulsan-coated hydrocarbon prevents reattachment of the RAG-1 cells. The released capsule-deficient bacteria are hydrophobic and readily adhere to fresh hydrocarbon substrate.
Many hydrocarbon-degrading microorganisms produce extracellular emulsifying agents (Desai and Banat 1997; Rosenberg and Ron 1997). In some cases, emulsifier production is induced by growth on hydrocarbons (Hisatsuka et al. 1971). Mutants that do not produce the emulsifier grow poorly on hydrocarbons (Itoh and Suzuki 1972). Pretreatment of oil with emulsifying agents can both inhibit and stimulate oil biodegradation (e.g., Foght et al. 1989; Nakahara et al. 1981; Tiehm 1994; Thibault et al. 1996; Liu et al. 1995; Zhang and Miller 1994). As discussed above, emulsification may be a by-product of a cell/hydrocarbon detachment process. An entire chapter of this book is devoted to bioemulsifiers (Rosenberg and Ron 1997).
Acinetobacter sp. HOI-N accumulates extracellular membrane vesicles of 20–50 nm in diameter when grown on hexadecane (Kappeli and Finnerty 1980). The isolated vesicles partition exogenously supplied hydrocarbons in the form of a microemulsion. These vesicles appear to play a role in the uptake of alkanes. Miller and Bartha (1989) have been able to overcome the difficulties involved in the transport of water-insoluble, solid hydrocarbons by using unilamellar vesicles. A Pseudomonas isolate grew on octadecane (C18) and hexatriacontane (C36) with Ks values of 2,450 and 2,700 mg·l−1, compared to 60 and 41 mg·l−1, respectively, when the hydrocarbon was presented in the form of liposomes. The data clearly demonstrate the importance of transport in the microbial metabolism of recalcitrant hydrocarbons.
Applications
Petroleum microbiology began as an applied subject, and the applied aspects continue to provide the primary impetus for research in this field. Current areas of applied interest are:
-
1.
Microbial spoilage of petroleum products
-
2.
Treatment of oil spills and disposal of petroleum wastes
-
3.
Enhanced oil recovery
-
4.
Production of surface-active agents
-
5.
Hydrocarbons as substrates in industrial fermentation processes
Biodeterioration of petroleum products, such as fuels, lubricating oils, and oil emulsions, has obvious economic implications. Genner and Hill (1981) have reviewed the data on the microbial spoilage of petroleum products and emphasized that spoilage only occurs when the petroleum products come in contact with water. In addition to avoiding water, spoilage can sometimes be retarded by the use of biocides (Rogers and Kapian 1968) or membrane filtration.
In considering the microbial treatment of oil spills, it is essential to distinguish between open systems (e.g., the ocean) and closed ones (e.g., oil storage tanks). In the latter case, it is possible to supplement the system with appropriate sources of nitrogen, phosphates, oxygen, and seed bacteria to enhance microbial growth and petroleum degradation, emulsification, or both. Two early published accounts of the use of these fundamental microbiological principles to enhance oil conversion in a restricted area are the treatment of oily ballast water from an oil tanker (Gutnick and Rosenberg 1977) and of contaminated soil (Raymond et al. 1976). More recently, petroleum pollution has been treated by composting (Kirchmann and Ewnetu 1998), by thermophilic bacteria (Mueller and Nielsen 1996), in soil-water slurries (Zhang and Bouwer 1997), and by using water-insoluble fertilizers (Rosenberg et al. 1996; Knezevich et al. 2006). In an open system, such as the sea, the ability of resident bacteria to extensively degrade a large oil slick is limited primarily by the concentration of nitrogen and phosphorus. Since there is no economical technology for overcoming these nutrient limitations in an open system, there is at present no practical microbial solution for oil spills at sea.
The use of microorganism in tertiary oil recovery has been the subject of several international conferences and literature reviews (e.g., Westlake 1984; Moses and Springham 1982). After primary and secondary recovery (waterflooding) processes, approximately 70 % of the reservoir oil remains underground, trapped in pore spaces and bound to inorganic minerals. The potential use of microorganisms in situ to release this oil depends on the anaerobic production of organic solvents, such as ethanol and butanol, gasses, such as methane and carbon dioxide, and organic acids. These materials can help overcome the physical forces holding the oil in the reservoir. Also, acid production can dissolve carbonates thus increasing the permeability of the reservoir. In addition, microbial products could enhance oil recovery by producing surface-active material and viscosity-altering polymers. Although the evidence for the positive role of microorganism in enhanced oil recovery is limited to a few poorly controlled experiments (Hitzman 1983), the enormous potential of this technology warrants further investigation. In recent years, interest in bioemulsifiers and other microbial surface-active agents has been growing. Many of these compounds are produced by hydrocarbon-degrading microorganisms (Rosenberg 1986; Rosenberg and Ron 1997; Desai and Banat 1997). The advantages of microbially produced surfactants include (1) biodegradability and controlled inactivation, (2) diversity of structure and function for different applications, (3) selectivity for specific hydrocarbon/water interfaces, and (4) characteristic surface modifications.
The use of hydrocarbons as inexpensive raw materials for the production of single-cell protein (SCP) was stimulated by the publications of Champagnat and Llewelyn (1962) and Champagnat et al. (1963). During the 1960s, many large oil and fermentation companies were involved in large-scale research and development projects for the conversion of petroleum fractions into SCP. Although the anticipated market for SCP in human and animal nutrition was not realized, these technological developments have provided a rich source of information about how bacteria grow on petroleum, how a continuous process can be scaled-up, and how bulk products can be recovered economically. In the 1970s, several fermentation plants were operating with capacities of 100,000 t of SCP per year. These were the largest biotechnology plants ever built. Because of the increased cost of hydrocarbon feedstock and more stringent governmental regulations governing its use in fermentation industry, there are presently no large-scale commercial fermentation processes based on hydrocarbon substrates. There are, however, a number of excellent microbial processes that have already been developed; these could be activated under the right set of economic conditions. These include processes for producing alcohols, organic acids, and ketones from specific alkanes; single-cell (food) oil from mixed n-paraffins; and large numbers of microbiological metabolites, including vitamins, amino acids, pigments, polysaccharides, enzymes, and alkanes.
References
American Public Health Association (1995) Standard methods for the examination of water, sewage and industrial wastes. American Public Health Association, New York
Antoniewski J, Schaefer R (1972) Researches sur les reactions des coenoses microbiennes de sols impregnes par des hydrocarbures. Modification de l’activite respiratoire. Ann Inst Pasteur 123:805–819
Asperger O, Kleber HP (1991) Metabolism of alkanes y Acinetobacter. In: Towner KJ, Bergogne-Berezine E, Fewson CA (eds) The biology of Acinetobacter. Plenum Press, New York, pp 323–351
Atlas RM (1981) Microbial degradation of petroleum hydrocarbons: an environmental perspective. Microbiol Rev 45:180–209
Atlas RM, Bartha R (1972) Degradation and mineralization of petroleum by two bacteria isolated from coastal waters. Biotechnol Bioeng 14:297–305
Atlas RM, Bartha R (1973a) Stimulated biodegradation of oil slicks using oleophilic fertilizers. Environ Sci Technol 7:535–541
Atlas RM, Bartha R (1973b) Abundance, distribution and oil biodegradation potential of microorganisms in Raritan Bay. Environ Pollut 4:291–300
Atlas RM, Schofield EA (1975) Petroleum biodegradation in the Arctic. In: Bourquin AW, Ahearn DG, Meyers SP (eds) Impact on the use of microorganisms on the aquatic environment. Environmental Protection Agency, Corvallis, pp 183–198. EPA-660-3-75-001
Atlas RM, Horowitz A, Busdosh M (1978) Prudhoe crude oil in Arctic marine ice, water and sediment eco-systems; degradation and interactions with microbial and benthic communities. J Fish Res Board Can 35:585–590
Austin B, Calomiris JJ, Walker JD, Colwell RL (1977a) Numerical taxonomy and ecology of petroleum-degrading bacteria. Appl Environ Microbiol 34:60–68
Austin B, Colwell RR, Walker JD, Calomiris JJ (1977b) The application of numerical taxonomy to the study of petroleum degrading bacteria isolated from the aquatic environment. Dev Ind Microbiol 18:685–695
Barabas G, Sorkhoh NA, Fardoon F, Radwan SS (1995) n-Alkane-utilization by oligocarbophilic actinomycete strains from oil-polluted Kuwaiti desert soil. Actinomycetol 9:13–18
Bartha R, Atlas RM (1977) The microbiology of aquatic oil spills. Adv Appl Microbiol 22:225–266
Beauchop T, Elsden SR (1960) The growth of organisms in relation to their energy supply. J Gen Microbiol 23:457–469
Bertrand JC, Dour JM, Azoulay E (1976) Metabolisme des hydrobarbures chez une bacterie marine. Biochemie 58:843–854
Bossert L, Bartha R (1984) The fate of petroleum in oil ecosystems. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 435–473
Buckley EN, Jones RB, Pfaender FF (1976) Characterization of microbial isolates from an estuarine eco-system: relationship of hydrocarbon utilization to ambient hydrocarbon concentration. Appl Environ Microbiol 32:232–237
Burback BL, Perry JJ (1993) Biodegradation and biotransformation of groundwater pollutant mixtures by Mycobacterium vaccae. Appl Environ Microbiol 59:1025–1029
Byrom JA, Beastall S, Scotland S (1970) Bacterial degradation of crude oil. Mar Pollut Bull 1:25–26
Cappello S, Yakimov MM (2010) Alcanivorax. In: Timmis K (ed) Handbook of hydrocarbon and lipid microbiology. Springer, Heidelberg
Cerniglia CE (1984) Microbial transformation of aromatic hydrocarbons. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 95–128
Cerniglia CE (1992) Biodegradation of polycyclic aromatic hydrocarbons. Biodegradation 3:351–368
Chakrabarty AM, Chou G, Gunsalas LC (1973) Genetic regulation of octane dissimilation plasmid in Pseudomonas. Proc Natl Acad Sci 70:1137–1140
Champagnat A, Llewelyn DAB (1962) Protein from petroleum. New Sci 16:612–613
Champagnat A, Verne C, Laine B, Filosa J (1963) Biosynthesis of protein-vitamin concentrates. Nature 197:13–14
Churchill SAJP (1999) Isolation and characterization of a Mycobacterium species capable of degrading three-and four-ring aromatic and aliphatic hydrocarbons. Appl Environ Microbiol 65:549–552
Colwell RR, Walker JD, Nelson JD Jr (1973) Microbial ecology and the problem of petroleum degradation in Chesapeake Bay. In: Ahearn DG, Meyers SR (eds) The microbial degradation of oil pollutants. Center for Wetland Resources, Baton Rouge, pp 186–197. Publ. No. LSU-SG-73-91
Cook WL, Massey JL, Ahearn DG (1973) The degradation of crude oil by yeasts and its effects on Lebistes reticulatis. In: Ahearn DG, Meyers SP (eds) The microbial degradation of oil pollutants. Center for Wetland Resources, Baton Rouge, pp 252–297. Publ. No. LSU-SG-73-01
Coty VF (1967) Atmospheric nitrogen fixation of hydrocarbon-oxidizing bacteria. Biotechnol Bioeng 9:25–32
Crow SA, Hood MA, Meyers SP (1975) Microbiological aspects of oil intrusion in southeastern Louisiana. In: Bourquin AW, Ahearn DG, Meyers SP (eds) Impact of the use of microorganism on the aquatic environment. Environmental Protection Agency, Corvallis, pp 221–227. EPA-660-3-75-001
Cundell AM, Traxler RW (1973a) The isolation and characterization of hydrocarbon utilizing bacteria from Chedabucto Bay, Nova Scotia. In: Proceedings of joint conference on prevention and control of oil spills. American Petroleum Institute, Washington, DC, pp 421–426
Cundell AM, Traxler RW (1973b) Microbial degradation of petroleum at low temperature. Mar Pollut Bull 4:125–127
Cundell AM, Traxler RW (1976) Psychrophillic hydrocarbon degrading bacteria from Narragansett Bay, Rhode Island, USA. Mater Org 11:1–17
Desai JD, Banat IM (1997) Microbial production of surfactants and their commercial potential. Microbiol Mol Biol Rev 61:47–64
Dibble JT, Bartha R (1976) Effect of iron on the biodegradation of petroleum in seawater. Appl Environ Microbiol 31:544–550
Evans WC, Fernley HN, Griffiths E (1965) Oxidative metabolism of phenanthrene and anthracene by soil pseudomonads: the ring-fission mechanism. Biochem J 95:819–831
Fehler SWG, Light RJ (1970) Biosynthesis of hydrocarbons in Anabaena variabilis. Incorporation of (methyl-14C)-and (methyl2H3)-methionine into 7-and 8-methyl-heptadecanes. Biochemistry 9:418–422
Floodgate GD (1973) A threnody concerning the biodegradation of oil in natural waters. In: Ahearn DG, Meyers SP (eds) The microbial degradation of oil pollutants. Center for Wetland Resources, Baton Rouge, pp 17–22. Publication No. LSU-SG-73-01
Floodgate GD (1984) The fate of petroleum in marine ecosystems. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 355–398
Foght JM, Gutnick DL, Westlake DWS (1989) Effect of emulsan on biodegradation of crude oil by pure and mixed bacterial cultures. Appl Environ Microbiol 55:36–42
Geiselbrecht AD, Herwig RP, Deming JW, Staley JT (1996) Enumeration and phylogenetic analysis of polycyclic aromatic hydrocarbon-degrading marine bacteria from Puget Sound sediments. Appl Environ Microbiol 62:3344–3349
Genner C, Hill EC (1981) Fuels and oils. In: Rose AH (ed) Microbial biodeterioration. Academic, London, pp 260–306
Gibson DT (1968) Microbial degradation of aromatic compounds. Science 161:1093–1097
Gibson DT (1971) Microbial degradation of hydrocarbons. In: Goldberg ED (ed) Physical and chemical sciences research report I. Dahlem workshop report on the nature of sea water, pp 667–696
Gibson DT (1977) Biodegradation of aromatic petroleum hydrocarbons. In: Wolfe DA (ed) Fate of and effect of petroleum hydrocarbons in marine eco-systems and organisms. Pergamon Press, New York, pp 34–46
Giger W, Blumer M (1974) Polycyclic aromatic hydrocarbons in the environment: isolation and characterization by chromatography, visible, ultraviolet and mass spectrometry. Anal Chem 46:1663–1671
Griffol M, Selifonov SA, Chapman PJ (1994) Evidence for a novel pathway in the degradation of fluorene by Pseudomonas sp. Strain. Appl Environ Microbiol 60:2438–2449
Gunkel W, Trekel HH (1967) Zur Methodik der quantitative Erfassung olabbauender Sakterien in verolten Sedimenten und Boden, Ol-Wassergemischen, Olen and Teerartigen Substanzen. Helgolander wiss Meeresunters 16:336–348
Gutnick DL, Rosenberg E (1977) Oil tankers and pollution: a microbiological approach. Ann Rev Microbiol 31:379–396
Haines JR, Wrenn BA, Holder EL, Strohmeier KL, Herrington RT, Venosa AD (1996) Measurement of hydrocarbon-degrading microbial populations by a 96-well plat most-probable-number procedure. J Ind Microbiol 16:36–41
Hardwood JL, Russel NJ (1984) Lipids in plants and microbes. George Allen & Unwin, London, pp 110–111
Hisatsuka K, Nakahara T, Sano N, Yamada K (1971) Formation of rhamnolipid by Pseudomonas aeruginosa and its function in hydrocarbon fermentation. Agric Biol Chem 35:686–692
Hitzman DO (1983) Petroleum microbiology and the history of its role in enhanced oil recovery. In: Donaldson ES, Clark SB (eds) Proceedings of 1982 intentional conference on the microbial enhancement of oil recovery technology transfer branch. Bartlesville Energy Technology Center, Bartlesville, pp 162–218
Hollinger C, Zehnder AJ (1996) Anaerobic biodegradation of hydrocarbons. Curr Opin Biotechnol 3:326–330
Hood MA, Bishop WS Jr, Bishop FW, Meyers SP, Whelan T III (1975) Microbial indicators of oil-rich salt marsh sediments. Appl Microbiol 30:982–987
Horowitz A, Atlas RM (1977a) Continuous open flow-through system as a model for oil degradation in the Arctic Ocean. Appl Environ Microbiol 33:647–653
Horowitz A, Atlas RM (1977b) Response of microorganism to an accidental gasoline spoilage in an Arctic freshwater ecosystem. Appl Environ Microbiol 33:1252–1258
Horowitz A, Gutnick D, Rosenberg E (1975) Sequential growth of bacteria on crude oil. Appl Microbiol 30:10–19
Hunt JM, Miller RJ, Whelan JL (1980) Formation of C6,-C7 hydrocarbons from bacterial degradation of naturally occurring terpenoids. Nature (London) 288:577–578
Itoh S, Suzuki T (1972) Effect of rhamnolipids on growth of Pseudomonas aeruginosa mutant deficient in n-paraffin utilizing ability. Agric Biol Chem 6:2233–2235
Jensen V (1975a) Decomposition of oil wastes in soil. In: Kilbertus G, Reisinger O, Mourey A, Cancela da Fonseca J (eds) Proceedings of the first international conference on biodegradation and humification 1974. University of Nancy, Nancy
Jensen V (1975b) Bacterial flora of soil after application of oily waste. Oikios 26:152–158
Jobson A, Cook FD, Westlake DWS (1972) Microbial utilization of crude oil. Appl Microbiol 23:1082–1089
Jones JG, Edington WA (1968) An ecological survey of hydrocarbon-oxidizing microorganisms. J Gen Microbiol 52:381–390
Juttner F (1976) Beta-Cyclocitral and alkanes in microcystis (Cyanophyceae). Zeitschrift fur Naturforschung 31c:491–495
Kappeli O, Finnerty WR (1980) Characteristics of hexadecane partition by the growth medium of Acinetobacter sp. Biotechnol Bioeng 22:495–503
Kincannon CB (1972) Oily waste disposal by soil cultivation process. Government Printing Office, Washington, DC. EPA Publ. No. R2-72-110
Kirchmann H, Ewnetu W (1998) Biodegradation of petroleum-based oil wastes through composting. Biodegradation 9:151–156
Kiyohara H, Nagao L, Kauno L, Yano L (1982) Phenanthrene-degrading enzyme phenotype of Alcaligenes faecalis AFK2. Appl Environ Microbiol 43:458–461
Knezevich V, Koren O, Ron EZ, Rosenberg E (2006) Petroleum bioremediation in seawater using guano as the fertilizer. Bioremediat J 10:83–91
Kolattukudy PE, Buckner JS, Brown L (1972) Direct evidence for a decarboxylation mechanism in the biosynthesis of alkanes in B. oleracea. Biochem Biophys Res Commun 47:1306–1313
Leahy JG, Batchelor PJ, Morcomb SM (2003) Evolution of the soluble diiron monooxygenases. FEMS Microbiol Rev 27:449–479
Lindstrom JE, Prince RC, Clark JC, Grossman MJ, Yeager TR, Braddock JF, Brown EJ (1991) Microbial populations and hydrocarbon biodegradation potentials in fertilized shoreline sediments affected by the T/V Exxon Valdez oil spill. Appl Environ Microbiol 57:2514–2522
Liu Z, Jacobson AM, Luthy RG (1995) Biodegradation of naphthalene in aqueous nonionic surfactant systems. Appl Environ Microbiol 61:145–151
Makula RA, Lockwood PJ, Finnerty WR (1975) Comparative analysis of the lipids of Acinetobacter species grown on hexadecane. J Bacteriol 121:250–258
Margesin R, Schinner F (1997) Efficiency of indigenous and inoculated cold-adapted soil microorganisms for biodegradation of diesel oil in alpine soils. Appl Environ Microbiol 63:2660–2664
Markovetz AJ (1971) Subterminal oxidation of aliphatic hydrocarbons by microorganism. CRC Crit Rev Microbiol 1:225–238
McKee JE, Laverty FB, Hertel RM (1972) Gasoline in groundwater. J Water Pollut Contr Fed 44:293–302
Mihelcic JR, Luthy RG (1988) Degradation of polycyclic aromatic compounds under various redox conditions in soil-water system. Appl Environ Microbiol 54(1):1182–1187
Mikkelson JD, von Wettstein-Knowles P (1978) Biosynthesis of beta-diketones and hydrocarbons in barley spike epicuticular wax. Arch Biochem Biophys 188:172–181
Miller RM, Bartha R (1989) Evidence from liposome encapsulation for transport-limited microbial metabolism of solid alkanes. Appl Environ Microbiol 55:269–274
Mimura A, Takeda I, Wakasa R (1973) Some characteristic phenomena of oxygen transfer in hydrocarbon fermentation. Biotechnol Bioeng Symp (4):467–484. Wiley, New York
Mironov OC (1970) Role of microorganism growing on oil in the self purification and indication of oil pollution in the sea. Oceanology 10:650–656
Mironov OC, Lebed AA (1972) Hydrocarbon oxidizing bacteria in the North Atlantic. Hydrobiol J 8:74
Moses V, Springham DG (1982) Bacteria and the enhancement of oil recovery. Applied Science, London
Mueller RF, Nielsen PH (1996) Characterization of thermophilic consortia from two souring oil reservoirs. Appl Environ Microbiol 62:3083–3087
Mulkins-Phillips GJ, Stewart JE (1974a) Effect of environmental parameters on bacterial degradation of Bunker C. oil, crude oils, and hydrocarbons. Appl Microbiol 28:915–922
Mulkins-Phillips GJ, Stewart JE (1974b) Distribution of hydrocarbon utilizing bacteria in north western Atlantic waters and coastal sediments. Can J Microbiol 20:955–962
Nakahara T, Hisatsuka K, Minoda Y (1981) Effect of hydrocarbon emulsification on growth and respiration of microorganism in hydrocarbon media. J Ferm Technol 59:415–418
Nieder M, Shapiro J (1975) Physiological function of Pseudomonas putida PpG6 (Pseudomonas oleovarans) alkane hydroxylase: monoterminal oxidation of alkanes and fatty acids. J Bacterial 122:93–98
Odu CTI (1978) Fermentation characteristics and biochemical reactions of some organisms isolated from oil-polluted soils. Environ Pollut 15:271–276
Pérez-Pantoja D, González B, Pieper DH (2010) Aerobic degradation of aromatic hydrocarbons. In: Timmis K (ed) Handbook of hydrocarbon and lipid microbiology. Springer, Heidelberg
Perry JJ (1977) Microbial metabolism of cyclic hydrocarbons by microorganisms isolated from soil. Can J Microbiol 14:403–407
Perry JJ (1984) Microbial metabolism of cyclic alkanes, R61-98. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 61–67
Pirnik MP (1977) Microbial oxidation of methyl branched alkanes. CRC Crit Rev Microbiol 5:413–422
Powlowski J, Sealy J, Shingler V, Cadieux E (1997) On the role of DmpK, an auxiliary protein associated with multicomponent phenol hydroxylase from Pseudomonas sp. CF600. J Biol Chem 272:945–951
Rabus R, Wilkes H, Schramm A, Harms G, Behrends A, Amann R, Widdel F (1999) Anaerobic utilization of alkylbenzenes and n-alkanes from crude oil in an enrichment culture of denitrifying bacteria affiliating with the beta-subclass of Proteobacteria. Environ Microbiol 1:145–157
Ratajczak A, Geibdorfer W, Hillen W (1998) Expression of alkane hydroxylase from Acinetobacter sp. Strain ADP1 is induced by a broad range of n-alkanes and requires the transcriptional activator AlkR. J Bacteriol 180:5822–5827
Raymond RL, Hudson JO, Jamison VW (1976) Oil degradation in soil. Appl Environ Microbiol 31:522–535
Reisfeld A, Rosenberg E, Gutnick D (1972) Microbial degradation of crude oil: factors affecting the dispersion in sea water by mixed and pure cultures. Appl Microbiol 24:363–368
Robertson B, Arhelger S, Kinney PJ, Button DL (1973) Hydrocarbon degradation in Alaskan waters. In: AhearnDO, Meyers SP (eds) The microbial degradation of oil pollutants. Center for Wetland Resources, Baton Rouge, pp 171–184. Publication No. LSU-SG-73-001
Rogers MR, Kapian AM (1968) Screening of prospective biocides for hydrocarbon fuels. Dev Ind Microbiol 9:448–476
Rosenberg E (1986) Microbial surfactants. CRC Crit Rev Biotechnol 3:109–132
Rosenberg E, Ron EZ (1997) Bioemulsans: microbial polymeric emulsifiers. Curr Opin Biotechnol 8:313–316
Rosenberg M, Rosenberg E (1985) Bacterial adherence at the hydrocarbon-water interface. Oil Petrochem Pollut 2:155–162
Rosenberg M, Bayer EA, Delaria J, Rosenberg E (1982) Role of thin fimbriae in adherence and growth of Acinetobacter calcoaceticus RAG-1 on hexadecane. Appl Environ Microbiol 44:929–937
Rosenberg E, Kaplan N, Pines O, Rosenberg M, Gutnick D (1983) Capsular polysaccharides interfere with adherence of Acinetobacter. FEMS Microbiot Lett 17:157–161
Rosenberg E, Brown DR, Demain AL (1985) The influence of gramicidin S on hydrophobicity of germinating Bacillus brevis spores. Arch Microbiol 142:51–54
Rosenberg E, Rosenberg M, Shoham Y, Kaplan N, Sar N (1989) Adhesion and desorption during the growth of Acinetobacter calcoaceticus on hydrocarbons. In: Cohen Y, Rosenberg E (eds) Microbial mats. ASM, Washington, DC, pp 218–226
Rosenberg E, Legmann R, Kushmaro A, Taube R, Adler E, Ron E (1992) Petroleum bioremediation—a multiphase problem. Biodegradation 3:337–350
Rosenberg E, Legmann R, Kushmaro A, Adler E, Abir H, Ron EZ (1996) Oil bioremediation using insoluble nitrogen source. J Biotechnol 51:273–278
Rosenfeld WD (1947) Anaerobic oxidation of hydrocarbons by sulfate-reducing bacteria. J Bacteriol 54:664–665
Rueter P, Rabus R, Wilkes H, Aeckersberg F, Rainey FA, Jannasch HW (1994) Anaerobic oxidation of hydrocarbons in crude oil by new types of sulfate-reducing bacteria. Nature (London) 372:455–458
Schneiker S, Martins dos Santos VA, Bartels D, Bekel T, Brecht M, Buhrmester J, Chernikova TN, Denaro R, Ferrer M et al (2006) Genome sequence of the ubiquitous hydrocarbon-degrading marine bacterium Alcanivorax borkumensis. Nat Biotechnol 24:997–1004
Schocken MJ, Gibson DT (1984) Bacterial oxidation of the polycyclic aromatic hydrocarbon acenaphthalene. Appl Environ Microbiol 48:10–16
Senez JC, Azoulay E (1961) Dehydrogenation of d’hydrocarbures parafliniques par leis suspensions non-proliferants et les extracts de Pseudomonas aeruginosa. Biochimica et Biophysical Acta 47:307–316
Simon MJ, Osslund TD, Saunders R, Ensley BD, Suggs S, Harcourt A, Suen WC, Cruden DL, Gibson DT, Zylstra GJ (1993) Sequences of genes encoding naphthalene dioxygenase in Pseudomonas putida strains G7 and NCIB 9816-4. Gene 127:31–37
Singer ME, Finnerty WL (1984) Microbial metabolism of straight-chain and branched alkanes. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 1–60
Soli G (1973) Marine hydrocarbonoclastic bacteria: types and range of oil degradation. In: Ahearn DG, Meyers SP (eds) The microbial degradation of oil pollutants. Center for Wetland Resources, Baton Rouge, pp 141–146. Publ. No. UU-SG-73-001
Song H-G, Bartha R (1990) Effects of jet fuel spills on the microbial community of soil. Appl Environ Microbiol 56:646–651
Stevenson JJ (1966) Lipids in soil. J Am Oil Chem Soc 43:203–210
Thibault SL, Anderson M, Frankenberger WT Jr (1996) Influence of surfactants on pyrene desorption and degradation in soils. Appl Environ Microbiol 62:283–287
Tiehm A (1994) Degradation of polycyclic aromatic hydrocarbons in the presence of synthetic surfactants. Appl Environ Microbiol 60:258–263
Vestal R, Cooney JJ, Crow S, Berger J (1984) The effects of hydrocarbons on aquatic microorganisms. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 475–506
Walker JD, Colwell RR (1974) Microbial degradation of model petroleum at low temperatures. Microbiol Ecol 1:63–95
Walker JD, Colwell RR (1975) Factors affecting the enumeration and isolation of Actinomyces from Chesapeake Bay and south eastern Atlantic Ocean sediments. Mar Biol 30:193–201
Walker JD, Colwell RR (1976a) Measuring potential activity of hydrocarbon degrading bacteria. Appl Environ Microbiol 31:189–197
Walker JD, Colwell RR (1976b) Enumeration of petroleum-degrading microorganism. Appl Environ Microbiol 31:195–207
Walker JD, Seesman PA, Herbert TL, Colwell RR (1976) Petroleum hydrocarbons: degradation and growth potential of deep-sea sediment bacteria. Environ Pollut 10:89–99
Ward DM, Brock TD (1976) Environmental factors influencing the rate of hydrocarbon oxidation in temperate lakes. Appl Environ Microbiol 31:764–772
Westlake DWS (1984) Heavy crude oils and oil shales: tertiary recovery of petroleum from oil-bearing formations. In: Atlas RM (ed) Petroleum microbiology. Macmillan, New York, pp 537–552
Westlake DWS, Jobson A, Philippe R, Cooke FD (1974) Biodegradability and crude oil composition. Can J Microbiol 20:915–928
Whyte LG, Hawari J, Zhou E, Bourbonniere L, Inniss WE, Greer CW (1998) Biodegradation of variable-chain-length alkanes at low temperatures by a psychrotrophic Rhodococcus sp. Appl Environ Microbiol 64:2578–2584
Winters L, Parker PL, Van Baalen C (1969) Hydrocarbons of the blue-green algae: geochemical significance. Science 163:467–468
Wyndham RC, Costenon JW (1981) Heterotrophic potentials and hydrocarbon biodegradation potentials of sediment microorganisms within the Athabasca oil sands deposit. Appl Environ Microbiol 41:783–790
Zhang W, Bouwer EJ (1997) Biodegradation of benzene, toluene and naphthalene in soil-water slurry microcosms. Biodegradation 8:167–175
Zhang Y, Miller RM (1994) Effect of a Pseudomonas rhamnolipid biosurfactant on cell hydrophobicity and biodegradation of octadecane. Appl Environ Microbiol 60:2101–2106
ZoBell CE (1964) The occurrence, effects and fate of oil polluting the sea. Adv Water Pollut Res 3:85–118
ZoBell CE, Prokop JF (1966) Microbial oxidation of mineral oils in Barataria Bay bottom deposits. Zeitschrifl fur Allgemeine Mikrobiologie 6:143–162
Author information
Authors and Affiliations
Corresponding author
Editor information
Editors and Affiliations
Rights and permissions
Copyright information
© 2013 Springer-Verlag Berlin Heidelberg
About this entry
Cite this entry
Rosenberg, E. (2013). Hydrocarbon-Oxidizing Bacteria. In: Rosenberg, E., DeLong, E.F., Lory, S., Stackebrandt, E., Thompson, F. (eds) The Prokaryotes. Springer, Berlin, Heidelberg. https://doi.org/10.1007/978-3-642-30141-4_66
Download citation
DOI: https://doi.org/10.1007/978-3-642-30141-4_66
Publisher Name: Springer, Berlin, Heidelberg
Print ISBN: 978-3-642-30140-7
Online ISBN: 978-3-642-30141-4
eBook Packages: Biomedical and Life SciencesReference Module Biomedical and Life Sciences