Populations of heavy fuel oil-degrading marine microbial community in presence of oil sorbent materials


Christoph Gertler, School of Biological Sciences, Environment Centre for Wales, Bangor University, Deiniol Road, Bangor, Gwynedd, LL57 2UW, United Kingdom. E-mail: c.gertler@bangor.ac.uk


Aims:  To investigate the feasibility of applying sorbent material X-Oil® in marine oil spill mitigation and to survey the interactions of oil, bacteria and sorbent.

Methods and Results:  In a series of microcosms, 25 different treatments including nutrient amendment, bioaugmentation with Alcanivorax borkumensis and application of sorbent were tested. Microbial community dynamics were analysed by DNA fingerprinting methods, RISA and DGGE. Results of this study showed that the microbial communities in microcosms with highly active biodegradation were strongly selected in favour of A. borkumensis. Oxygen consumption measurements in microcosms and gas chromatography of oil samples indicated the fast and intense depletion of linear alkanes as well as high oxygen consumption within 1 week followed by consequent slower degradation of branched and polyaromatic hydrocarbons.

Conclusion:  Under given conditions, A. borkumensis was an essential organism for biodegradation, dominating the biofilm microbial community formation and was the reason of emulsification.

Significance and Impact of the Study:  This study strongly emphasizes the pivotal importance of A. borkumensis as an essential organism in the initial steps of marine hydrocarbon degradation. Interaction with the sorbent material X-Oil® proved to be neutral to beneficial for biodegradation and also promoted the growth of yet unknown micro-organisms.


Marine ecosystems are permanently challenged with hydrocarbons of different composition and origin. The threat of oil pollution not only from natural sources such as marine oil seeps but also by anthropogenic activities endangers the marine biodiversity. As oil is transported over the seas, tanker disasters and wars have been responsible for cataclysmic oil spills all over the world, with dramatic oil spills during Persian Gulf War in 1991 being the most significant. Dependent on oil type and magnitude of the spill, enormous sums have to be spent to clean up beaches or remove oil slicks from sea surface or its bottom as can be seen in two recent disasters, the T/V ‘Prestige’ oil spill and the destruction of Jiyyeh power plant near Beirut, Lebanon, in July 2006. In the first case, a spill of more than 80 000 m³ of heavy fuel oil, which caused clean-up costs of more than $2·8 billion US occurred. Owing to the inaccessibility of the T/V ‘Prestige’s wreck, more than 20 l of oil per day still continue to leak from the ship’s tanks. The Jiyyeh event has led to the spill of 19 000 m³ of heavy fuel oil that polluted 150 km of shoreline and severely damaged the environment and fisheries in Lebanon. Exactly US$64 million of international aid have so far been granted, which is still not sufficient for a complete clean-up. Not only do accidents involving tankers and refineries contribute to the pollution of the sea, but the drastic increase of shipping activity and illegal waste dumping in the present decade has led to problems of a chronic pollution with oil hydrocarbons.

The biodegradation of oil hydrocarbons is a process well established in nature and known to man for a long time (Sohngen 1913). Mostly limited due to the low mineral nutrient levels in seawater, biodegradation of hydrocarbons is conducted by numerous genera of bacteria, fungi and algae (Head et al. 2006). Knowledge of oil-degrading bacteria and their nutritional requirements encouraged scientists to look for ways of employing this self-cleaning function of the seas since the seventies (Atlas and Bartha 1972a,b,c; Jobson et al. 1972; Reisfeld et al. 1972; Cerniglia and Perry 1973). A highlight of this research was a large-scale recovery operation following the T/V ‘Exxon Valdez’ oil spill in Alaska waters (Brown 1991). Two major components, biostimulation (addition of nutrients) and bioaugmentation (addition of hydrocarbon-degrading bacteria), for in situ biodegradation have been applied several times in the field with various successes (Lindstrom et al. 1991).

Two major spills of heavy fuel oil, the grounding of Russian tanker T/V ‘Nakhodka’ (Kasai et al. 2001) on the northern coast of Japan and the ‘Prestige’ disaster (Diez et al. 2005; Franco et al. 2006) on the Galician coast of Spain have recently been investigated closely for signs of biodegradation. Intense microbiological and molecular biological surveys made in the case of T/V ‘Nakhodka’ spill revealed different oil-degrading communities utilizing beached oil on several types of sediments and shoreline types (Harayama et al. 2004). Single dominant members of the oil-degrading consortia were identified by culture-independent methods. Most dominant among these bacteria was Alcanivorax borkumensis (Yakimov et al. 1998; Golyshin et al. 2003), which outgrows other bacteria in the presence of oil in marine environments (Hara et al. 2003; Roling et al. 2004). Its unique properties include obligate hydrocarbon metabolism selective for linear saturated hydrocarbons (Hara et al. 2004) and biosurfactant production (Abraham et al. 1998). Because of its ubiquity in marine environments and abundance in hydrocarbon-polluted seawater (Hara et al. 2003; Roling et al. 2004; Brakstad and Lodeng 2005), the genome and biochemistry of A. borkumensis has been extensively analysed (Dutta and Harayama 2001; Golyshin et al. 2003; Hara et al. 2003; van Beilen et al. 2004; Sabirova et al. 2006, 2008; Schneiker et al. 2006). Being a paradigm of marine hydrocarbonoclastic bacteria, this organism was chosen for bioaugmentation trials in the present study.

As A. borkumensis is not able to degrade aromatic heavy fuel oil constituents, other micro-organisms have to take part in later stages of heavy fuel oil degradation. Organisms like Cycloclasticus pugetti capable of degrading aromatic compounds (Kasai et al. 2002) were reported to play an important role at these stages of biodegradation (Maruyama et al. 2003; Yakimov et al. 2005). Numerous trials in microcosm (Lepo et al. 2003; Yakimov et al. 2005) or mesocosm scale (Bachoon et al. 2001; Roling et al. 2004; Yoshida et al. 2005) have been conducted to examine changes within microbial communities, considering mostly contaminations of sediment or open seawater.

In spite of the biotechnological and ecological progress of marine microbiology, so far no applicable biotechnological solution to marine oil spills has been invented and introduced into the world market, although there is a need for such an application for chronically polluted waters, i.e. oil terminals, harbours, for shallow waters, i.e. the Wadden Sea in German Bight or in third-world countries that cannot afford oil spill mitigation equipment. Recently, the application of oil sorbent material was added to the toolbox of oil spill mitigation techniques. Both synthetic (Wei et al. 2003) and natural (Khan et al. 2004; Pasila 2004; Suni et al. 2004; Annunciado et al. 2005) products are applied to bind and remove oil from the environment. Natural sorbents mostly consist of fibrous plant materials, i.e. peat moss or hemp (Pasila 2004). The major advantages of these materials are low production costs and they are nontoxic. Oil sorbent materials represent a yet unstudied alternative technique for oil spill mitigation purposes. Sorbents are easily storable, inexpensive and can be handled by untrained personnel. The major objective of these materials is the removal of oil by absorption to their hydrophobic surfaces. Thus, the spreading, dispersion or sedimentation of spilled oil can be prevented. Oil sorbents are also a potential key technology for offshore oil bioremediation purposes, as hydrocarbons can be immobilized and a matrix is provided for oil-degrading bacteria.

In this study, a novel type of recycling-based oil absorbent material X-Oil (Hellmann-tech, Lehrte, Germany) was applied. Made from textile and leather waste or shoes and consisting of textile or leather fibres, this material provides a biological support matrix and exhibits excellent oil absorption capacities. Using two culture-independent methods for analysis of 16S rRNA gene sequences, population changes were inspected and correlated to biodegradation in the microcosms. Important members of the oil-degrading communities were identified by sequencing of the 16S rRNA gene. In addition, multivariate statistics were applied to study the growth of oil-degrading microbial consortia and their population alterations. This constellation of experiments is considered to identify the basic conditions of bioremediation involving the oil absorption and to inspect its effects on marine heavy fuel oil degradation. This is the first study to investigate changes in bacterial community composition during the process of marine biodegradation of a heavy fuel oil adhering on an artificial oil-binding matrix.

Material and methods


Alcanivorax borkumensis SK2T (DSM 11573T; Yakimov et al. 1998) was used for augmentation experiments. It was routinely maintained on the solid ONR7a medium (Dyksterhouse et al. 1995), in vapours of hexadecane. The biomass of A. borkumensis SK2 was collected with an inoculation loop, resuspended in sterile liquid ONR 7a medium. The cell numbers were counted with a Neubauer chamber and diluted to a concentration of 5 × 108 cells ml−1. Exactly 250 μl of the cell suspension was added to each parallel of microcosms, 2–7 and 20–25, respectively.

Microcosm experiments

Twenty-five different microcosms were established in duplicates in 0·5 l Pyrex bottles. All microcosms were filled with seawater taken from the eastern port of the island of Helgoland (54°11_N, 7°33_E) on 10th August 2004. A part of the seawater samples were diluted at a ratio of 4 : 1 with distilled water and autoclaved. Several samples were amended with a fertilizer solution described previously (Gunkel 1967). Briefly, 150 ml of a 6·98 mmol l−1 Na2HPO4 × 2H2O, 2·87 mmol l−1 K2HPO4 and 9·35 mmol l−1 NH4Cl fertilizer solutions were mixed with 100 ml of distilled water and added to 750 ml of seawater. Table 1 shows the exact contents of each microcosm and the corresponding code numbers. The oil absorbent material X-Oil® (Hellmann-tech, Lehrte, Germany) was added in some experiments to serve as a model for oil spill-mitigation strategies. Except for controls, all microcosms were amended by 2·5 ml of bunker C heavy fuel oil (IFO 180) supplied by the Shell company (Stade, Germany). All microcosms contained 500 ml of aqueous phase and 0·5% heavy fuel oil (v/v) and were incubated aerobically in the dark on a shaker at 90 rev min−1 at the constant temperature of 17°C, which was the water temperature at the sampling site at the beginning of the experiment. Exactly 50 ml of microcosm was removed weekly for analysis and replaced by 50 ml of seawater from the very first seawater sampling, which was autoclaved, diluted to 80% with distilled water and stored at 4°C to prevent precipitation of sea salt. Sampling of each microcosm was performed on a weekly basis. In parallel to each sampling, the concentration of dissolved oxygen in every microcosm was measured by Oxyscan Graphic system (UMS, Meinersen, Germany). Out the samples taken weekly, bacterial biomass was collected via filtration with a 0·2 μm nitrocelluose filter (Sartorius, Göttingen) after 1, 2, 3, 4 and 6 weeks from 10 ml microcosm samples for consequent DNA extraction. Exactly 40 ml of microcosm water was stored at −20°C for hydrocarbon extraction.

Table 1.   Composition of microcosms used in the experiments and correlating code numbers. Augmentation (*) was performed by addition of Alcanivorax borkumensis biomass to a final concentration of 5 × 105 cells ml−1. Alcanivorax borkumensis biomass was grown in ONR7 medium containing 2% sodium pyruvate (w/v) at 25°C for 10 days. Seawater was taken as described in text. For sterilization, seawater was diluted to 80% with distilled water and autoclaved at 120°C for 20 min. Supplemented oil was heavy fuel oil IFO 180 (bunker C) kindly provided by Shell (Stade, Germany). For binding oil, two types of X-Oil materials made from leather (X-Oil (1)) or chopped shoes (X-Oil (2)), respectively, were applied. Both types of binding materials showed identical results in all experimental parameters. The fertilizers applied in this experiment have been prepared according to Gunkel (1967) as described in text
Microcosm no. Augmentation*Autoclaved seawaterNonsterile seawaterOilX-Oil (1)X-Oil (2)Fertillizer
  1. *‘+’ indicates addition or amendment of corresponding factor and ‘−’ stands for lack of thereof.


Extraction and gas chromatography of hydrocarbons from microcosms

Hydrocarbons from 40 ml of the microcosm samples were extracted thrice with N-pentane (Riedel-de-Haen, Seelze, Germany). Pentane phases were combined and concentrated by evaporation at room temperature overnight. Concentrated hydrocarbons were solubilized in 2 ml of N-pentane after gravimetric determination of extracted hydrocarbon mass. Heptamethyl-nonane was added to the resuspended hydrocarbons prior to high-resolution gas chromatography-mass spectrometry (GC-MS), which was conducted as described previously (Dutta and Harayama 2001; Wang et al. 2002).

DNA extraction and PCR

DNA from samples was extracted by phenol-chloroform as described previously (Wichels et al. 2004). Genomic DNA obtained was visualized on a 0·8% agarose gel and amplified by polymerase chain reaction (PCR) for RISA (Ribosomal Intergenic Spacer Analysis) (Ranjard et al. 2000, 2001) and denaturing gradient gel electrophoresis (DGGE; Wichels et al. 2004; Sigler et al. 2004). For RISA, the primers 132f (5′-CCGGGTTTCCCCATTCGG-3′) and 1522r (5′-TGCGGCTGGATCCCCTCCTT-3′) were used to amplify the intergenic spacers between 16S and 23S subunits of ribosomal RNA gene sequences. Exactly 100 μl of the PCR reaction mix contained 10 μl of 10× Taq buffer (Eppendorf), 20 μl of 5× Master Enhancer (Eppendorf), 300 μmol l−1 of each dNTP, 50 pmol of each primer and 2 U of Taq DNA Polymerase (Eppendorf). Exactly 10 ng of genomic DNA was added to each reaction. The amplification started with a denaturing step at 95°C for 3 min and 25 cycles at 95°C for 1 min, 53°C for 1 min and 72°C for 1 min followed by an extension step of 5 min at 72°C. The PCR reactions were performed in an Eppendorf Mastercycler. The presence of PCR products was proven by electrophoresis on a 1·4% (w/v) agarose gel and resolved on 8% polyacrylamide gels (QBiogene) in 0·5× TAE buffer with a Biorad Dcode gel electrophoresis device. About 10–20 μl of each PCR product was loaded on the polyacrylamide gels together with three 100 bp ladders (Invitrogen) for comparison and run at 50 V, 800 mA for 16 h. The gels were stained for 20 min with 10 ml of 0·5× TAE buffer containing SYBR-Gold stain (1 : 10 000; Sigma), illuminated on a ultraviolet (UV) table (2011 Macrovue Transilluminator; LKB Bromma, Pharmacia Biotech, Sweden). Band patterns were photographed on Polaroid 665 film with Polaroid MP4 equipment. Negative films were scanned and RISA profiles straightened, aligned and normalized by BioNumerics Gelcompare software (Applied Maths, Sint-Martens-Latern, Belgium).

For DGGE, the template DNA was amplified by a touchdown PCR described by Wichels et al. (2004) with primers 907r (5′-CCT ACG GGA GGC AGC AG-3′) and 341f including GC-clamp (5′-CGC CCG CCG CGC CCC GCG CCC GGC CCG CCG CCC CCG CCC CCC TAC GGG AGG CAG CAG CCT ACG GGA GGC AGC AG-3′). Exactly 100 μl of the PCR reaction mix contained 10 μl of 10× Taq buffer (Eppendorf), 20 μl of 5× Master Enhancer (Eppendorf), 300 μmol l−1 of each dNTP (Perkin Elmer), 50 pmol of each primer, 2 U of Taq DNA Polymerase (Eppendorf) and 1 μl of diluted sample genomic DNA (c. 20 ng).

A ‘touchdown’ PCR was used, starting with a denaturing step of 94°C for 5 min. Each of the following cycles consisted of a denaturing step of 94°C for 1 min and was completed by an elongation step of 72°C for 1 min. The annealing step of 1 min each in ‘touchdown’ PCR included initial annealing temperature of 65°C, which was decreased by 0·5°C in each cycle until a ‘touchdown’ of 55°C. Twelve further cycles were conducted at this temperature. Final primer extension was performed at 72°C for 10 min followed by 22 cycles of extension starting at 71°C and decreasing by 1°C per cycle in order to avoid heteroduplices. All PCR reactions were conducted in an Eppendorf Mastercycler. PCR products were inspected on 1·2% agarose gels. The PCR products were analysed as described before in a Biorad D-Code gel electrophoresis apparatus using 6% (w/v) polyacrylamide denaturing gradient gels with linear gradients from 15% to 70% for analysis of 20–55% bands (where 100% denaturant is 7 mol l−1 urea and 40% (v/v) formamide) for excision of DGGE. About 10–40 μl of each PCR product was loaded on the gels and run in 0·5× TAE buffer at 150 V, 200 mA at 60°C for 10 h. The gels were stained and photographed as described before (see RISA).

Phylogenetic analysis of RISA fingerprinting analysis

For comparison and analysis of RISA band patterns, Bionumerics Gelcompare software (Applied Maths) was used. For normalisation of all band patterns, 100 bp ladders were used as references. To compare the band patterns, a band-matching analysis was performed. Within all profiles, the bands were assigned to common classes. During this procedure, densitometric values of the profiles were included (Muylaert et al. 2002) resulting in a band-matching table. Bray-Curtis similarity of the RISA profiles was calculated using a square root transformation (Clarke and Warwick 2001) for each sample and the whole dataset. Ordination of the similarity matrices was performed by nonmetric multidimensional scaling (MDS; Clarke and Warwick 2001). For MDS, PRIMER 5 software was used (Clarke and Gorley 2001). For clearness, the MDS plots were presented in two dimensions although the stress levels were higher than in the three-dimensional plots including all the samples.

Sequencing of DNA fragments from DGGE gels

Selected DGGE bands were excised and eluted for 5 h at 37°C in 150 μl elution buffer containing 0·5 mol l−1 CH3COONH4, 1 mmol l−1 EDTA and 0·1% (w/v) sodium dodecyl sulphate (SDS). The solutions were centrifuged at 4°C, 10 000 g for 1 min, supernatant transferred in Eppendorf tubes and precipitated with 400 μl of ethanol at −20°C for 12–24 h. The resulting pellets were resuspended in 10 μl of MilliQ water. Exactly 1 μl of the solution was used for a PCR using primers 907r and 344f-IR (5′-ACG GGA GGC AGC AG-3′) using the PCR programme described before. Re-amplified PCR fragments resulting from DGGE bands gel-purified as described before were sequenced using Big Dye Terminator Kit (Applied Biosystems, Foster City, USA) in an Eppendorf Mastercycler. The sequencing reactions were cleaned with DyeEx Kit (Qiagen, Hilden, Germany) and analysed in an automated DNA sequencer according to the protocols provided by the manufacturer (Perkin Elmer). DNA sequences were assembled with CAP subroutine of BioEdit Sequence Alignment Editor according to Hall (1999) and aligned with ClustalW software (http://www.ebi.ac.uk/clustalw). The sequences were compared with those deposited in the GenBank using the Blast algorithm (http://www.ncbi.nlm.nih.gov/BLAST; Altschul et al.1990). The DNA sequences obtained in this study were deposited in the EBI database and can be referred to under the accession numbers FM211072FM211124.

Cloning of DNA fragments from DGGE gels

Owing to contamination of some DGGE bands with fragments of several other bands, the cloning of former DGGE fragments was necessary. PCR fragments from such bands were cloned into Escherichia coli DH 10B with TOPO TA cloning kit (Invitrogen) and One Shot TOP 10 cells according to the instructions of the supplier.

The clones were spread onto Luria-Bertani (LB) agar containing 50 μg ml−1 of kanamycin and 50 μg ml−1 of X-Gal for blue/white selection and incubated at 37°C overnight. Several resulting white clones were grown in 5 ml of LB medium containing 50 μg ml−1 of kanamycin for 16 h at 37°C. The plasmids were isolated with Nucleospin Plasmid Mini kit (Macherey-Nagel, Düren, Germany) and used for sequencing as described before.

DNA quantification

In order to approximately estimate the biomass of the microbial community, we determined the concentration of the DNA as the principal biomass constituent. The DNA yield was quantified using Pico Green stain (Hoechst, Germany) according to the supplier’s protocol.

Phylogenetic analysis of the sequences

For sequence assembly BioEdit software (version 2·3–2·5; see before) was used. Several relevant 16S rRNA gene sequences from the micro-organisms with the validly published names were selected from GenBank and added to the sequences obtained before. Multiple alignment of this set of sequences was conducted by ClustalW software (http://www.ebi.ac.uk/clustalw/). The resulting alignment was refined by BioEdit software and analysed with Phylip 3.5 software (http://www.evolution.genetics.washington.edu/phylip.html). For receiving robust phylogenetic trees, Phylip subroutines ‘seqboot’, ‘dnadist’ and ‘fitch’ were used to perform bootstrapping, distance matrix analysis and neighbour joining analysis as described previously (Yakimov et al. 2002). A total of 100 data sets was analysed for bootstrapping. Outgroup rooting with the 16S rDNA sequence of Methanococcus thermolithtrophicus was performed to prevent potential bias. A consensus tree was visualized with Treeview software (http://taxonomy.zoology.gla.ac.uk/rod/treeview.html).


Microcosm experiments

During the 6 weeks of experimentation, significant changes in several microcosms were observed. Microcosms with N/P supplementation showed a massive emulsification of the bunker C fuel changing the microcosm liquid from transparent to a black colour. This effect was already visible in the microcosm 7 after less than 7 days. An exceptionally fast emulsification could be seen in three microcosms containing the X-Oil® sorbent (microcosms 7, 19 and 25). Over the entire experiment, no emulsification was ever observed in nonsupplemented microcosms or controls, where bunker C fuel remained on the microcosm water surface as an oil slick. Phase-contrast microscopy of stained microcosms revealed a biofilm formation on X-Oil® fibres as well as on micelles in the microcosm’s aqueous phase (Supporting Information: Fig. S1a,b). The average concentration of dissolved oxygen in the negative control and the initial concentration in all microcosms was c. 10 mg ml−1. In all the microcosms, concentration of dissolved oxygen decreased within 7 days to 2–4 mg l−1 and slowly converged to 8 mg ml−1 within the following 4 weeks. Oxygen concentration and depletion is displayed in Fig. S2 of Supporting Information. In an A. borkumensis control microcosm containing an initial dose of 105 cells in sterile seawater, oil and nutrients, a steady decrease of dissolved oxygen concentration was observed, reaching a minimum of 2 mg l−1 after 2 weeks and a steady increase to 10 mg l−1 after that point in time. As it can be seen in Fig. S2, nutrient-amended microcosms (microcosm nos 7, 19, 25) showed a stronger depletion of oxygen than the nonsupplemented ones and a consequent slow increase to a level of 7 mg l−1 of dissolved oxygen from that point in time.

Both microcosm parallels treated with X-Oil®, soluble N/P-sources and augmentation with A. borkumensis (microcosm no. 25) showed a clarification of the emulsification after 6 weeks of experiment. DNA yields in nonbiostimulated microcosms were extremely low, whereas in nutrient-amended microcosms a steady increase of DNA could be measured. The increase in the cases of pure cultures and seawater microcosms was steady and ended at the concentration of 1500–2000 ng ml−1. In the case of augmented microcosms, a quick increase to a level of 2000 ng ml−1 was observed within the first two weeks of the experiment. This DNA yield remained constant for the remaining four weeks (data not shown).

GC analysis of oil samples

An exemplary quantitative analysis of selected microcosms is presented in Fig. 1. Results are shown in relation to the contents of untreated heavy fuel oil and a sterile control showing fuel oil extracted from a sterile seawater microcosm. Figure 1 displays the relative concentration of oil samples from the three microcosms treated with nutrients and sorbent material that contain different microbial communities in relation to the residues extracted from a control sample of heavy fuel oil. Samples were taken after 7 and 36 days, respectively.

Figure 1.

 Quantitative analysis of bunker C heavy fuel oil (Shell, Stade, Germany). All concentrations refer to a control sample of bunker C heavy fuel oil IFO 180 (Shell). WSC, sterile control: artificially weathered fuel oil in sterile seawater; Abo-PC, samples from pure culture of Alcanivorax borkumensis (microcosm no. 7); SW, microcosm based on pure seawater (microcosm no. 19); Aug-SW, microcosms based on seawater initially amended with 50 000 cells ml−1 (microcosm no. 25). All three types of microcosms were treated with nutrients and X-Oil material. The samples were taken after 1 and 6 weeks (7 days and, respectively, 36 days). Aliphatic HC (inline image), Branched HC (inline image), Polyaromatic HC (inline image).

As can be seen in Fig. 1, a significant decrease of aliphatic branched and polyaromatic hydrocarbons below the level of a sterile control (WSC) was seen in all the three sets of microcosms. Intensity and rate of this degradation obviously depended upon the inoculum or abundant microbial community.

The amount of transformed oil in microcosms containing nonsterile seawater (microcosm nos 19 and 25) in general was higher than in pure cultures of A. borkumensis. Moreover, the relative concentrations of all components measured after 36 days of the experiment were generally lower than those measured after 7 days. Interestingly, the relative concentration of polyaromatic hydrocarbons detected after 7 days was relatively similar between 58% and 45%, but decreased in microcosms containing the original seawater community within the following 29 days.

Aliphatic compounds on the other hand decreased in a different pattern depending on the type of the microcosm. After 36 days, more than 95% of the aliphatic compounds were transformed in the augmented microcosms, whereas the seawater-based microcosm (microcosm no. 19) still contained 18% whereas with the pure culture of A. borkumensis more than 30% of the residual hydrocarbons were remaining.

RISA fingerprints of bacterial communities and their statistical analysis

Selected results of RISA for several microcosm sets are displayed in Fig. 2a–c. The effects of single or combined treatments on the microbial community composition can be observed. All microcosms presented were initially set on plain seawater thus containing typical free-living marine bacteria. Effects of three factors of bioremediation (namely N/P supplementation, the addition of X-Oil® and A. borkumensis inoculum alone and in various combinations) on community composition can be seen. Changes in band patterns allowed a preliminary evaluation of the importance of a single parameter that might help in the development of future applications.

Figure 2.

 (a–c) Selected RISA profiles of experimental microcosms; profiles are vertically arranged in order of experiment time (1, 2, 3, 4, 6 weeks). White arrowheads point at bands that are affiliated for Alcanivorax borkumensis. Microcosm numbers of the samples presented are (from left to right) 14, 15, 20, 21, 24, 25.

In general, a reduction of the RISA band pattern diversity can be seen in all microcosms supplemented with soluble nutrients. In general, all nonsupplemented microcosms presented in Fig. 2a–c showed a total of more than 20 bands, especially the seawater microcosm (no. 14) presented in Fig. 2a. In nutrient-supplemented microcosms, this diversity of band patterns was strongly reduced; mostly to only two remaining bands marked with white arrows in Fig. 2a–c. Interestingly, this reduction seems to take place at different rates. In seawater microcosms, reduction to two bands can be seen after 4 weeks; in augmented microcosms after 3 weeks; and in augmented microcosms treated with X-Oil® sorbent even within the first 7 days. These phenomena were observed in the band patterns of both parallels of each microcosm.

MDS was used to visualize a matrix analysis comparing the large amount of band patterns gained and analysing effects of experiment time and treatment by connection of one or several experimental parameters to the landmarks of a MDS map. Figure 3 shows an MDS plot of one set of RISA profiles of five sampling points in time (1, 2, 3, 4, 6 weeks) for each microcosm type. To each landmark, experimental parameter oil spiking and nutrient amendment was connected. Obviously, the formation of two distinct clusters of samples can be seen, namely a large and widespread cluster of oil-spiked samples without nutrient addition and a condensed cluster of oil-spiked and N/P-supplemented samples. By addition of blank controls of a pure seawater sample, a pure culture sample of A. borkumensis and an initial sample of the augmented community, MDS clearly gives evidence that nutrient-amended oil-spiked samples altogether show a high similarity to a pure culture of A. borkumensis.

Figure 3.

 (a–h) Multivariate statistics of RISA fingerprints using multidimensional scaling (MDS); figures on the left-hand side show nonaugmented and pictures on the right-hand side show augmented microcosm communities. Impacts of addition of nitrogen/phosphorous source and X-Oil on communities are presented in (a–d). Numbers in the circles of (a) and (b) correspond to the week of sampling. (e–h) show changes in diversity and intensity of a band of 458 bp length, which was determined to be caused by Alcanivorax borkumensis. Stress levels for two-dimensional plots are given in the upper left corner of each figure.

Dominance of Alcanivorax borkumensis in experimental microcosms

For a closer look into the role of A. borkumensis in the experimental microcosms, the MDS plot presented in Fig. 3g, h was analysed with the ‘bubble plot’ subroutine of Primer 5 software. This tool enables the connection of a quantitative value to each landmark of the MDS plot. Therefore, the magnitude of a certain factor is proportional to the diameter of the circle in the position of the connected landmark. For demonstrating a potential dominance of an organism, the factor of a low diversity and abundance/intensity of a specific band were indicated in the MDS. According to a blank control of A. borkumensis SK2, a specific band of 458 bp was identified. Figure 3e, f show the number of detected bands per RISA band pattern, whereas Fig. 3g, h show the intensity of the 458 bp band. Comparison of Fig. 3e with 3g shows the clear decrease of diversity and number of bands correlate to the intensity of specific bands formed by A. borkumensis.

DGGE analysis of communities and phylogenetic affiliation of 16S rRNA genes from individual bands

In parallel to RISA analysis, the DGGE was performed for a phylogenetic analysis of bacterial populations involved in the oil degradation process. Yet, only samples from weeks 1, 3 and 6 derived from oil-spiked microcosms were applied. Resulting DGGE gels (Fig. 4a), contained samples from oil-spiked seawater-based microcosms and those from Fig. 4b contained those taken from augmented microcosms. A selection of distinct bands are marked in Fig. 4a, b and the corresponding phylogenetic trees can be seen in Fig. 5a, b. In general, there was a higher diversity of band patterns registered in samples containing sorbent and N/P source. This was found particularly in the upper part of the gel, which contained a low concentration of denaturing agents. Several significant bands marked with 15, 44, 70 and 78 were detected in these parts of the DGGE gels. These bands mostly belong either to Alphaproteobacteria and particularly form a cluster of species closely related to the genus Roseobacter or to the Cytophaga-Flavobacterium group. Both phylogenetic groups comprise common members of the natural marine microflora. Two bands that commonly appear in most band patterns belong to Thalassospira lucentensis and Ruegeria atlantica. Because of their similar GC content, both bands could not be clearly distinguished under the given conditions. A very prominent and intense band was observed as well among augmented and N/P-supplemented samples as in the seawater microcosms, marked with 45 and 73, respectively. These DGGE bands were attributed to A. borkumensis because of their 100% sequence identity to the rRNA gene fragment of this organism. Samples treated with X-Oil contained an intense and unique band that was marked with 39, 49 and 81. These were affiliated with a previously unknown and unidentified species of Spirochaeta. The three bands furthermore had 16S rRNA gene sequences identical each to other. In contrast to this, the samples of microcosms not supplied with nutrients (microcosm nos 14, 20 and 24) showed band patterns of a similar type.

Figure 4.

 (a) Denaturing gradient gel electrophoresis (DGGE) profiles of seawater microcosms (microcosm no., from left to right, 14, 15, 18, 19); the numbers correspond to excised bands; (b) DGGE profiles of augmented microcosms (microcosm no. from left to right, 20, 21, 24, 25); the numbers correspond to excised bands.

Figure 5.

Figure 5.

 (a) Phylogenetic consensus tree of phylotypes resulting from denaturing gradient gel electrophoresis (DGGE) band excision and sequencing; source of samples was seawater-based microcosm only. Each number in the tree is equivalent to the DGGE band presented in Fig. 4a. Robustness of the tree was examined by bootstrapping performed for 100 data sets. Open circles on the branching nodes of the tree indicate bootstrap values between 50% and 75%, whereas closed circles indicate 75% or more trees that show an identical branching. (b) Phylogenetic consensus tree of phylotypes resulting from DGGE band excision and sequencing; the source of the samples was augmented seawater-based microcosm only. The numbers of the sequences correspond to the DGGE band from Fig. 4b. Robustness of the tree was examined by bootstrapping performed for 100 data sets. Open circles on the branching nodes of the tree indicate bootstrap values between 50% and 75%, whereas closed circles indicate 75% or more trees that show an identical branching.

Figure 5.

Figure 5.

 (a) Phylogenetic consensus tree of phylotypes resulting from denaturing gradient gel electrophoresis (DGGE) band excision and sequencing; source of samples was seawater-based microcosm only. Each number in the tree is equivalent to the DGGE band presented in Fig. 4a. Robustness of the tree was examined by bootstrapping performed for 100 data sets. Open circles on the branching nodes of the tree indicate bootstrap values between 50% and 75%, whereas closed circles indicate 75% or more trees that show an identical branching. (b) Phylogenetic consensus tree of phylotypes resulting from DGGE band excision and sequencing; the source of the samples was augmented seawater-based microcosm only. The numbers of the sequences correspond to the DGGE band from Fig. 4b. Robustness of the tree was examined by bootstrapping performed for 100 data sets. Open circles on the branching nodes of the tree indicate bootstrap values between 50% and 75%, whereas closed circles indicate 75% or more trees that show an identical branching.


This is the first study to address the biodegradation of a common, yet specific pollutant in marine systems in combination with a novel type of oil sorbent material and a variety of other typical tools of marine oil spill mitigation, biostimulation (application of mineral nutrients to the polluted site) and bioaugmentation (addition of micro-organisms to site). In contrast to many other studies that concern the impact of single factors of the biotechnological clean-up of oil or the natural attenuation of oil pollutions in the sea, the experiments presented here focus on finding an applicable and practical solution to this grave environmental problem.

Emulsification and biofilm formation appeared in all nutrient-amended microcosms prior to the 14th day of the experiment, which led to an increase of the surface area between oil and water and hence improved bioavailability of the pollutant. This can be attributed to the activity of the well-known oil-degrading marine microbe A. borkumensis. Attachment to oil droplets has been described for this organism (Yakimov et al. 1998) and is a key issue for its lifestyle. It is attributed to the production of specific cell-bound glucose lipids (Abraham et al. 1998). GC of emulsified oil revealed that aliphatic linear and branched components of bunker C heavy fuel oil were degraded over a short period of time, whose rate was dependent on the initial microbial communities in the microcosms. The removal of heavy fuel oil was visible until the end of the experiment as photooxidation was excluded owing to the experimental design. Solubilization of components into the aquifer and evaporation of components (weathering) was simulated in a separate microcosm containing bunker C heavy fuel oil and sterile seawater. However, residual amounts of oil in microcosms containing micro-organisms were significantly lower in comparison with this control.

In parallel, observation of solubilized oxygen levels showed quick and intense respiration in pure cultures of A. borkumensis within sterile seawater (microcosms 5–7), which was in contrast to weaker but long-lasting effects in all other microcosms. Obviously, conditions in microcosms 5–7 led to the rapid metabolic activity of A. borkumensis and subsequent consumption of the hydrocarbon fraction that this organism is able to degrade. Conversely, microbial consortia in microcosms 19 and 25 took longer to establish, which explains the discrepancy described before. Increased respiration of microcosms 19 and 25 can also be explained by oxidation of a larger spectrum or quantity of hydrocarbons as can be seen in Fig. 1, underlining the efficiency of the consortia formed in both microcosms.

Composition of oil-degrading consortia

The majority of phylotypes detected in all excised DGGE bands were derived from the group of Alphaproteobacteria, which has commonly been observed in seawater sample studies (Eilers et al. 2000, 2001; Brakstad and Lodeng 2005). Most of these phylotypes are clustered around Roseobacter spp., yet a very common and prominent band showed a high degree of similarity (90%) to Thalassospira lucentensis, a member of Oceanospirillaceae, which is common in long-term seawater cultures (Lopez-Lopez et al. 2002). Phylotypes with high similarity to Ruegeria atlantica and Terasakiella pusillum were often detected in microcosms without nutrient supply. These did not display a large activity despite being spiked with oil. DGGE bands of these microcosms mostly contained phylotypes belonging to Alphaproteobacteria, except for a single phylotype identical with Microbacterium schleiferi (a member of Actinobacteria). It is probable that these organisms unspecifically grow owing to the high amounts of biomass and secondary metabolites or the excess of nutrients in the oil-depleting microcosms. Surprisingly, all nutrient-amended microcosm samples displayed a high diversity of DGGE bands that was accompanied by differing compositions of communities, i.e. a multitude of bands clustering within the Cytophaga-Flavobacterium group. Yet, it cannot be determined if this abundance is because of the greater amount of biomass in this microcosm, as the faint bands in this part of the gel could also be found in nonbiostimulated microcosms. Among the bands detected in nutrient-amended microcosms, Sulfitobacter pontiacus was identified. Sulfitobacter strains also were observed in a recent study (Brakstad and Lodeng 2005) and are considered common micro-organisms in North Sea waters. As mentioned previously, an unusual phylotype of spirochete was detected in microcosms containing exclusively X-Oil®. However, as the phylotype could not be cultured, its potential influence on oil degradation therefore is unknown. Very prominent and abundant within N/P-supplemented microcosms was the phylotype of A. borkumensis. As this micro-organism was detected in both nutrient-amended and nonsupplemented treatments, the appearance of A. borkumensis in the microcosms must be independent of biomass amendment (bioaugmentation).

As it has been observed in the RISA profiles (Fig. 2a–c), the addition of nutrients to an oil-spiked microcosm led to a dramatic reduction in the number of bands, thus showing a strong selective pressure in favour of few or a single species of micro-organism. The resulting bands were assigned to A. borkumensis suggesting its importance in these microcosms verified by the results of MDS in Fig. 3g, h.

Interestingly though, the DNA band characteristic for A. borkumensis at 458 bp was not only very distinct in samples containing low numbers of bands but also in those with a larger diversity of PCR amplicons. Considering Fig. 3e, f, which is a congruent display of the same MDS plot, it is clear that nutrient supply leads to strong selection in favour of A. borkumensis, although at the same time its presence was also detectable in nonsupplemented samples. A possible explanation for this could be the presence of strain(s) of Alcanivorax spp. in the initial seawater. Several Alcanivorax ssp. with differing nutrient requirements have been discovered recently (Roling et al. 2002) some showing a high consumption of nutrients, others using far lower amounts. On the one hand, the present results underline the importance of this organism in a given experimental setting though on the other, indicate their ability to outgrow competitors under in situ conditions, which has been reported previously (Hara et al. 2003).

Augmentation and its impact on microbial communities

Bioaugmentation is one of the most controversial issues of bioremediation. On the one hand, the formation of complex microbial consortia for efficient biodegradation of hydrocarbons and other hydrophobic pollutants is very important. Given that a single organism has no capability to degrade all constituents of oil, syntrophic interactions between members of the consortia are thought to be essential in this regard. On the other hand, the complex community undergoes certain changes of predominant organisms upon petroleum oil degradation process, for instance Alcanivorax and Cycloclasticus, which are known to utilize, correspondingly, aliphatic and aromatic hydrocarbons (for review, see Yakimov et al. 2007), suggesting that only a few of the community members perform the majority of the degradation process. Therefore, it stands to reason that increasing the population density of these key organisms to a high level might enhance the rate of oil biodegradation.

To prove this beneficial effect of bioaugmentation in experimental microcosms, RISA profiles of augmented microcosms were compared with the microcosms initially treated with plain seawater using MDS analysis; Fig. 3b, d show that the RISA profiles of the augmented samples are clustered around a blank control of a pure culture of A. borkumensis, in contrast to a rather widespread dispersion of the nonaugmented samples in Fig. 3a, c. Because of its unique features, A. borkumensis seems to be a primary colonizer of oil-contaminated marine environments, which makes it extremely valuable for biodegradation and bioaugmentation purposes.

Nevertheless, a pure culture of A. borkumensis applied in microcosm 7 showed poor results regarding degradation of hydrocarbons in Fig. 1. This may be a result of poor adaptation of the laboratory culture used for this experiment and might point to the practical problems of using such cultures in oil mitigation field work, as both marine seawater and augmented communities indicated a different behaviour concerning degradation of hydrocarbons.

Another potential explanation for this effect is the low catabolic diversity and thus functionality within a monoculture. As presented in several studies (Griffiths et al. 2000; Giller et al. 2004), the functionality, resilience and resistance to stressors of an ecosystem strongly depend on the diversity of its members. An ecosystem based on A. borkumensis alone analysed inter alia in this work is an excellent example for this theory, as this organism is strongly limited to a few metabolic pathways for linear and branched aliphatic hydrocarbons; however, it shall be remembered that other community members may facilitate the degradation of aromatic compounds and some minor oil constituents. As mentioned in a previous study (Loreau et al. 2001), high biodiversity represents a ‘buffer’ for fluctuations of environmental parameters and enhances the metabolic potential of an ecosystem. This may also explain the presence of numerous members in the microbial community although its composition is dominated by A. borkumensis. However, bioaugmentation of seawater microcosms led to an increase of biodegradation of aromatic compounds that are no substrates of A. borkumensis (Fig. 1). Given the genes for pathways (and even their parts) of aromatic hydrocarbons degradation are missing in the genome of A. borkumensis, a possible explanation for this could be the acceleration of the development of an oil-degrading consortium facilitated by A. borkumensis, as can be seen in Fig. 3a–h. Augmentation with a significant number of the nonmotile cells of A. borkumensis enhances the probability of cells and oil making contact. Thus, owing to the whole-cell- and biosurfactant-based emulsification the duration of the lag phase in the formation of the consortium is reduced.

Application of oil adsorbent material

Although different kinds of oil sorbent materials of both natural and synthetic origins already have been tested for their applicability in biodegradation and oil spill mitigation, little is known about their effects on or interactions with oil-degrading bacteria. In fact, because of the multitude of materials and their unique properties, each material being exposed to the marine environment has to be tested individually. Because of the textile origin of the product used in this study, interference of the compounds being leached out of the material was assumed, especially because of heavy metals like chromium (III), which is commonly applied in shoe or textile production.

Apart from this chemical interference, an influence of the material containing significant amounts of leather on microbial communities and the introduction of micro-organisms indigenous to X-Oil® was probable. Yet, isolation of genomic DNA from several grams of pure X-Oil® material failed because of extremely small quantities of biomass available on its surface. Influence of X-Oil® on microbial communities was examined with MDS but no significant effect on microbial communities (either positive or negative) was observed. Figure 3c shows no significant grouping of samples indicating an influence of X-Oil®. While in Fig. 3d, a clustering of samples is visible, the same observation can be made in Fig. 3b and therefore be ascribed to nutrient amendment. Surprisingly, results from DGGE indicated the growth of specific bacteria in microcosms treated with X-Oil®, i.e. a novel yet uncultured species of spirochete and members of Cytophaga–Flavobacterium group, which were also detected by Brakstad and Lodeng (2005) in a similar experiment. Most likely, the differences in communities promoted by X-Oil® are very subliminal and thus not detectable by RISA. The selective pressures of oil and nutrients in this experiment obviously were much stronger than the one resulting from the addition of the oil sorbent, which consists of inert or poorly degradable textiles.

As observed by phase-contrast microscopy, X-Oil® did bind free oil thus removing the pollutant from the environment. It also provided a matrix for biofilm and emulsion formation. According to the present data, X-Oil® has to be considered as a novel type of microbial habitat with unusual physical-chemical properties and should be investigated further. It was proven to be very useful for mechanical removal and absorption of oil in the seawater microcosms. Obviously, the role of an artificial support matrix in biodegradation is not fully understood. It can be assumed that oil-degrading microbial consortia forming in the water body and those forming biofilms on oil-spiked sorbents might differ in composition and activity.


Results of this study indicate that owing to its exceptional adaptation to oil-polluted marine environments and its strong dominance in case of adequate nutrient supply, A. borkumensis is a major organism initiating and mainly conducting the degradation of aliphatic hydrocarbons. As it is not able to degrade all components of the oil type used in the experiment, it obviously promotes the growth of microbes using further compounds, i.e. extracellular polysaccharides. Discussing the remarkable adaptation of A. borkumensis to oil degradation, this study underlines its importance and additionally highlights the necessity of its cooperation with other members of the microbial community in seawater, whose identity, metabolic capabilities and role for the in situ process of marine oil degradation have still to be analysed on a larger scale. An attempt of augmentation with A. borkumensis nevertheless demonstrated quicker growth of other marine microbes possibly because of the provision of better access to the substrate by oil emulsification. Therefore, it is foreseen that the application of A. borkumensis for this purpose would be beneficial. Using a novel type of sorbent material, no interference between sorbent and marine biodegradation was observed but the feasibility of combining bioaugmentation with this micro-organism and the oil sorbent used in this experiment can be assumed that will enable the construction of novel bioremediation tools for offshore oil mitigation.


The authors thank all staff of Helgoland Marine Science Station (BAH) who assisted in this work, especially Dr Antje Wichels and Hilke Döpke. They are especially grateful to Melanie Sapp for her kind assistance and help, essential for this study. This work was funded by GenoMik initiative of German Ministry for Science and Education (BMBF). K.N. Timmis acknowledges the Fonds der Chemischen Industrie for generous support. The authors also thank Joachim Hellmann for the kind provision of X-Oil® material and Mark C. Malpass for proofreading the manuscript.