Journal of Hazardous Materials 300 (2015) 75–83
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Marine biodegradation of crude oil in temperate and Arctic water samples Mette Kristensen a , Anders R. Johnsen b , Jan H. Christensen a,∗ a Analytical Chemistry, Section for Environmental Chemistry and Physics, Department of Plant and Environmental Sciences, University of Copenhagen, Thorvaldsensvej 40, 1871 Frederiksberg C, Denmark b Geological Survey of Denmark and Greenland (GEUS), Department of Geochemistry, Øster Voldgade 10, 1350 Copenhagen K, Denmark
h i g h l i g h t s
g r a p h i c a l
a b s t r a c t
• Biodegradation was assessed after exposure to 100 mg L−1 crude oil in microcosms. • The degradation order in Disko Bay samples were nalkanes > alkyltoluenes > PAHs. • The degradation order in North Sea samples were PAHs > alkyltoluenes > n-alkanes. • The differences in degradation orders will affect risk assessment in arctic regions
a r t i c l e
i n f o
Article history: Received 25 February 2015 Received in revised form 11 June 2015 Accepted 18 June 2015 Available online 24 June 2015 Keywords: Oil hydrocarbon fingerprint Polycyclic aromatic hydrocarbon Biodegradation Marine Arctic environment Gas chromatography-mass spectrometry TOC graphic
a b s t r a c t Despite increased interest in marine oil exploration in the Arctic, little is known about the fate of Arctic offshore oil pollution. Therefore, in the present study, we examine the oil degradation potential for an Arctic site (Disko Bay, Greenland) and discuss this in relation to a temperate site (North Sea, Denmark). Biodegradation was assessed following exposure to Oseberg Blend crude oil (100 mg L−1 ) in microcosms. Changes in oil hydrocarbon fingerprints of polycyclic aromatic hydrocarbons (PAHs), alkyl-substituted PAHs, dibenzothiophenes, n-alkanes and alkyltoluenes were measured by gas chromatography-mass spectrometry (GC–MS). In the Disko Bay sample, the degradation order was n-alkanes > alkyltoluenes (para- > meta- > ortho-isomers) > PAHs and dibenzothiophenes, whereas, the degradation order in the North Sea samples was PAHs and dibenzothiophenes > alkyltoluenes > n-alkanes. These differences in degradation patterns significantly affect the environmental risk of oil spills and emphasise the need to consider the specific environmental conditions when conducting risk assessments of Arctic oil pollution. © 2015 Published by Elsevier B.V.
1. Introduction
∗ Corresponding author. http://dx.doi.org/10.1016/j.jhazmat.2015.06.046 0304-3894/© 2015 Published by Elsevier B.V.
Focus on oil pollution is developing around the Arctic and other high-latitude environments due to climate change and intensified
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exploration for new oil resources. The Arctic regions differ from temperate regions in many ways, which may influence the fate of oil pollution. Low temperature, low light intensity [1,2], ice coverage [2] and lack of microbial adaptation [3,4] all indicate that oil pollution of the Arctic marine environment may be more persistent than oil pollution in temperate and tropic climates. In 2011, marine oil exploration started west of Greenland with 20 licensed areas covering 201.131 km2 [5]. Oil exploration and exploitation in this region is associated with an increased risk of accidental oil spills due to sea ice, drifting icebergs, and the depth at which the drillings take place (>350 m), but there is no knowledge on the fate of oil pollution in deeper parts of the water column in the Arctic. For the Macondo Deep Water Horizon blowout in the Gulf of Mexico, pre-spill adaptation of the microbial degrader community led to surprisingly fast oil degradation deep in the water column, probably caused by microbial adaptation from natural oil seeps [6]. Therefore, we wanted to test whether a similar adaptation was present in the marine water column at Disko Bay (Western Greenland). When investigating the fate of oil spills, unfortunately, it is common to analyse only for selected compounds in the complex mixture of crude oil or to quantify only the sum of total petroleum hydrocarbon. When focusing on a few target compounds, information on the entire mixture is limited. Many compounds in crude oil show large variations in physical-chemical properties and represent different classes that are important in different time aspects of an oil spill. The composition of n-alkanes, alkyltoluenes and monoaromatics changes rapidly in the initial phase, whereas, polycyclic aromatic hydrocarbons (PAHs), alkyl-substituted PAHs and NSO aromatics (nitrogen-, sulphur- and oxygen containing compounds) are more recalcitrant and may be present for years or decades [7]. Since the 1980s, chemical fingerprinting of oil hydrocarbons (‘oil hydrocarbon fingerprinting’) has been used for the identification of oil spill sources and for assessing weathering effects [8]. Chemical fingerprinting gives the relative composition of hundreds of compounds in a complex mixture by using specific analytical methods. A general trend in oil hydrocarbon biodegradation is the degradation order of n-alkanes > monoaromatic compounds (including benzene, toluene, ethylbenzene and xylenes known as BTEX) > branched and cyclic alkanes > polycyclic aromatic compounds (decreased degradation with increased number of aromatic rings), and a negative correlation between degradation and degree of alkyl substitution [8–10]. n-alkyltoluenes are not commonly included in the chemical fingerprinting of oil spills, but have been suggested as a complementary analysis to n-alkanes in the preliminary screening of oil pollution [11]. The analysis of alkyltoluenes can provide additional information about biodegradation rates and patterns in the initial stage of an oil spill, as well as in source identification. The aim of our study was to evaluate the natural potential for biodegradation of crude oil in subsurface Arctic water, presumably with limited microbial adaptation to oil hydrocarbons. Biodegradation was assessed by analysing n-alkanes, alkyltoluenes, PAHs and dibenzothiophenes to give a complex fingerprint of the crude oil changes. Additionally, the degradation potential of a typical shallow, temperate environment with oil production and shipping was evaluated to emphasise the significant differences in degradation due to environmental conditions. Diagnostic ratios are a commonly used method in oil spill assessments to distinguish between degradation processes. The method is robust and insensitive to changes in extraction efficiency or analysis. Diagnostic biodegradation ratios, such as nC17 /pristane, nC18 /phytane, PAH isomers or other compounds with the same physical-chemical properties but different susceptibilities to microbial attack, are used as markers of biodegradation,
and to identify sources of hydrocarbon pollution [12–14]. A change in diagnostic ratios is evidence for biodegradation, which is especially important in subsurface spills where biodegradation is the only true degradation process. To examine the biodegradation potential of the two environments under laboratory conditions, a microcosm study was set up with the addition of typical North Sea crude oil (Oseberg Blend) to subsurface water from the Disko Bay, Greenland (Arctic) and to surface water from the North Sea, Denmark (temperate) in 1 L microcosms. The degradation of crude oil and changes in the microbial degrader communities was followed over a 10-week incubation period. 2. Materials and methods 2.1. Chemicals: The North Sea crude oil, Oseberg Blend (Statoil, Norway), was used for oil incubations For solid phase extraction and gas chromatography - mass spectrometry (GC–MS), methanol (HPLC grade, Rathburn), npentane (HPLC grade, VWR Chemicals), acetone (HPLC grade, Rathburn), dichloromethane (HPLC grade, Rathburn), ammonia solution (28–30%, MERCK) and ammonium acetate (≥98%, MERCK) were used. For instrument performance assessment, a GC–MS tune mixture was used. This was composed of benzidine (99.9%, Sigma), 1,1,1-Trichloro-2,2-bis(4-chlorophenyl) ethane (99.7%, Sigma), pentachlorophenol (99.9%, Sigma, Supelco) and decafluorotriphenylphosphine (99.3%, Sigma, Supelco) (50 g mL−1 of each). For further quality assurance and quality control, a reference sample composed of 19 PAHs with 2–6 aromatic rings and 15 deuterated PAHs with 2–6 aromatic rings was analysed between samples. Internal standard and recovery standard solutions consisted of mixtures of deuterated PAHs and nitrogen- and oxygen-containing polycyclic aromatic compounds in methanol. The internal standard stock solution contained a mixture of d8 -naphthalene (13.42 g mL−1 , 99%), d8 -dibenzothiophene (8.82 g mL−1 , 98%), d10 -acenaphthene (7.96 g mL−1 , 98%), d10 phenanthrene (13.05 g mL−1 , 98%), d10 -pyrene (8.27 g mL−1 , 98%) and d10 -fluorene (7.68 g mL−1 , 98%) from the Cambridge Isotope Lab and d9 -acridine (12.50 g mL−1 , 98.7%, CHIRON AS), d8 -carbazole (12.50 g mL−1 , 98.9%, CHIRON AS), d8 -antraquinonone (9.05 g mL−1 ) and d5 -phenol (13.78 g mL−1 , 98%, Sigma–Aldrich). The recovery standard stock solution contained d10 -anthracene (7.07 g mL−1 , 98%, Dr. Ehrenstorfer Gmb), and d8 -acenaphthylene (7.76 g mL−1 , 98%), d10 -fluoranthene (6.91 g mL−1 , 98%), d12 -benzo[a]anthracene (7.12 g mL−1 , 98%), d12 -benzo[a]pyrene (8.40 g mL−1 , 98%) and d12 -indenol(1,2,3–c and d) pyrene (6.84 g mL−1 , 98%) from the Cambridge Isotope Lab, and hydrochloric acid (37%, VWR Chemicals). Substrates used for Most-Probable-Number enumeration (MPN) were hexadecane (99%, Sigma–Aldrich), m-xylene (>99%, Fluka), 2-methylnaphthalene (97%, Sigma–Aldrich), 1-naphtol (≥99%, Sigma–Aldrich), 1-naphthoic acid (98%, Alfa Aesar) and silicone oil AR20 (Sigma–Aldrich). Dilutions were made in pre-mixed Bushnell–Haas minimal mediums containing magnesium sulphate (0.2 g L−1 ), calcium chloride (0.02 g L−1 ), monopotassium dihydrogen phosphate (1.0 g L−1 ), diammonium hydrogen phosphate (1.0 g L−1 ), potassium nitrate (1.0 g L−1 ) and ferric chloride (0.05 g L−1 ) (DifcoTM , Becton, Dickinson and Company, Sparks, MD21152, USA) [15]. 2.2. Sampling Two locations, the North Sea and the Disko Bay, represented typical temperate and Arctic marine environments. The North Sea
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is influenced by oil exploration and extensive shipping traffic, whereas, the Disko Bay in Western Greenland is a pristine area of great interest for future oil exploration and exploitation [16]. Sampling of the North Sea was carried out on the 6th of July 2013 from a small fishing boat. The sample was collected from a depth of 1.5 m 1.8 km north–west of Hirtshals Harbor (northern Denmark) at the position 57◦ 36 18“N, 09◦ 56 14“E. The sample was stored for four days at 5 ◦ C until use. Sampling of the Disko Bay was carried out on June 29th, 2013. Approximately 50 L of water was collected from a depth of 150 m using a 10 L Niskin water sampler (KC Denmark A/S, Silkeborg, Denmark) at a position 5 km south of Qeqertarsuaq at 69◦ 11 16“N, 53◦ 31 06“W. The Disko Bay sample was stored at 5 ◦ C for 11 days until initiation of the microcosm experiment. The temperature of both samples was continuously logged during transportation. The North Sea sample never exceeded 20 ◦ C, and the Arctic sample never exceeded 10 ◦ C. 2.3. Microcosm set-up Subsamples of 750 mL with 100 mg L−1 of Oseberg Blend crude oil were incubated for 10 weeks in 1 L glass flasks at 15 ◦ C (North Sea subsamples) or 5 ◦ C (Disko Bay subsamples). These temperatures represented typical water temperatures at the sampling times. Triplicate microcosms were sacrificed after 0, 4, 8, 15, 36 and 71 days of incubation. The microbial degrader community was analysed for all these incubation times, whereas the composition of the oil phase was analysed only for days 0, 15 and 71. The day 0 samples were sacrificed after nine hours of incubation to ensure equilibrium between the oil and water phase. A parallel series of autoclaved controls, with an addition of 100 mg L−1 Oseberg Blend, were used to estimate the physical removal processes. A series of controls without oil were used to determine the inherent changes in the microbial communities that were not caused by oil. For the first 24 h of incubation, the flasks were sealed with Teflon-lined lids to avoid evaporation and allow time for phase equilibration of the oil compounds between the oil phase and the water phase. The lids were then removed, and the microcosms were covered with oxygen- and carbon dioxide-permeable low-density polyethylene film (cling film). During incubation, the microcosms were shaken for five minutes at 120 rev min−1 approximately every 48 h to facilitate aeration. The microcosms were incubated in the dark to avoid photo-transformation of the oil compounds. Subsamples of the water phase were collected at each sampling day for MPN enumeration of hydrocarbon-degrading microbes. The microcosms were then acidified with 10 mL of 1 M hydrochloric acid to stop further biodegradation and left overnight for separation of the oil- and water phases before chemical analysis of the oil phase. 2.4. MPN enumeration of hydrocarbon-degrading microorganisms The size of microbial subpopulations that could utilise specific hydrocarbons as the sole source of carbon and energy were estimated with a modified version of a previously published microplate method [17] where, hydrocarbon growth substrates are added to the microplate wells in a separate silicone phase (silicone oil AR20, Sigma–Aldrich) to avoid toxic effects. The concentrations in the silicone phase were as follows: 10% hexadecane, 2% 2-methylnaphthalene, 1% m-xylene, 0.5% 1-naphtol and 0.5% 1naphthoic acid. The substrates were pasteurised for five minutes in a 70 ◦ C water bath in tightly closed glass scintillation vials with aluliners in the lids. A half strength Bushnell–Haas minimal medium [15] was supplemented with 30 g L−1 of NaCl (BH-medium) and autoclaved for 20 min at 121 ◦ C in tightly closed red-cap bottles. The precipitate was removed after autoclavation by decanting into sterile blue-cap bottles. For MPN enumerations, 1 mL subsamples were
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sampled from each microcosm. Subsamples from triplicate microcosms were pooled to give one mixed sample for each treatment at each time point. Ten-fold dilution series’ of the mixed samples were prepared in the BH-medium. From each dilution (10−1 –10−8 ), four times 200 L were added to six microtiter plates. Then, 15 L of a hydrocarbon substrate solution was added to each microplate well (one substrate per microplate), a microplate lid was put on and the microplates were placed in airtight plastic boxes. A small vial with extra substrate solution (1 mL) was placed on top of the plates to compensate for evaporation. Plates with sample dilutions and silicone oil, but no hydrocarbons, served as negative controls. The plates were incubated at 10 ◦ C in the dark. A vial with fresh substrate was added every three weeks. After 10 weeks of incubation, the absorbance of each microplate well was read at 450 nm on a ThermoMax microplate reader (Molecular Devices). Wells were scored growth-positive when they had an absorbance higher than 0.1, and MPN estimates were calculated according to Hurley and Roscoe [18] from the distribution of growth-positive and growthnegative microplate wells. The detection limit was calculated as one growth-positive well in the lowest dilution, giving a LogMPN of 1.1. 2.5. Chemical analysis Five hundred and fifty millilitres of the water phase was removed, and the residual oil phase was extracted two times with 5 mL dichloromethane. To all extracts, 200 L internal standard stock solution was added, samples were volume adjusted to 5 mL by evaporation in a fume hood at room temperature and 200 L recovery standard was added. Oil phase samples were diluted tenfold to give an adequate signal for GC–MS analysis of approximately 2000 mg oil L−1 solvent. The oil composition was analysed by gas chromatography (HP-7890A, Agilent Technology) with a 60 m ZB-5 capillary column (25 mm inner diameter and 0.25 m film thickness) interfaced to a quadrupole mass spectrometer (HP-5975C, Agilent Technology). Measurements were done by selected ion monitoring (SIM) of 55 ions divided into 12 groups using electron ionisation. One L sample was injected by pulsed splitless injection at 315 ◦ C and helium was used as a carrier gas with a flow rate of 1.1 mL s−1 . Oven temperature was ramped from initial 40 ◦ C (hold 2 min) to 100 ◦ C by 25 ◦ C s−1 followed by 5 ◦ C s−1 up to 315 ◦ C (hold 13.4 min). The method is described in detail in Gallotta and Christensen (2012) [19]. 2.5.1. Quality assurance and quality control Oil phase extracts were analysed in three batches of 23 vials, including samples and test solutions. Test solutions of dichloromethane, reference samples (with a known mixture of PAHs and deuterated PAHs), mixture samples (a mixture of thirteen samples) and a GC–MS tune mixture were analysed between samples and batches. These solutions were used for quality control of analysis, e.g. daily variations, cross contamination, peak shape, chromatographic resolution and sensitivity, and for verification of tuning, injection port and column performance [19]. 2.5.2. Data processing The data consisted of GC–MS SIM chromatograms of 44 oil phase extracts (three sampling days, triplicates, two locations, two exposures (autoclaved and oil exposed) and eight 0 mg L−1 microcosms as biological controls). The compound peak area was determined by manual integration according to the European Committee for Standardization, CEN oil spill identification guideline (CEN/TR 15,522-2:2009, www.cen.eu) in Agilent MSD Productivity ChemStation (E.02.02.1431). All calculations were made in Excel 2007 (Microsoft Office).
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Fig. 1. Most probable number enumeration (MPN, cells mL−1 ) of specific hydrocarbon degrader sub-populations (n = 1 ± one standard error).
3. Results 3.1. MPN enumeration of hydrocarbon-degrading microorganisms MPN enumerations were carried out for five growth substrates (Fig. 1) representing n-alkane degraders (hexadecane), monoaromatic degraders (m-xylene), diaromatic degraders (2methylnaphthalene) and polar aromatic degraders (1-naphthol
and 1-naphthoic acid). The MPNs gave several unexpected results. First, we did not detect any cultivable degraders able to grow with 1-naphthol or 1-naphthoic acid as the sole source of carbon and energy. The absence of degraders was especially unexpected for 1-naphthoic acid, which is an intermediate in the microbial metabolisation of 1-methylnaphthalenes [20]. Second, it was striking that the addition of crude oil did not result in massive increases of cultivable hydrocarbon-degrader MPNs. It was especially interesting that hexadecane degraders, which were found in
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Fig. 2. nC8 -nC32 alkanes in samples (triangles) after 15 (filled symbols) and 71 (open symbols) days of incubation, normalised to 17␣,21-hopane and compared to day 0 (n = 3 ± one relative standard deviation). Additionally, n-alkanes in the corresponding autoclaved controls are depicted (diamonds).
high initial numbers in the North Sea sample, increased less than 10-fold in microcosms spiked with crude oil and soon dropped to the same level as in the unspiked control microcosms. The most significant change, induced by the addition of crude oil, was the increase in 2-methylnaphthalene degrader MPNs in both the North Sea and the Disko Bay microcosms. Degraders of hexadecane, m-xylene and 2-methylnaphthalene were present in high numbers already at day zero in the North Sea samples, whereas, in the Disko Bay samples, m-xylene degraders and especially 2methylnaphthalene degraders were detectable after enrichment for 15 and 36 days, respectively. This might indicate that the microbial degrader community from the North Sea was more preadapted to oil degradation than the Arctic microbial community from Disko Bay. 3.2. n-alkanes and alkyltoluenes Changes in nC8 -nC32 alkanes compared to day 0 are shown in Fig. 2. The data was normalised to 17␣,21-hopane because this compound is highly recalcitrant in the environment and largely unaffected by biological degradation [12]. Surprisingly, only shortchain n-alkanes were significantly affected in samples from the North Sea (Fig. 2). After 71 days of incubation, nC8 –nC14 alkanes were reduced to 0.1–75.0% of the initial values, whereas, alkanes with chain lengths above nC14 remained almost unaffected with recoveries of 90.4–104.5% at day 71. According to the United States Environmental Protection Agency definition, nC8 -nC14 alkanes are classified as volatile organic compounds (VOCs) (BP 50-260 ◦ C), nC15 –nC24 alkanes are classified as semi-volatile organic compounds (SVOCs) (BP 240–400 ◦ C) and longer n-alkanes are classified as non-volatile (non-VOCs) (www.epa.gov). From this definition, it is clear that only volatile alkanes up to nC14 were removed by
evaporation in the North Sea samples, both in the intact samples and in the autoclaved controls. For the Disko Bay samples, we observed a much larger effect on the n-alkanes (Fig. 2). The amounts of VOCs were reduced to 0.0-4.4%, SVOCs to 5.5–42.9% and non-VOCs to 59.7–102.4% compared to day 0. In the autoclaved controls, only the VOCs were affected (0.0–88.3% after 71 days compared to day 0). The large removal of semi- and non-volatile n-alkanes, compared to the autoclaved controls, demonstrated that n-alkane biodegradation was a key factor in oil degradation only in the Disko Bay samples. Additionally, SVOCs were not expected to show large removal from evaporation in the Disko Bay samples, which were incubated at lower temperatures than the North Sea samples. The diagnostic ratios of nC17 /pristane and nC18 /phytane were calculated to confirm biodegradation. Fig. 3 shows the changes in diagnostic ratios, normalised to autoclaved controls, when expressed relative to the day 0 ratios. No changes in the diagnostic alkane biodegradation ratios were seen in the autoclaved controls for the North Sea samples or the Disko Bay samples (99.9–100.9% compared to day 0, data shown in Supplementary information). This demonstrated that physical removal processes, e.g. evaporation, as assumed, were negligible with respect to these biodegradation ratios. Therefore, the decrease in nC17 /pristane and nC18 /phytane in the intact samples was solely caused by alkane biodegradation. In samples from the North Sea, alkane biodegradation ratios decreased only slightly (to 93.6 ± 3.3% of their initial values for nC17 /pristane and to 96.3 ± 3.1% for nC18 /phytane). This confirms the lack of n-alkane biodegradation in the North Sea water seen in Fig. 2. For the Disko Bay samples, both nC17 /pristane and nC18 /phytane were reduced significantly (to 9.2 ± 8.0% and 9.2 ± 6.8%, respectively), verifying that biodegradation was the only significant process in the removal of n-alkanes in the Disko Bay water.
Fig. 3. Diagnostic ratios of nC17 /pristane and nC18 /phytane normalised to the corresponding autoclaved controls and compared to day 0 (n = 3 ± one relative standard deviation).
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Table 1 Boiling point, octanol-water partitioning coefficient (LogKow ), retention time and chemical formula for alkyltoluenes (group I, II and II) and their corresponding n-alkanes. BP, ◦ C
LogKow
RT, min
Chemical formula
nC16 Group I alkyltoluene
286.4 296.4
8.2 7.0
23.6 22.8–23.3
C16 H34 C16 H26
nC17 Group II alkyltoluene
302.0 310.9
8.7 7.5
25.8 25.2–25.6
C17 H36 C17 H28
nC18 Group III alkyltoluene
316.3 324.4
9.2 8.0
27.9 27.4–27.9
C18 H38 C18 H30
Alkyltoluene structure (ortho-isomer)
Calculated in EPIWEB 4.1. Ortho substituted alkyltoluenes are used for calculations.
The initial loss of low molecular weight alkyltoluenes is comparable to that of n-alkanes. Three groups of meta-, para- and ortho-alkyltoluenes were chosen with retention times corresponding to nC16 (group I), nC17 (group II) and nC18 alkanes (group III), and thereby, were estimated to be of equivalent boiling points to these n-alkanes (Table 1). In Fig. 4, the three groups of alkyltoluenes are presented as compared to day 0. All data were normalised to 17␣,21-hopane and autoclaved controls. In the North Sea samples (Fig. 4), there were no changes in the alkyltoluenes at day 15 or 71 (98.1–175.8% compared to day 0). This was in accordance with results for n-alkanes that were removed only by evaporation of the short-chain alkanes up to nC14 . In samples from the Disko Bay, we detected the removal of alkyltoluenes for both days 15 and 71. The group I alkyltoluenes showed 23.3 ± 27.2%, 48.6 ± 29.6% and 102.4 ± 8.9% of initial values after 71 days of incubation for para-, meta- and ortho-alkyltoluenes, respectively. For group II and III alkyltoluenes, the isomers varied between 46.0 and 103.5%. nC16 to nC18 alkanes in the Disko Bay microcosms were reduced to 7.1–9.7% of their initial values at day 71. The degree of n-alkane removal in the Disko Bay water was, hence, much higher than that observed for alkyltoluenes of corresponding alkane chain lengths. Furthermore, the alkyltoluenes were removed in the isomer specific order of para- > meta- » ortho-alkyltoluene, which is in accordance with previous studies showing lower biodegradation of the ortho-isomer of xylene [21].
3.3. Polycyclic aromatic hydrocarbons (PAHs) and dibenzothiophenes Changes in naphthalene, phenanthrene, fluorene, pyrene, chrysene and the sulphur-containing dibenzothiophene, together with their alkyl-substituted homologues (C0 –C4 ) were measured by oil hydrocarbon fingerprinting on GC–MS. The percentage of the five PAHs, dibenzothiophene and their alkyl-substituted homologues is shown in Fig. 5. All data was normalised to 17␣,21-hopane and expressed relative to the equivalent autoclaved controls, thereby excluding effects from physical weathering processes. In the North Sea microcosms, we found biodegradation at day 71 of C0 - – C4 -naphthalenes (0.0–66.5% compared to autoclaved controls), C0 - – C2 -dibenzothiophenes (3.4–80.1%), C0 - – C2 -phenanthrenes (4.2–77.6%), C0 - – C3 -fluorenes (6.5–85.3%) and C0 -pyrenes (85.8 ± 7.0%). Neither C1 - – C3 - chrysenes, C1 - – C2 pyrenes, C3 - – C4 -phenanthrenes nor C3 - – C4 -dibenzothiophenes were biodegraded. The non-substituted PAHs and dibenzothiophene were degraded to the largest extent for all compound groups, showing that the degradation was negatively correlated with alkyl substitution. Additionally, the biodegradation of PAHs was decreased as the number of aromatic rings increased. In the Disko Bay samples, only C0 - and C1 -naphthalenes showed evidence of biodegradation by decreased amounts at day 71 (54.0 ± 13.7% and 64.5 ± 8.0%, respectively). From the MPN esti-
Fig. 4. Percent of alkyltoluenes (alkylT) normalised to 17␣,21-hopane and the corresponding autoclaved controls compared to day 0 (n = 3 ± one relative standard deviation). Group I alkyltoluenes have retention times corresponding to nC16 -, group II to nC17 - and group III to nC18 alkanes.
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Fig. 5. PAHs, dibenzothiophenes and the alkyl-substituted homologues normalised to 17␣,21-hopane and compared to the corresponding autoclaved controls (n = 3 ± one relative standard deviation). The North Sea samples are depicted in the left part of the graphs, while the Disko Bay samples are depicted to the right.
mates, it was clear that microorganisms capable of degrading more metabolically complex hydrocarbons, in this case, expressed by growth on m-xylene and 2-methylnaphthalene, were at low levels in the Disko Bay microcosms compared to the North Sea microcosms, even after 71 days of incubation (Fig. 1). The low content of cultivable degrader microorganisms might explain the lack of degradation of the metabolically complex hydrocarbons with more than two aromatic rings and with alkyl substitution.
We also calculated diagnostic biodegradation ratios for the PAHs and the dibenzothiophenes (Fig. 6). In the North Sea samples, all ratios indicated biodegradation of PAHs. The diagnostic ratios for the dibenzothiophenes remained almost constant after 71 days of incubation, with values of 108.4 ± 3.8% for 2 + 3-methyldibenzothiophene/4-methyldibenzothiophene and 91.1 ± 9.8% for 1-methyldibenzothiophene/4methyldibenzothiophene compared to day 0 (100%). The
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Fig. 6. Diagnostic ratios of PAHs and dibenzothiophenes normalised to corresponding autoclaved controls and compared to day 0 for North Sea and Disko Bay samples (n = 3 ± one relative standard deviation). mN: methylnaphthalenes, mF: methylfluorenes, mP: methylphenanthrenes and mDBT: methyldibenzothiophenes.
diagnostic ratios of alkyl-substituted fluorenes showed the largest responses with decreases to 63.0 ± 5.9% for 2 + 3methylfluorene/4-methylfluorene and to 57.0 ± 10.7% for 1-methylfluorene/4-methylfluorene at day 71. For day 71, it was not possible to calculate a reliable estimate of the 2methylnaphthalene/1-methylnaphthalene diagnostic ratio since both isomers were almost completely removed. In samples from the Disko Bay, only the diagnostic ratio of 2methylnaphthalene/1-methylnaphthalene decreased (56.6 ± 2.2% at day 71), indicating degradation of the two-ring PAHs. The diagnostic ratios of the three-ring PAHs (fluorenes and phenanthrenes) and the dibenzothiophenes remained constant during the incubation (99.4–104.7% at day 71 compared to day 0). The lack of changes in the diagnostic ratios for three-ring PAHs and dibenzothiophenes proved the limited potential for biodegradation of metabolically complex aromatics in the Disko Bay sample. Autoclaved controls showed no changes in the diagnostic ratios, except for 2-methylnaphthalene/1-methylnaphthalene where the autoclaved controls were reduced to 72.2 ± 2.1% for the North Sea samples and 86.0 ± 0.7% at day 71 for the Disko Bay samples (data shown in Supplementary information). This change, however, seems to be by physical–chemical processes rather than biological degradation, since we did not detect any microbial activity in the autoclaved controls as determined by [3 H]-Leucine incorporation at days 0, 15 and 71 (unpublished results).
environments, the degree of biodegradation decreased with alkyl substitution, which has also been found in several previous studies of crude oil biodegradation [9,10,22]. Water samples from the North Sea showed a clear potential for biodegradation of the metabolically complex compounds, whereas the subsurface samples from the Arctic Disko Bay environment only showed the degradation of simple hydrocarbons like the n-alkanes. Extensive biodegradation of PAHs and dibenzothiophenes compared to n-alkanes, as seen in the North Sea samples, is the opposite of the expected degradation order, but other studies have indicated a similar pattern [22,23]. A recent study of oil degradation in Arctic sea-water indicated the substantial loss of oil at −1 ◦ C with significant biodegradation of 2- – 4-ring PAHs and their alkyl-substituted homologues [24]. These results are different from ours, but it was not indicated whether sterile controls were included in their study. Abiotic removal processes might therefore have caused the different degradation patterns. It may be problematic to directly extrapolate from our limited microcosm study to a real Arctic subsurface spill, but our results indicate that aromatic hydrocarbons with more than two rings may show high persistence in Arctic subsurface waters. These persistent compounds may detrimentally affect marine organisms and Arctic marine ecosystems, and ought to influence future oil spill responses in the Arctic. Our results indicate that risk assessment data from the shallow North Sea is not relevant to subsurface spills in the Disko Bay area, and probably also of little relevance to other pristine places in the marine Arctic.
4. Discussion In the present study, we compared the potential for microbial oil degradation in water from the Disko Bay and the North Sea. The sampling depth of 150 meters in Disko Bay was chosen to study the oil spill response of a subsurface Arctic environment. Surface samples from the North Sea, on the other hand, were chosen for comparison because this environment is well characterized with respect to oil degradation. Lessons from this environment might therefore be applied for oil spill risk assessment of Arctic environments, where, data are scarce. Our results, however, demonstrated a completely different pattern of crude oil degradation in the two environments. The degradation order in the North Sea samples was PAHs and dibenzothiophenes (naphthalenes > dibenzothiophenes > phenanthrenes > fluorenes) > alkyltoluene > n-alkanes, whereas, in the Disko Bay samples, the order was reversed: n-alkanes > alkyltoluenes (para> meta- > ortho-isomer) > PAHs and dibenzothiophenes. In both
Acknowledgements The authors would like to thank the University of Copenhagen for funding this project and Katrine Scheibye for collaboration on the topic of oil degradation in Arctic environments. Peter Jensen is acknowledged for help during field sampling at Hirtshals, Denmark, as is the staff at Arctic Station, University of Copenhagen for their assistance during field sampling at Disko Bay, Greenland. This study was funded by the European Commission (KILLSPILL project, grant agreement 312139). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jhazmat.2015.06. 046
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[12] Z.D. Wang, M. Fingas, D.S. Page, Oil spill identification, J. Chromatogr. A 843 (1999) 369–411. [13] J.H. Christensen, G. Tomasi, Practical aspects of chemometrics for oil spill fingerprinting, J. Chromatogr. A 1169 (2007) 1–22. [14] M. Tobiszewski, J. Namiesnik, PAH diagnostic ratios for the identification of pollution emission sources, Environ. Pollut. 162 (2012) 110–119. [15] L.D. Bushnell, H. Haas, F.: The utilization of certain hydrocarbons by microorganisms, J. Bacteriol. 41 (1941) 653–673. [16] A. Mosbech, D. Boertmann, M. Jespersen, Strategic Environmental Impact Assessment of hydrocarbon activities in the Disko West area, NERI Tech. Rep. (2007). [17] A.R. Johnsen, S. Henriksen, Microplate MPN-enumeration of monocyclic- and dicyclic-aromatic hydrocarbon degraders via substrate phase-partitioning, Biodegradation 20 (2009) 581–589. [18] M.A. Hurley, M.E. Roscoe, Automated statistical-analysis of microbial enumeration by dilution series, J. Appl. Bacteriol. 55 (1983) 159–164. [19] F.D.C. Gallotta, J.H. Christensen, Source identification of petroleum hydrocarbons in soil and sediments from Iguacu River Watershed, Parana, Brazil using the CHEMSIC method (CHEMometric analysis of Selected Ion Chromatograms), J. Chromatogr. A 1235 (2012) 149–158. [20] M.C. Mahajan, P.S. Phale, C.S. Vaidyanathan, Evidence for the involvement of multiple pathways in the biodegradation of 1- and 2-methylnaphthalene by Pseudomonas putida CSV86, Arch. Microbiol. 161 (1994) 425–433. [21] N. Jimenez, M. Vinas, J. Sabate, S. Diez, J.M. Bayona, A.M. Solanas, J. Albaiges, The Prestige oil spill. 2. Enhanced biodegradation of a heavy fuel oil under field conditions by the use of an oleophilic fertilizer, Environ. Sci. Technol. 40 (2006) 2578–2585. [22] N.M. Fayad, Overton, E, A unique biodegradation pattern of the oil spilled during the 1991 Gulf War, Mar. Pollut. Bull. 30 (1995) 239–246. [23] D.M. Jones, A.G. Douglas, R.J. Parkes, J. Taylor, W. Giger, C. Schaffner, The recognition of biodegraded petroleum derived aromatic hydrocarbons in recent marine sediments, Mar. Pollut. Bull. 14 (1983) 103–108. [24] K. McFarlin, M. Prince, R.C. Perkins, R. Leigh, MB, Biodegradation of Dispersed Oil in Arctic Seawater at-1 degrees, C, PLoS One (2014) 9.
Contents lists available at ScienceDirect
Journal of Hazardous Materials journal homepage: www.elsevier.com/locate/jhazmat
Marine biodegradation of crude oil in temperate and Arctic water samples Mette Kristensen a , Anders R. Johnsen b , Jan H. Christensen a,∗ a Analytical Chemistry, Section for Environmental Chemistry and Physics, Department of Plant and Environmental Sciences, University of Copenhagen, Thorvaldsensvej 40, 1871 Frederiksberg C, Denmark b Geological Survey of Denmark and Greenland (GEUS), Department of Geochemistry, Øster Voldgade 10, 1350 Copenhagen K, Denmark
h i g h l i g h t s
g r a p h i c a l
a b s t r a c t
• Biodegradation was assessed after exposure to 100 mg L−1 crude oil in microcosms. • The degradation order in Disko Bay samples were nalkanes > alkyltoluenes > PAHs. • The degradation order in North Sea samples were PAHs > alkyltoluenes > n-alkanes. • The differences in degradation orders will affect risk assessment in arctic regions
a r t i c l e
i n f o
Article history: Received 25 February 2015 Received in revised form 11 June 2015 Accepted 18 June 2015 Available online 24 June 2015 Keywords: Oil hydrocarbon fingerprint Polycyclic aromatic hydrocarbon Biodegradation Marine Arctic environment Gas chromatography-mass spectrometry TOC graphic
a b s t r a c t Despite increased interest in marine oil exploration in the Arctic, little is known about the fate of Arctic offshore oil pollution. Therefore, in the present study, we examine the oil degradation potential for an Arctic site (Disko Bay, Greenland) and discuss this in relation to a temperate site (North Sea, Denmark). Biodegradation was assessed following exposure to Oseberg Blend crude oil (100 mg L−1 ) in microcosms. Changes in oil hydrocarbon fingerprints of polycyclic aromatic hydrocarbons (PAHs), alkyl-substituted PAHs, dibenzothiophenes, n-alkanes and alkyltoluenes were measured by gas chromatography-mass spectrometry (GC–MS). In the Disko Bay sample, the degradation order was n-alkanes > alkyltoluenes (para- > meta- > ortho-isomers) > PAHs and dibenzothiophenes, whereas, the degradation order in the North Sea samples was PAHs and dibenzothiophenes > alkyltoluenes > n-alkanes. These differences in degradation patterns significantly affect the environmental risk of oil spills and emphasise the need to consider the specific environmental conditions when conducting risk assessments of Arctic oil pollution. © 2015 Published by Elsevier B.V.
1. Introduction
∗ Corresponding author. http://dx.doi.org/10.1016/j.jhazmat.2015.06.046 0304-3894/© 2015 Published by Elsevier B.V.
Focus on oil pollution is developing around the Arctic and other high-latitude environments due to climate change and intensified
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exploration for new oil resources. The Arctic regions differ from temperate regions in many ways, which may influence the fate of oil pollution. Low temperature, low light intensity [1,2], ice coverage [2] and lack of microbial adaptation [3,4] all indicate that oil pollution of the Arctic marine environment may be more persistent than oil pollution in temperate and tropic climates. In 2011, marine oil exploration started west of Greenland with 20 licensed areas covering 201.131 km2 [5]. Oil exploration and exploitation in this region is associated with an increased risk of accidental oil spills due to sea ice, drifting icebergs, and the depth at which the drillings take place (>350 m), but there is no knowledge on the fate of oil pollution in deeper parts of the water column in the Arctic. For the Macondo Deep Water Horizon blowout in the Gulf of Mexico, pre-spill adaptation of the microbial degrader community led to surprisingly fast oil degradation deep in the water column, probably caused by microbial adaptation from natural oil seeps [6]. Therefore, we wanted to test whether a similar adaptation was present in the marine water column at Disko Bay (Western Greenland). When investigating the fate of oil spills, unfortunately, it is common to analyse only for selected compounds in the complex mixture of crude oil or to quantify only the sum of total petroleum hydrocarbon. When focusing on a few target compounds, information on the entire mixture is limited. Many compounds in crude oil show large variations in physical-chemical properties and represent different classes that are important in different time aspects of an oil spill. The composition of n-alkanes, alkyltoluenes and monoaromatics changes rapidly in the initial phase, whereas, polycyclic aromatic hydrocarbons (PAHs), alkyl-substituted PAHs and NSO aromatics (nitrogen-, sulphur- and oxygen containing compounds) are more recalcitrant and may be present for years or decades [7]. Since the 1980s, chemical fingerprinting of oil hydrocarbons (‘oil hydrocarbon fingerprinting’) has been used for the identification of oil spill sources and for assessing weathering effects [8]. Chemical fingerprinting gives the relative composition of hundreds of compounds in a complex mixture by using specific analytical methods. A general trend in oil hydrocarbon biodegradation is the degradation order of n-alkanes > monoaromatic compounds (including benzene, toluene, ethylbenzene and xylenes known as BTEX) > branched and cyclic alkanes > polycyclic aromatic compounds (decreased degradation with increased number of aromatic rings), and a negative correlation between degradation and degree of alkyl substitution [8–10]. n-alkyltoluenes are not commonly included in the chemical fingerprinting of oil spills, but have been suggested as a complementary analysis to n-alkanes in the preliminary screening of oil pollution [11]. The analysis of alkyltoluenes can provide additional information about biodegradation rates and patterns in the initial stage of an oil spill, as well as in source identification. The aim of our study was to evaluate the natural potential for biodegradation of crude oil in subsurface Arctic water, presumably with limited microbial adaptation to oil hydrocarbons. Biodegradation was assessed by analysing n-alkanes, alkyltoluenes, PAHs and dibenzothiophenes to give a complex fingerprint of the crude oil changes. Additionally, the degradation potential of a typical shallow, temperate environment with oil production and shipping was evaluated to emphasise the significant differences in degradation due to environmental conditions. Diagnostic ratios are a commonly used method in oil spill assessments to distinguish between degradation processes. The method is robust and insensitive to changes in extraction efficiency or analysis. Diagnostic biodegradation ratios, such as nC17 /pristane, nC18 /phytane, PAH isomers or other compounds with the same physical-chemical properties but different susceptibilities to microbial attack, are used as markers of biodegradation,
and to identify sources of hydrocarbon pollution [12–14]. A change in diagnostic ratios is evidence for biodegradation, which is especially important in subsurface spills where biodegradation is the only true degradation process. To examine the biodegradation potential of the two environments under laboratory conditions, a microcosm study was set up with the addition of typical North Sea crude oil (Oseberg Blend) to subsurface water from the Disko Bay, Greenland (Arctic) and to surface water from the North Sea, Denmark (temperate) in 1 L microcosms. The degradation of crude oil and changes in the microbial degrader communities was followed over a 10-week incubation period. 2. Materials and methods 2.1. Chemicals: The North Sea crude oil, Oseberg Blend (Statoil, Norway), was used for oil incubations For solid phase extraction and gas chromatography - mass spectrometry (GC–MS), methanol (HPLC grade, Rathburn), npentane (HPLC grade, VWR Chemicals), acetone (HPLC grade, Rathburn), dichloromethane (HPLC grade, Rathburn), ammonia solution (28–30%, MERCK) and ammonium acetate (≥98%, MERCK) were used. For instrument performance assessment, a GC–MS tune mixture was used. This was composed of benzidine (99.9%, Sigma), 1,1,1-Trichloro-2,2-bis(4-chlorophenyl) ethane (99.7%, Sigma), pentachlorophenol (99.9%, Sigma, Supelco) and decafluorotriphenylphosphine (99.3%, Sigma, Supelco) (50 g mL−1 of each). For further quality assurance and quality control, a reference sample composed of 19 PAHs with 2–6 aromatic rings and 15 deuterated PAHs with 2–6 aromatic rings was analysed between samples. Internal standard and recovery standard solutions consisted of mixtures of deuterated PAHs and nitrogen- and oxygen-containing polycyclic aromatic compounds in methanol. The internal standard stock solution contained a mixture of d8 -naphthalene (13.42 g mL−1 , 99%), d8 -dibenzothiophene (8.82 g mL−1 , 98%), d10 -acenaphthene (7.96 g mL−1 , 98%), d10 phenanthrene (13.05 g mL−1 , 98%), d10 -pyrene (8.27 g mL−1 , 98%) and d10 -fluorene (7.68 g mL−1 , 98%) from the Cambridge Isotope Lab and d9 -acridine (12.50 g mL−1 , 98.7%, CHIRON AS), d8 -carbazole (12.50 g mL−1 , 98.9%, CHIRON AS), d8 -antraquinonone (9.05 g mL−1 ) and d5 -phenol (13.78 g mL−1 , 98%, Sigma–Aldrich). The recovery standard stock solution contained d10 -anthracene (7.07 g mL−1 , 98%, Dr. Ehrenstorfer Gmb), and d8 -acenaphthylene (7.76 g mL−1 , 98%), d10 -fluoranthene (6.91 g mL−1 , 98%), d12 -benzo[a]anthracene (7.12 g mL−1 , 98%), d12 -benzo[a]pyrene (8.40 g mL−1 , 98%) and d12 -indenol(1,2,3–c and d) pyrene (6.84 g mL−1 , 98%) from the Cambridge Isotope Lab, and hydrochloric acid (37%, VWR Chemicals). Substrates used for Most-Probable-Number enumeration (MPN) were hexadecane (99%, Sigma–Aldrich), m-xylene (>99%, Fluka), 2-methylnaphthalene (97%, Sigma–Aldrich), 1-naphtol (≥99%, Sigma–Aldrich), 1-naphthoic acid (98%, Alfa Aesar) and silicone oil AR20 (Sigma–Aldrich). Dilutions were made in pre-mixed Bushnell–Haas minimal mediums containing magnesium sulphate (0.2 g L−1 ), calcium chloride (0.02 g L−1 ), monopotassium dihydrogen phosphate (1.0 g L−1 ), diammonium hydrogen phosphate (1.0 g L−1 ), potassium nitrate (1.0 g L−1 ) and ferric chloride (0.05 g L−1 ) (DifcoTM , Becton, Dickinson and Company, Sparks, MD21152, USA) [15]. 2.2. Sampling Two locations, the North Sea and the Disko Bay, represented typical temperate and Arctic marine environments. The North Sea
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is influenced by oil exploration and extensive shipping traffic, whereas, the Disko Bay in Western Greenland is a pristine area of great interest for future oil exploration and exploitation [16]. Sampling of the North Sea was carried out on the 6th of July 2013 from a small fishing boat. The sample was collected from a depth of 1.5 m 1.8 km north–west of Hirtshals Harbor (northern Denmark) at the position 57◦ 36 18“N, 09◦ 56 14“E. The sample was stored for four days at 5 ◦ C until use. Sampling of the Disko Bay was carried out on June 29th, 2013. Approximately 50 L of water was collected from a depth of 150 m using a 10 L Niskin water sampler (KC Denmark A/S, Silkeborg, Denmark) at a position 5 km south of Qeqertarsuaq at 69◦ 11 16“N, 53◦ 31 06“W. The Disko Bay sample was stored at 5 ◦ C for 11 days until initiation of the microcosm experiment. The temperature of both samples was continuously logged during transportation. The North Sea sample never exceeded 20 ◦ C, and the Arctic sample never exceeded 10 ◦ C. 2.3. Microcosm set-up Subsamples of 750 mL with 100 mg L−1 of Oseberg Blend crude oil were incubated for 10 weeks in 1 L glass flasks at 15 ◦ C (North Sea subsamples) or 5 ◦ C (Disko Bay subsamples). These temperatures represented typical water temperatures at the sampling times. Triplicate microcosms were sacrificed after 0, 4, 8, 15, 36 and 71 days of incubation. The microbial degrader community was analysed for all these incubation times, whereas the composition of the oil phase was analysed only for days 0, 15 and 71. The day 0 samples were sacrificed after nine hours of incubation to ensure equilibrium between the oil and water phase. A parallel series of autoclaved controls, with an addition of 100 mg L−1 Oseberg Blend, were used to estimate the physical removal processes. A series of controls without oil were used to determine the inherent changes in the microbial communities that were not caused by oil. For the first 24 h of incubation, the flasks were sealed with Teflon-lined lids to avoid evaporation and allow time for phase equilibration of the oil compounds between the oil phase and the water phase. The lids were then removed, and the microcosms were covered with oxygen- and carbon dioxide-permeable low-density polyethylene film (cling film). During incubation, the microcosms were shaken for five minutes at 120 rev min−1 approximately every 48 h to facilitate aeration. The microcosms were incubated in the dark to avoid photo-transformation of the oil compounds. Subsamples of the water phase were collected at each sampling day for MPN enumeration of hydrocarbon-degrading microbes. The microcosms were then acidified with 10 mL of 1 M hydrochloric acid to stop further biodegradation and left overnight for separation of the oil- and water phases before chemical analysis of the oil phase. 2.4. MPN enumeration of hydrocarbon-degrading microorganisms The size of microbial subpopulations that could utilise specific hydrocarbons as the sole source of carbon and energy were estimated with a modified version of a previously published microplate method [17] where, hydrocarbon growth substrates are added to the microplate wells in a separate silicone phase (silicone oil AR20, Sigma–Aldrich) to avoid toxic effects. The concentrations in the silicone phase were as follows: 10% hexadecane, 2% 2-methylnaphthalene, 1% m-xylene, 0.5% 1-naphtol and 0.5% 1naphthoic acid. The substrates were pasteurised for five minutes in a 70 ◦ C water bath in tightly closed glass scintillation vials with aluliners in the lids. A half strength Bushnell–Haas minimal medium [15] was supplemented with 30 g L−1 of NaCl (BH-medium) and autoclaved for 20 min at 121 ◦ C in tightly closed red-cap bottles. The precipitate was removed after autoclavation by decanting into sterile blue-cap bottles. For MPN enumerations, 1 mL subsamples were
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sampled from each microcosm. Subsamples from triplicate microcosms were pooled to give one mixed sample for each treatment at each time point. Ten-fold dilution series’ of the mixed samples were prepared in the BH-medium. From each dilution (10−1 –10−8 ), four times 200 L were added to six microtiter plates. Then, 15 L of a hydrocarbon substrate solution was added to each microplate well (one substrate per microplate), a microplate lid was put on and the microplates were placed in airtight plastic boxes. A small vial with extra substrate solution (1 mL) was placed on top of the plates to compensate for evaporation. Plates with sample dilutions and silicone oil, but no hydrocarbons, served as negative controls. The plates were incubated at 10 ◦ C in the dark. A vial with fresh substrate was added every three weeks. After 10 weeks of incubation, the absorbance of each microplate well was read at 450 nm on a ThermoMax microplate reader (Molecular Devices). Wells were scored growth-positive when they had an absorbance higher than 0.1, and MPN estimates were calculated according to Hurley and Roscoe [18] from the distribution of growth-positive and growthnegative microplate wells. The detection limit was calculated as one growth-positive well in the lowest dilution, giving a LogMPN of 1.1. 2.5. Chemical analysis Five hundred and fifty millilitres of the water phase was removed, and the residual oil phase was extracted two times with 5 mL dichloromethane. To all extracts, 200 L internal standard stock solution was added, samples were volume adjusted to 5 mL by evaporation in a fume hood at room temperature and 200 L recovery standard was added. Oil phase samples were diluted tenfold to give an adequate signal for GC–MS analysis of approximately 2000 mg oil L−1 solvent. The oil composition was analysed by gas chromatography (HP-7890A, Agilent Technology) with a 60 m ZB-5 capillary column (25 mm inner diameter and 0.25 m film thickness) interfaced to a quadrupole mass spectrometer (HP-5975C, Agilent Technology). Measurements were done by selected ion monitoring (SIM) of 55 ions divided into 12 groups using electron ionisation. One L sample was injected by pulsed splitless injection at 315 ◦ C and helium was used as a carrier gas with a flow rate of 1.1 mL s−1 . Oven temperature was ramped from initial 40 ◦ C (hold 2 min) to 100 ◦ C by 25 ◦ C s−1 followed by 5 ◦ C s−1 up to 315 ◦ C (hold 13.4 min). The method is described in detail in Gallotta and Christensen (2012) [19]. 2.5.1. Quality assurance and quality control Oil phase extracts were analysed in three batches of 23 vials, including samples and test solutions. Test solutions of dichloromethane, reference samples (with a known mixture of PAHs and deuterated PAHs), mixture samples (a mixture of thirteen samples) and a GC–MS tune mixture were analysed between samples and batches. These solutions were used for quality control of analysis, e.g. daily variations, cross contamination, peak shape, chromatographic resolution and sensitivity, and for verification of tuning, injection port and column performance [19]. 2.5.2. Data processing The data consisted of GC–MS SIM chromatograms of 44 oil phase extracts (three sampling days, triplicates, two locations, two exposures (autoclaved and oil exposed) and eight 0 mg L−1 microcosms as biological controls). The compound peak area was determined by manual integration according to the European Committee for Standardization, CEN oil spill identification guideline (CEN/TR 15,522-2:2009, www.cen.eu) in Agilent MSD Productivity ChemStation (E.02.02.1431). All calculations were made in Excel 2007 (Microsoft Office).
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Fig. 1. Most probable number enumeration (MPN, cells mL−1 ) of specific hydrocarbon degrader sub-populations (n = 1 ± one standard error).
3. Results 3.1. MPN enumeration of hydrocarbon-degrading microorganisms MPN enumerations were carried out for five growth substrates (Fig. 1) representing n-alkane degraders (hexadecane), monoaromatic degraders (m-xylene), diaromatic degraders (2methylnaphthalene) and polar aromatic degraders (1-naphthol
and 1-naphthoic acid). The MPNs gave several unexpected results. First, we did not detect any cultivable degraders able to grow with 1-naphthol or 1-naphthoic acid as the sole source of carbon and energy. The absence of degraders was especially unexpected for 1-naphthoic acid, which is an intermediate in the microbial metabolisation of 1-methylnaphthalenes [20]. Second, it was striking that the addition of crude oil did not result in massive increases of cultivable hydrocarbon-degrader MPNs. It was especially interesting that hexadecane degraders, which were found in
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Fig. 2. nC8 -nC32 alkanes in samples (triangles) after 15 (filled symbols) and 71 (open symbols) days of incubation, normalised to 17␣,21-hopane and compared to day 0 (n = 3 ± one relative standard deviation). Additionally, n-alkanes in the corresponding autoclaved controls are depicted (diamonds).
high initial numbers in the North Sea sample, increased less than 10-fold in microcosms spiked with crude oil and soon dropped to the same level as in the unspiked control microcosms. The most significant change, induced by the addition of crude oil, was the increase in 2-methylnaphthalene degrader MPNs in both the North Sea and the Disko Bay microcosms. Degraders of hexadecane, m-xylene and 2-methylnaphthalene were present in high numbers already at day zero in the North Sea samples, whereas, in the Disko Bay samples, m-xylene degraders and especially 2methylnaphthalene degraders were detectable after enrichment for 15 and 36 days, respectively. This might indicate that the microbial degrader community from the North Sea was more preadapted to oil degradation than the Arctic microbial community from Disko Bay. 3.2. n-alkanes and alkyltoluenes Changes in nC8 -nC32 alkanes compared to day 0 are shown in Fig. 2. The data was normalised to 17␣,21-hopane because this compound is highly recalcitrant in the environment and largely unaffected by biological degradation [12]. Surprisingly, only shortchain n-alkanes were significantly affected in samples from the North Sea (Fig. 2). After 71 days of incubation, nC8 –nC14 alkanes were reduced to 0.1–75.0% of the initial values, whereas, alkanes with chain lengths above nC14 remained almost unaffected with recoveries of 90.4–104.5% at day 71. According to the United States Environmental Protection Agency definition, nC8 -nC14 alkanes are classified as volatile organic compounds (VOCs) (BP 50-260 ◦ C), nC15 –nC24 alkanes are classified as semi-volatile organic compounds (SVOCs) (BP 240–400 ◦ C) and longer n-alkanes are classified as non-volatile (non-VOCs) (www.epa.gov). From this definition, it is clear that only volatile alkanes up to nC14 were removed by
evaporation in the North Sea samples, both in the intact samples and in the autoclaved controls. For the Disko Bay samples, we observed a much larger effect on the n-alkanes (Fig. 2). The amounts of VOCs were reduced to 0.0-4.4%, SVOCs to 5.5–42.9% and non-VOCs to 59.7–102.4% compared to day 0. In the autoclaved controls, only the VOCs were affected (0.0–88.3% after 71 days compared to day 0). The large removal of semi- and non-volatile n-alkanes, compared to the autoclaved controls, demonstrated that n-alkane biodegradation was a key factor in oil degradation only in the Disko Bay samples. Additionally, SVOCs were not expected to show large removal from evaporation in the Disko Bay samples, which were incubated at lower temperatures than the North Sea samples. The diagnostic ratios of nC17 /pristane and nC18 /phytane were calculated to confirm biodegradation. Fig. 3 shows the changes in diagnostic ratios, normalised to autoclaved controls, when expressed relative to the day 0 ratios. No changes in the diagnostic alkane biodegradation ratios were seen in the autoclaved controls for the North Sea samples or the Disko Bay samples (99.9–100.9% compared to day 0, data shown in Supplementary information). This demonstrated that physical removal processes, e.g. evaporation, as assumed, were negligible with respect to these biodegradation ratios. Therefore, the decrease in nC17 /pristane and nC18 /phytane in the intact samples was solely caused by alkane biodegradation. In samples from the North Sea, alkane biodegradation ratios decreased only slightly (to 93.6 ± 3.3% of their initial values for nC17 /pristane and to 96.3 ± 3.1% for nC18 /phytane). This confirms the lack of n-alkane biodegradation in the North Sea water seen in Fig. 2. For the Disko Bay samples, both nC17 /pristane and nC18 /phytane were reduced significantly (to 9.2 ± 8.0% and 9.2 ± 6.8%, respectively), verifying that biodegradation was the only significant process in the removal of n-alkanes in the Disko Bay water.
Fig. 3. Diagnostic ratios of nC17 /pristane and nC18 /phytane normalised to the corresponding autoclaved controls and compared to day 0 (n = 3 ± one relative standard deviation).
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Table 1 Boiling point, octanol-water partitioning coefficient (LogKow ), retention time and chemical formula for alkyltoluenes (group I, II and II) and their corresponding n-alkanes. BP, ◦ C
LogKow
RT, min
Chemical formula
nC16 Group I alkyltoluene
286.4 296.4
8.2 7.0
23.6 22.8–23.3
C16 H34 C16 H26
nC17 Group II alkyltoluene
302.0 310.9
8.7 7.5
25.8 25.2–25.6
C17 H36 C17 H28
nC18 Group III alkyltoluene
316.3 324.4
9.2 8.0
27.9 27.4–27.9
C18 H38 C18 H30
Alkyltoluene structure (ortho-isomer)
Calculated in EPIWEB 4.1. Ortho substituted alkyltoluenes are used for calculations.
The initial loss of low molecular weight alkyltoluenes is comparable to that of n-alkanes. Three groups of meta-, para- and ortho-alkyltoluenes were chosen with retention times corresponding to nC16 (group I), nC17 (group II) and nC18 alkanes (group III), and thereby, were estimated to be of equivalent boiling points to these n-alkanes (Table 1). In Fig. 4, the three groups of alkyltoluenes are presented as compared to day 0. All data were normalised to 17␣,21-hopane and autoclaved controls. In the North Sea samples (Fig. 4), there were no changes in the alkyltoluenes at day 15 or 71 (98.1–175.8% compared to day 0). This was in accordance with results for n-alkanes that were removed only by evaporation of the short-chain alkanes up to nC14 . In samples from the Disko Bay, we detected the removal of alkyltoluenes for both days 15 and 71. The group I alkyltoluenes showed 23.3 ± 27.2%, 48.6 ± 29.6% and 102.4 ± 8.9% of initial values after 71 days of incubation for para-, meta- and ortho-alkyltoluenes, respectively. For group II and III alkyltoluenes, the isomers varied between 46.0 and 103.5%. nC16 to nC18 alkanes in the Disko Bay microcosms were reduced to 7.1–9.7% of their initial values at day 71. The degree of n-alkane removal in the Disko Bay water was, hence, much higher than that observed for alkyltoluenes of corresponding alkane chain lengths. Furthermore, the alkyltoluenes were removed in the isomer specific order of para- > meta- » ortho-alkyltoluene, which is in accordance with previous studies showing lower biodegradation of the ortho-isomer of xylene [21].
3.3. Polycyclic aromatic hydrocarbons (PAHs) and dibenzothiophenes Changes in naphthalene, phenanthrene, fluorene, pyrene, chrysene and the sulphur-containing dibenzothiophene, together with their alkyl-substituted homologues (C0 –C4 ) were measured by oil hydrocarbon fingerprinting on GC–MS. The percentage of the five PAHs, dibenzothiophene and their alkyl-substituted homologues is shown in Fig. 5. All data was normalised to 17␣,21-hopane and expressed relative to the equivalent autoclaved controls, thereby excluding effects from physical weathering processes. In the North Sea microcosms, we found biodegradation at day 71 of C0 - – C4 -naphthalenes (0.0–66.5% compared to autoclaved controls), C0 - – C2 -dibenzothiophenes (3.4–80.1%), C0 - – C2 -phenanthrenes (4.2–77.6%), C0 - – C3 -fluorenes (6.5–85.3%) and C0 -pyrenes (85.8 ± 7.0%). Neither C1 - – C3 - chrysenes, C1 - – C2 pyrenes, C3 - – C4 -phenanthrenes nor C3 - – C4 -dibenzothiophenes were biodegraded. The non-substituted PAHs and dibenzothiophene were degraded to the largest extent for all compound groups, showing that the degradation was negatively correlated with alkyl substitution. Additionally, the biodegradation of PAHs was decreased as the number of aromatic rings increased. In the Disko Bay samples, only C0 - and C1 -naphthalenes showed evidence of biodegradation by decreased amounts at day 71 (54.0 ± 13.7% and 64.5 ± 8.0%, respectively). From the MPN esti-
Fig. 4. Percent of alkyltoluenes (alkylT) normalised to 17␣,21-hopane and the corresponding autoclaved controls compared to day 0 (n = 3 ± one relative standard deviation). Group I alkyltoluenes have retention times corresponding to nC16 -, group II to nC17 - and group III to nC18 alkanes.
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Fig. 5. PAHs, dibenzothiophenes and the alkyl-substituted homologues normalised to 17␣,21-hopane and compared to the corresponding autoclaved controls (n = 3 ± one relative standard deviation). The North Sea samples are depicted in the left part of the graphs, while the Disko Bay samples are depicted to the right.
mates, it was clear that microorganisms capable of degrading more metabolically complex hydrocarbons, in this case, expressed by growth on m-xylene and 2-methylnaphthalene, were at low levels in the Disko Bay microcosms compared to the North Sea microcosms, even after 71 days of incubation (Fig. 1). The low content of cultivable degrader microorganisms might explain the lack of degradation of the metabolically complex hydrocarbons with more than two aromatic rings and with alkyl substitution.
We also calculated diagnostic biodegradation ratios for the PAHs and the dibenzothiophenes (Fig. 6). In the North Sea samples, all ratios indicated biodegradation of PAHs. The diagnostic ratios for the dibenzothiophenes remained almost constant after 71 days of incubation, with values of 108.4 ± 3.8% for 2 + 3-methyldibenzothiophene/4-methyldibenzothiophene and 91.1 ± 9.8% for 1-methyldibenzothiophene/4methyldibenzothiophene compared to day 0 (100%). The
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Fig. 6. Diagnostic ratios of PAHs and dibenzothiophenes normalised to corresponding autoclaved controls and compared to day 0 for North Sea and Disko Bay samples (n = 3 ± one relative standard deviation). mN: methylnaphthalenes, mF: methylfluorenes, mP: methylphenanthrenes and mDBT: methyldibenzothiophenes.
diagnostic ratios of alkyl-substituted fluorenes showed the largest responses with decreases to 63.0 ± 5.9% for 2 + 3methylfluorene/4-methylfluorene and to 57.0 ± 10.7% for 1-methylfluorene/4-methylfluorene at day 71. For day 71, it was not possible to calculate a reliable estimate of the 2methylnaphthalene/1-methylnaphthalene diagnostic ratio since both isomers were almost completely removed. In samples from the Disko Bay, only the diagnostic ratio of 2methylnaphthalene/1-methylnaphthalene decreased (56.6 ± 2.2% at day 71), indicating degradation of the two-ring PAHs. The diagnostic ratios of the three-ring PAHs (fluorenes and phenanthrenes) and the dibenzothiophenes remained constant during the incubation (99.4–104.7% at day 71 compared to day 0). The lack of changes in the diagnostic ratios for three-ring PAHs and dibenzothiophenes proved the limited potential for biodegradation of metabolically complex aromatics in the Disko Bay sample. Autoclaved controls showed no changes in the diagnostic ratios, except for 2-methylnaphthalene/1-methylnaphthalene where the autoclaved controls were reduced to 72.2 ± 2.1% for the North Sea samples and 86.0 ± 0.7% at day 71 for the Disko Bay samples (data shown in Supplementary information). This change, however, seems to be by physical–chemical processes rather than biological degradation, since we did not detect any microbial activity in the autoclaved controls as determined by [3 H]-Leucine incorporation at days 0, 15 and 71 (unpublished results).
environments, the degree of biodegradation decreased with alkyl substitution, which has also been found in several previous studies of crude oil biodegradation [9,10,22]. Water samples from the North Sea showed a clear potential for biodegradation of the metabolically complex compounds, whereas the subsurface samples from the Arctic Disko Bay environment only showed the degradation of simple hydrocarbons like the n-alkanes. Extensive biodegradation of PAHs and dibenzothiophenes compared to n-alkanes, as seen in the North Sea samples, is the opposite of the expected degradation order, but other studies have indicated a similar pattern [22,23]. A recent study of oil degradation in Arctic sea-water indicated the substantial loss of oil at −1 ◦ C with significant biodegradation of 2- – 4-ring PAHs and their alkyl-substituted homologues [24]. These results are different from ours, but it was not indicated whether sterile controls were included in their study. Abiotic removal processes might therefore have caused the different degradation patterns. It may be problematic to directly extrapolate from our limited microcosm study to a real Arctic subsurface spill, but our results indicate that aromatic hydrocarbons with more than two rings may show high persistence in Arctic subsurface waters. These persistent compounds may detrimentally affect marine organisms and Arctic marine ecosystems, and ought to influence future oil spill responses in the Arctic. Our results indicate that risk assessment data from the shallow North Sea is not relevant to subsurface spills in the Disko Bay area, and probably also of little relevance to other pristine places in the marine Arctic.
4. Discussion In the present study, we compared the potential for microbial oil degradation in water from the Disko Bay and the North Sea. The sampling depth of 150 meters in Disko Bay was chosen to study the oil spill response of a subsurface Arctic environment. Surface samples from the North Sea, on the other hand, were chosen for comparison because this environment is well characterized with respect to oil degradation. Lessons from this environment might therefore be applied for oil spill risk assessment of Arctic environments, where, data are scarce. Our results, however, demonstrated a completely different pattern of crude oil degradation in the two environments. The degradation order in the North Sea samples was PAHs and dibenzothiophenes (naphthalenes > dibenzothiophenes > phenanthrenes > fluorenes) > alkyltoluene > n-alkanes, whereas, in the Disko Bay samples, the order was reversed: n-alkanes > alkyltoluenes (para> meta- > ortho-isomer) > PAHs and dibenzothiophenes. In both
Acknowledgements The authors would like to thank the University of Copenhagen for funding this project and Katrine Scheibye for collaboration on the topic of oil degradation in Arctic environments. Peter Jensen is acknowledged for help during field sampling at Hirtshals, Denmark, as is the staff at Arctic Station, University of Copenhagen for their assistance during field sampling at Disko Bay, Greenland. This study was funded by the European Commission (KILLSPILL project, grant agreement 312139). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jhazmat.2015.06. 046
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