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Ecotoxicology and Human Environmental Health
Does microbial biodegradation of water-soluble components of oil reduce the toxicity to early life stages of fish? Bjørn Henrik Hansen, Julia Farkas, Trond Nordtug, Dag Altin, and Odd Gunnar Brakstad Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b06408 • Publication Date (Web): 07 Mar 2018 Downloaded from http://pubs.acs.org on March 8, 2018
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Does microbial biodegradation of water-soluble
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components of oil reduce the toxicity to early
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life stages of fish?
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Bjørn Henrik Hansen†*, Julia Farkas†, Trond Nordtug†, Dag Altin‡ & Odd Gunnar
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Brakstad†
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†
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‡
SINTEF Ocean AS, Postboks 4762 Torgarden, 7465 Trondheim, Norway BioTrix, 7020 Trondheim, Norway
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*Corresponding author: Bjørn Henrik Hansen. E-mail:
[email protected]. Phone: +47
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98283892
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Key words: Oil; PAH; biodegradation; ecotoxicity; cardiac toxicity; developmental toxicity
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Abstract
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Microbial degradation following oil spills results in metabolites from the original oil.
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Metabolites are expected to display lower bioaccumulation potential and acute toxicity to
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marine organisms due to microbial-facilitated incorporation of chemical functional groups
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and a general decrease in lipophilicity. The toxicity and characterization of metabolites are
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poorly studied. The purpose of the present work was to evaluate the toxicity of degraded (0-
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21 days) water-soluble oil components. Low-energy water accommodated fraction (LE-WAF)
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of a weathered crude oil was prepared with nutrient amended seawater at 5°C, kept in dark,
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and sampled at 0, 10, 14 and 21 days. Samples were extracted with dichloromethane and
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toxicity experiments were conducted with reconstituted extracts. Toxicity experiments were
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conducted for 4 days on developing cod (Gadus morhua) embryos during a critical period of
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their heart development. After exposure, embryos were kept in clean seawater and observed
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until 5 days post hatch. Survival, hatching, morphometric aberrations and cardiac function
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was studied. The expected decrease in sub-lethal toxicity during the biodegradation period
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was not found; indicating that metabolites formed during biodegradation likely contributed to
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larvae toxicity.
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Table of Contents (TOC)/Abstract Art.
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Introduction
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In the event of an oil spill at sea, oil will be affected by abiotic and biotic processes that
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change the physical-chemical properties of the oil. These 'weathering' processes include
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dispersion, dissolution, spreading, emulsification, UV-oxidation and microbial degradation.1
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In cold marine environments, like the Arctic, biodegradation processes are expected to be
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slower than in temperate environments.2,
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seawater (SW) after oil spills is associated with oxidative processes. Aerobic n-alkane
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degradation is associated with oxygenases and dehydrogenases, in which the alkane is
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converted to alcohols and further to acetyl-coA4, while aromatic HCs are degraded primarily
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by dioxygenases.5 The resulting metabolites are more water-soluble and thus are attributed
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with lower octanol-water partitioning coefficient (Kow). Often, acute effect concentrations
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(e.g LC50) of HCs are predicted using LogKow vs LogLC50-regressions6, resulting in
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reduced acute toxicity estimates of metabolites compared to their original parent components.
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The toxicity of oil to marine species depends on the characteristics of the oil, and for pelagic
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organisms, dissolved oil components are expected to display the highest bioavailability and
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subsequently have the highest contribution to toxicity7-10. Biodegradation likely reduces the
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toxicity of oil spills over time due to the rate of biodegradation of dissolved oil components.
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Oil spill models are often utilized to perform risk assessments and assessments of
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environmental harm. The overall fate of the oil is predicted using empirical data of different
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weathering processes, including biodegradation rates of known oil components or component
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groups.11 Metabolites resulting from microbial degradation are rarely accounted for when
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predicting damage to natural resources. Unfortunately, metabolized oil components are often
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omitted from modelled assessments as knowledge pertaining to their fate and effects is
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limited. Metabolites are generally not expected to contribute to overall toxicity since they
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Biodegradation of oil hydrocarbons (HCs) in
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display a lower potential for bioaccumulation compared to their more lipophilic parent
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compounds. This, however, lacks verification.
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Relations between biodegradation and acute toxicity have been investigated in several studies,
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but mainly in soil or groundwater 12-15 or using bacterial cultures16, 17. Only a few studies have
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systematically investigated the relationships between biodegradation and ecotoxicity to
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marine organisms18, 19 and in general, toxicity data are limited for Arctic species.20
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In recent years, the sensitivity of early life stages (ELS) of fish to oil exposure has received
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increased attention, particularly after the Deepwater Horizon incident in the Gulf of Mexico in
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2010.21-23 It has been shown that exposure to low concentrations of weathered crude oils may
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cause craniofacial and jaw deformations and cardiotoxicity manifested as pericardial oedema,
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reduced heart rate (bradycardia), arrhythmia, contractility defects, reduced stroke volume and
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reduced cardiac output.21,
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larvae following acute spills.26 It has been argued that these effects are PAH-dependent27,
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however, it should not be ruled out that these effects might be mediated by the presence of
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unknown components like metabolites.
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The purpose of the current work was to study how biodegradation impacted the toxicity of
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WAF to ELS fish. We hypothesised that toxicity decreases as a function of WAF
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biodegradation. We hope that these results will generate relevant information and empirical
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data for improving risk and damage assessment processes following acute oil spills.
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Such effects have even been observed in field-collected fish
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Materials and Methods
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Water supply and WAF biodegradation. Natural seawater (SW) was used as microbial
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inoculum in the experiment. The SW was collected from a depth of 80 m (below thermocline)
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in a non-polluted Norwegian fjord (Trondheimsfjord; 63°26'N, 10°23'E), supplied by a 5 ACS Paragon Plus Environment
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pipeline system from the source to our laboratories (salinity of 34 ‰). The SW was
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acclimated to 5°C three days before start of the experiments, aerated with filter-sterilized air
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(0.22 µm), and fortified with mineral nutrients.28 The oil used in this experiment was Troll oil;
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a naphthenic oil that has been the subject of numerous laboratory experiments.7-9, 29, 30 Two 10
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L low energy water accommodated fractions (LE-WAFs) of crude oil were generated in glass
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bottles as described previously31 with an oil-to-water ratio of 1:100 using nutrient-enriched
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seawater.32. Slow stirring was used to prevent the formation of oil droplets and stirring was
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performed for 72 hours at 5°C. WAF preparations were collected from the glass port located
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at the bottom of the bottle. The two WAFs (100% stock solutions) were pooled and
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homogenized prior to sampling. Samples (800 ml) were acidified, liquid-liquid-extracted
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using dichloromethane (DCM), and analysed for C10-C36-total extractable material (C10-C36-
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TEM) using Gas Chromatography – Flame Ionization Detector (GC-FID). Semi-volatile
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organic components (SVOC), including PAHs and phenols, were also analysed using gas
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chromatography – mass spectrometry (GC-MS). The concentration of unknown components
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in the GC-FID chromatograms, here regarded as 'unresolved complex mixtures' (UCM) in the
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WAFs were estimated by subtracting the identified and quantified components (SVOC) from
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the TEM. Samples (40 ml) was also taken for analyses of volatile organic components (VOC)
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using Purge & Trap GC-MS using standard methodology.31 A complete list of target
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compounds are given in Supplementary Information (Table S1 and Table S2). Water samples
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were also analysed for oxygen concentrations using an oxygen meter (Model 59 with 5905
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BOD probe, YSI Inc., OH, USA). An aliquot of the 100% WAF (800 ml) was separately
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extracted with dichloromethane (DCM) for subsequent reconstitution for the toxicity tests
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(day 0, T0). Finally, the remaining 100% WAF was distributed into separate 2 L glass bottles
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filled to the rim and capped. These bottles were kept at 5°C until they were sampled at 10
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(T10), 14 (T14) and 21 (T21) days. Each time point was analysed for oxygen content, TEM,
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SVOC and VOC as described above. In addition, an aliquot of these biodegraded WAF
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samples was obtained at each time point and extracted using DCM for subsequent
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reconstitution and toxicity testing (800 ml),.
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Preparation of exposure solutions. When fish embryos were available, the biodegraded
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WAF extracts (t=0, t=10, t=14 and t=21 days) were reconstituted into filtered seawater to
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prepare the representative exposure solutions at each time point. Each DCM-extract, extracted
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from biodegraded WAFs (800 ml), was added to the bottom of a glass bottle (1 L) and flushed
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using nitrogen gas for 60 min to remove the DCM. Using filtered seawater (0.22 µm), the
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dried extract was reconstituted to the same volume of seawater as it was initially extracted
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(800 ml). Extracts were resolubilized into seawater using ultrasonication for 30 min. The
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reconstituted WAFs from each time point (t=0, t=10, t=14 and t=21 days) were subsequently
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diluted in filtered seawater to three nominal concentrations (10%, 50% and 100%
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(undiluted)). 'Solvent controls' (DCM controls) were prepared following the same
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reconstitution procedure as oiled-extracts, but contained clean DCM instead of extracts.
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Negative controls contained filtered seawater only. All prepared exposure solutions were
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cooled to 5°C and aeriated with filtered ambient air for 30 min to increase oxygen saturation.
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Aliquots of all reconstituted and diluted samples were extracted and analysed for TEM and
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SVOC as described above for exposure verification. Exposure solutions were then transferred
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into glass jars (80 ml) for the fish embryo exposure experiment.
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Fish exposure. A complete time line of the exposure experiment is given in Supplemental
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Information (Table S3). Fish embryos (day 0 post fertilization, 0 dpf) were obtained from the
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Austevoll Research Station, Norway. Embryos were shipped directly to our laboratory in a
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cooling container using airfreight. Embryos arrived less than 12 hours after fertilization and
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were immediately transferred to flow-through tanks (250 L) containing filtered seawater (1
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mm, 6°C). The flow-through rate ensured that the entire tank (250 L) of seawater was
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exchanged every 24 hours. Gentle aeration kept embryos moving continuously in the tanks.
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Dead embryos were removed from the tank daily.
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When embryos reached 9 dpf, they were transferred into the glass jars (80 ml) containing
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exposure solutions (472±63 embryos per jar). Embryos were exposed for 4 days until 13 dpf,
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and surviving embryos were transferred into clean seawater and monitored for an additional 9
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days (until 22 dpf). During the exposure (days 9-13 post fertilization), dead eggs were
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counted in all jars daily. During the post-exposure monitoring period (days 14-22 post
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fertilization), dead eggs were counted almost every day (days 16-18, 20 and 22 post
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fertilization). At the same time points, hatching and larvae mortality was monitored. The
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experiment was terminated at 5 days post hatch (dph).
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Seawater control groups included 8 replicates (N=8), and all treatments contained 4 replicates
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(N=4), with on average 472±63 embryos per replicate. Replicates were kept separate until
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being pooled at hatch. In the morning at largest hatch, newly hatched larvae from replicates
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were transferred into a new glass jar (80 ml) in order to provide a sufficient number of larvae
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at the exact same age to be sampled at the different time points. Oxygen saturation in the
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exposure containers were monitored throughout the experiment with a phase fluorometer
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(NeoFox with FOXY R-sensor, OceanOptics Inc., FL, US).
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Biological endpoints. The heart rates (HR) of individual embryos/larvae were monitored with
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automated video analyses. Videos of embryos (17 dpf) and individual larvae (1, 3 and 5 dph)
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were taken through a microscope (Eclipse 80i, Nikon Inc., Japan) equipped with a CMOS
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camera (MC170HD, Leica Microsystems, Germany). Microscopy images were also taken of
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larvae at 1, 3 and 5 dph for biometric analyses using Image J33, 34 and blinded deformation
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ranking analysis. Care was taken to make sure that the larvae from all groups were identical in
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age, so newly hatched larvae were transferred into new glass jars in the morning (being 99%) after
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incubation at 5°C for 21 days. The TEM fraction was also depleted, but to a lower extent
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2- to 6-ring PAH and phenols) were depleted by 98% (Supplemental Information Table S4).
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The TEM depletion was well in line with the oxygen consumption (24%), and both the
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oxygen consumption and TEM depletion could therefore be used as measures of
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mineralization. Since the experiment was performed in closed flasks without headspace,
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avoiding evaporation, biodegradation is likely the cause of the depletion. This was further
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substantiated by the temporary increase in the phenol concentrations, expected to be products
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of BTEX and PAH biodegradation.18,
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groups were mainly in agreement with recent results, comparing biodegradation of LE-WAF
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and gas compounds in natural SW at 5°C 37.
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Concentrations of the unresolved components from the GC-FID chromatograms (UCM
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fraction) was also reduced by almost 25% at T21, but the fraction of UCM increased, which
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includes undegraded unresolved components as well as metabolites (Fig. 2B). This is caused
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by an almost complete removal of the identified SVOC components (Fig 2C and D),
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particularly decalins (100% loss), naphthalenes (100% loss) and 2-3-ring PAHs (86.7% loss).
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Concentrations for 4-6 ring PAHs were very close to levels of quantification in all initial
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WAFs. As mentioned above, phenols increased at days 10 (38.5% increase) and day 14
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(164.2%) compared to the initial WAF, and returned to below T0 levels at day 21. This
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suggests that the biotransformation of cyclic organic oil components includes OH-substitution
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to generate phenols, and that these are readily broken down by the end of the three-week
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degradation period. The identified and quantified SVOC components represent a small
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fraction (
