AEM Accepted Manuscript Posted Online 19 June 2015 Appl. Environ. Microbiol. doi:10.1128/AEM.01470-15 Copyright © 2015, American Society for Microbiology. All Rights Reserved.
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Microbial community composition, functions and activities in the Gulf of Mexico, one
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year after the Deepwater Horizon accident.
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Etienne Yergeau1, Christine Maynard1, Sylvie Sanschagrin1, Julie Champagne1, David Juck1,
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Kenneth Lee2,3 and Charles W. Greer1*
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1
8
Canada
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2
National Research Council Canada, Energy Mining and Environment, Montréal, Quebec,
Centre for Offshore Oil, Gas and Energy Research (COOGER), Bedford Institute of
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Oceanography, Fisheries and Oceans Canada, Dartmouth, Nova Scotia, Canada
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3
12
Research Centre, Kensington, WA, Australia
Commonwealth Scientific and Industrial Research Organization (CSIRO), Australian Resources
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Keywords: Deepwater Horizon, microbial functions, microbial activities, Gulf of Mexico,
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metagenomics, metatranscriptomics
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Running title: Microbiology of the GOM one year after the DWH spill
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*
Corresponding author:
[email protected]; Tel: 514-496-6182
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Abstract
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Several studies have assessed the effects of the released oil on microbes, either during or
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immediately after the Deepwater Horizon accident. However, little is known about the potential
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longer-term persistent effects on microbial communities and their functions. In this study, one
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water column station near the wellhead (3.78 km SW of the wellhead), one water column
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reference station outside of the affected area (37.77 km SE of the wellhead), and deep-sea
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sediments near the wellhead (3.66 km SE of the wellhead) were sampled one year after the
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capping of the well. In order to analyze microbial community composition, function and activity,
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we used metagenomics, metatranscriptomics and mineralization assays. Mineralization of
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hexadecane was significantly higher at the wellhead station at a depth of ~1200 m as compared
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to the reference station. Community composition based on taxonomical or functional data
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showed that the samples taken at a depth of ~1200 m were significantly more dissimilar
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between the stations than at other depths (surface, 100 m, 750 m and >1500 m). Both Bacteria
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and Archaea showed reduced activity at depths of ~1200 m when comparing the wellhead
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station to the reference station, and their activity was significantly higher in surficial sediments
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as compared to 10 cm sediments. Surficial sediments also harbored significantly different active
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genera when compared to 5 and 10 cm sediments. For the remaining microbial parameters
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assessed, no significant differences could be observed between the wellhead and reference
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stations and between surface and 5-10 cm deep sediments.
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Introduction
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Following the explosion and sinking of the Deepwater Horizon (DWH) oil rig in the Gulf
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of Mexico, an estimated 3.26 to 4.9 million barrels of light crude oil was released at a depth of
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1544 m from April 20 to July 15, 2010, making it the largest and deepest offshore spill in United
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States history (3, 22). When including gaseous hydrocarbons, like methane, the total discharge
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was 40% higher than the abovementioned estimates (12). During the spill, a deep water oil
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plume was detected at depths of 1000-1200 m (4, 10), but this plume was no longer detectable
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after a few months (25), in agreement with the very high degradation rates observed in
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laboratory incubations (10). However, most microbiological research to date has focused on the
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effects of the oil spill with samples taken during the contamination event or shortly thereafter (2,
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10, 16, 17, 20, 24, 27, 33, 34, 39), and only one study reported on the bacterial communities at
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plume depth 1 year after the spill (41). In view of the high degradation rates observed and slow
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mixing of deep water, it was suggested that oxygen depletion at plume depth might persist for
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several years (1, 12, 26, 35). The cause, extent and duration of this oxygen depletion was
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subject to debate (11, 15, 16), and it is not clear how, and if, it would impact the microbial
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communities in the long term. Recent work also indicated that significant quantities of oil sank to
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the sea floor (38), potentially affecting microbial communities in the sediments.
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The microbial characterization of the water column shortly after the beginning of the spill
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identified Oceanospirillales as a dominant group of hydrocarbon degrading organisms making
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up as much as 90% of the of the 16S rRNA gene clone libraries (6, 10, 27, 33, 41). Shortly after
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this, other Gammaproteobacteria affiliated with Colwellia and Cycloclasticus appeared,
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indicative of a succession from alkane to aromatic degrading bacteria (6, 33, 39, 41). In
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addition, other phyla of bacteria (Alteromonas, Halomonas, Pseudoalteromonas) were observed
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in the water column (10, 39). A recent DNA-Stable Isotope Probing (SIP) study provided direct
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evidence that most of the abovementioned taxa were in fact capable of degrading various 3
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hydrocarbons (9). Following the spill, after the flow of hydrocarbons had been arrested,
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methylotrophs including known methane oxidizers, became dominant in the region of the plume
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(16). Microbial communities are at the base of several crucial biogeochemical processes in
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marine environments, including hydrocarbon degradation. Full ecosystem recovery is intimately
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linked to microbial community recovery. Microorganisms might also serve as highly sensitive
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bioindicators (37), as they have been shown to be sensitive to very low concentration of
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pollutants especially with regard to their transcriptome (43-45). For these reasons,
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microorganisms could be used as indicators of pollution and ecosystem recovery through the
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examination of their gene content and gene expression patterns.
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Two approaches were used to determine the potential effects of the DWH blowout on
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microbial communities more than one year after the event: 1) comparison of two water column
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stations, one very close to the well and the second 38 km away, outside the plume area; 2)
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depth profile of deep-sea sediment cores taken in the proximity of the Macondo well. We used a
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shotgun metagenomic and metatranscriptomic approach and compared the microbial functions,
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community composition and activities of the different stations with depth.
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Material and methods
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Sampling sites
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A map of the sampling sites is provided as Fig. 1. The wellhead water column station (BM-57,
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28.7051°, -88.4016°) was located at a distance of 3.78 km SW from the actual Deepwater
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Horizon wellhead and corresponded to the plume station BM-57 used by Hazen and colleagues
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(10). The reference water column station (A6, 28.6632°, -88.0095°) was located 37.77 km SE
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from the Deepwater Horizon wellhead, but in the same “dome” area and was outside the plume
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area during the spill. A series of 6 deep-sea sediment cores were collected on Nov. 16, 2011
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during a second cruise. The cores were collected from the vicinity of the Deepwater Horizon
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wellhead (around 28.715011°; -88.358703°, 3.66 km SE from the wellhead) at a depth of
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approximately 1600 m.
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Water and sediment sampling
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Water samples were collected between Sept. 9-16, 2011 using either a large bailing bucket
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(surface samples) or a CTD Niskin rosette equipped with 20 L bottles. For each depth, three
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replicate water samples were taken. Samples were returned to the boat and immediately
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transferred to 4 L carboys which were previously rinsed with 70% ethanol and sterile distilled
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water. Sample filtration was started immediately after transfer using Millipore GSWP (0.22 µm
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pore size, 47 mm diameter) filters and glass filter supports. Each 4 L water sample was filtered
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on two filters, resulting in a total of six filters per depth. The filters were then transferred to ice
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and then stored at -80˚C. Between samples, the glass filter supports were rinsed with 70%
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ethanol and sterile distilled water. For the shipping of filtered samples, coolers with dry ice were
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used and upon arrival at the lab, filters were stored at -80˚C until nucleic acid extraction was
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performed. Water samples destined for mineralization analysis were collected from the same 5
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carboys as used for filtration. Samples were placed in sterile 50 ml Falcon tubes and stored at
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4˚C. Samples were shipped on ice and upon arrival at the lab were placed immediately at 4˚C.
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Microcosms were started as soon as possible after arrival into the lab (within 24 h). Water for
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chemical analysis was also taken and kept at 4°C until processing.
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Sediments were frozen on-board the sampling vessel. Samples were shipped and
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received frozen and stored at -20˚C until processing. Sample processing was performed at -
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20˚C based on a protocol modified from Juck et al. (13). In brief, an approximately 5 cm strip of
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the core sample plastic sleeve was cut and removed (from top to bottom of the core) and a
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‘clean’ area of the core (i.e. not contacted by the sample sleeve) was exposed using a sterile
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chisel. Once this flat clean area was exposed, a sterile 1.4 cm drill bit was used to slowly drill
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into the core sample, parallel to the core surface. The drilled core sub-sample was then
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transferred to a sterile 50 ml Falcon tube and stored at -80˚C until extraction of nucleic acids
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was performed. Each core was sampled at 3 different depths – ‘0 cm’ was from the surface of
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the sediment to 1.4 cm, ‘5 cm’ was from approximately 4.3 to 5.7 cm from core surface and ’10
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cm’ was from approximately 9.3 to 10.7 cm from the sediment surface. From the remaining core
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samples, the material remaining at 0, 5 and 10 cm was sampled and used for hydrocarbon
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analysis as described below.
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Water microcosm mineralization assays
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Mineralization assays using microcosms were set up using 15 ml of seawater and
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(100,000 dpm) hexadecane (2.5 ppm), as sole carbon source, with no amendments added. The
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sealed bottles containing seawater from all the different depths for both water column stations
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and the spiked substrate were all incubated at 15˚C (the range of sample temperatures at the
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time of collection was 4˚C (bottom samples) to 30˚C (surface samples) with orbital shaking (140 6
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C labeled
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rpm) and a microcosm KOH trap (1 ml of 1.0 M KOH in a test tube). Sampling was performed at
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T=3, 7, 14, 22, 28, 35, 42, 49, 56 and 63 days. The amount of radioactive
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to the complete mineralization of the added carbon source (hexadecane) was determined by
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scintillation counting of the KOH solution recovered from the microcosm flasks and is presented
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as a percentage of
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During sampling of the KOH traps, atmospheric oxygen was introduced (through a 0.22 µm
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filter) into the microcosms to ensure sufficient aeration of the samples. The 700 m depth sample
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from the reference station was also used as a sterile abiotic control by autoclaving for 20 min
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and cooling to room temperature before addition of the radioactive spike.
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CO2 produced, due
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CO2 produced from the known quantity of carbon source added at T=0.
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Hydrocarbon analyses
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Water column samples were extracted for C10-C50 and PAH analyses using liquid-liquid
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extraction
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http://www.epa.gov/osw/hazard/testmethods/sw846/pdfs/3510c.pdf). Extracts of water were
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analyzed using high resolution gas chromatography (Agilent 6890 GC) coupled to a mass
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selective detector (Agilent 5973N) (Willmington, DE, USA) operated in the selective ion
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monitoring mode (SIM) using the following GC (MDN-5S column 30 m x 0.25mm id 0.25 μm film
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thickness, Supelco Canada) conditions: cool on-column injection with oven track mode (track 3
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°C higher that the oven temperature program) 80 °C hold 2 min ramp at 4 °C/min to 280 °C hold
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10 min. Deep-sea sediments were processed according to King and Lee (19) and the GC-MS
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conditions outlined for the water extracts were applied to sediment extracts.
(US
EPA
Method
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Total DNA extraction from filters (seawater)
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3510
C,
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From each water sample (2 stations X 5 depths X 3 replicates = 30 water samples), one filter
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was used and treated for DNA extraction, resulting in 30 DNA extracts. In the 50-ml Falcon
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tube containing the filter, 1.7 ml of Tris-EDTA (TE) pH 8.0 buffer was added with 45 µl of 20%
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(w/v) sodium dodecyl sulfate (SDS) and 9 µl of 20mg/ml proteinase K. The tube was incubated
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with gentle inversion at 37°C for one hour. At the end of the incubation, 300 µl of 5M NaCl was
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added in addition to 240 µl of a 10% (w/v) cetyl trimethylammonium bromide (CTAB) and 0.7M
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NaCl solution. The tube was incubated at 65°C for 10-min. The total DNA was extracted with 1
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volume of 24:1 chloroform/isoamyl alcohol. After centrifugation for 10 min at 3,000 x g, the
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upper phase was transferred and mixed with one volume of 25:24:1 phenol/chloroform/isoamyl
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alcohol. Following centrifugation at 16,000 x g for 10min at 4°C, the supernatant was
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precipitated by mixing 0.6 volume of isopropanol and 1/50 volume of glycogen (5 mg/mL),
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incubating one hour at -80°C and centrifuging at 12,000 x g for 30 min at 4°C. The DNA pellets
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were washed using 1ml of 80% (v/v) ice-cold ethanol and dried using a SpeedVac. The DNA
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was re-suspended in 50µl of nuclease-free water and treated with RNase If (NEB, Ipswich, MA)
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according to the manufacturer’s instructions. After the reaction was complete and the enzyme
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was inactivated, the DNA was purified with the QIAEX II Kit (QIAgen, Valencia, CA) and
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quantified using the PicoGreen assay (Invitrogen).
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Total RNA extraction from filters (seawater)
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For RNA extraction, one replicate seawater sample was used (5 depths X 2 stations = 10
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samples). All solutions were RNase free. In the 50-ml Falcon tube containing the filter, 1.6 mL of
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freshly prepared lysozyme (10 mg/ml in TE pH 8.0) and 80 uL of 20% SDS were added and the
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tube was then incubated at 64°C for 5 min. At the end of the incubation, 176 µl of 3M sodium
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acetate pH 5.2 and 1.6 mL of pre-warmed acid phenol was added to the lysate incubated at
8
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64°C for 6 min, with mixing every minute. The tube was transferred on ice for 2 min and then
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centrifuged at 16,000 x g for 10 min at 4°C. The upper phase was transferred and mixed with
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1.6 ml of chloroform before centrifugating at 16,000 x g for 2 min at 4°C. The upper aqueous
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phase was transferred, mixed with 20 µl of glycogen (5 mg/ml), 160 µL of 3M sodium acetate
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pH 5.2 and 4 mL of 100% ice-cold ethanol and incubated for 30 min on dry ice before
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centrifuging at 12,000 x g for 30 min at 4°C. The pellet was washed with 1 ml of 80% ice-cold
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ethanol and dried using a SpeedVac. The RNA was re-suspended in 400 µL of nuclease-free
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water (Ambion, Life Technologies, Burlington, Ontario, Canada) and pooled together in the
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same tube. The extracted total RNA was treated with Turbo DNase I (Ambion) and purified with
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RNeasy MinElute Cleanup Kit (Qiagen).
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DNA/RNA extraction (deep-sea sediments)
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DNA and RNA were extracted simultaneously from 2 g of sediment using the MoBio RNA
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PowerSoil Total RNA Isolation Kit with the RNA PowerSoil DNA Elution Accessory kit (MoBio
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Laboratories, Carlsbad, CA).
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Metagenomic sequencing
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Each DNA library was prepared for sequencing from 50-100 ng of DNA using the Ion Xpress
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Plus Fragment Library Kit (Life Technologies) with the Ion Xpress Barcode Adapters 1-16 (Life
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Technologies), using the Ion Shear Plus Reagents and a Pippin Prep instrument (SAGE
194
Science, Beverly, MA) for size-selection. Barcoded libraries were pooled in an equimolar ratio
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three by three. A total of 3.50 x 107 molecules were used in an emulsion PCR using the Ion
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OneTouch 200 Template Kit (Life Technologies) and the OneTouch instrument (Life
9
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Technologies). The sequencing of the pooled libraries was performed using the Personal
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Genome Machine (PGM) system with the Ion Sequencing 200 kit and 316 chips (Life
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Technologies). Sequencing statistics are shown in Table S1.
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Metatranscriptomic sequencing
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In order to get enough RNA for library preparation, RNA samples were amplified using the
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MessageAmp II-Bacteria Kit (Ambion) according to the manufacturer's protocol. The antisense
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RNA (aRNA) obtained was subjected to ribosomal RNA subtraction following the procedure of
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Stewart et al. (36) with the exception that the T7 promoter was coupled to the forward primer
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instead of the reverse primer. After subtraction, a 227 bp control RNA transcribed from the
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pSPT18 vector (positions 2867-3104 and 1-70) was added in a 1:1000 ratio (on a nanogram
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basis) to the total rRNA-subtracted RNA. This mixture was then reverse-transcribed using the
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SuperScript III kit (Invitrogen, Life Technologies). Illumina libraries were prepared following the
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protocol of Meyer and Kircher (30), with tags 1 to 34. The indexed libraries were pooled in an
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equimolar ratio and sent for eight lanes of Illumina HiSeq 2000 paired-end 2x100 bp sequencing
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at
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Canada).Sequencing statistics are shown in Table S2.
the
McGill
University
and
Genome
Quebec
Innovation
Centre
(Montreal,
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Bioinformatics
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Metagenomic sequences were submitted to MG-RAST where they were de-replicated using the
217
method of Gomez-Alvarez et al. (8) and trimmed using the dynamic trimming method of Cox et
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al. (5) in a way that each individual sequence would contain a maximum of 5 bases below a
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Phred score of 15. Within MG-RAST, significant matches were defined as having 60%
10
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sequence identity over at least 15 aa or 50 bp and with an e-value below 10-5. Metagenomic
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data was used as relative abundance by dividing the abundance of sequences for a particular
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organisms/gene by the total number of sequence retrieved from the sample. Metatranscriptomic
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data resulted in 544 files (34 samples x 2 reads x 8 lanes). Data from the different lanes were
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pooled together and the resulting 68 files were filtered using a custom-made Perl script, as
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follows. Paired-end reads were processed in parallel. Reads were first trimmed at the first
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occurrence of a low quality base (Phred score below 20) or when the adapter sequence was
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encountered. Following this step, sequences of less than 75 bp were removed from further
228
analyses. If only one of the paired reads was filtered out, then the remaining read was also
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removed. The filtered reads were then submitted to MG-RAST 3.0 (29) where mate-paired
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reads were joined using the fastq-join utility. Mate-paired reads that did not overlap were kept
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for downstream analyses. Within MG-RAST, significant matches were defined as having 60%
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sequence identity over at least 15 aa or 50 bp and with an e-value below 10-5. The number of
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sequences related to the pSPT18 vector in the filtered metatranscriptomic datasets was
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obtained by Blast using an e-value cutoff of 10-25 and this number was used to normalize the
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number of transcripts using the method of Moran et al. (31).
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Statistical analyses
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All statistical analyses were carried out in R (v. 2.13.2, The R foundation for statistical
239
computing, Vienna, Austria). Normal distribution of the data was tested using the “shapiro.test”
240
function. If necessary, data was then transformed using log or square root transformations.
241
Analysis of variance (ANOVA) was performed using the “aov” function while post-hoc Tukey
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honestly significant difference tests were carried out using the “TukeyHSD” function. If these
243
transformations failed to normalize the data, a non-parametric Kruskal-Wallis test was carried
11
244
out in lieu of ANOVA (function “kruskal.test”). Correlation analyses were carried out based on
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Spearman correlation using the “cor” function. Bray-Curtis dissimilarities were calculated using
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the “vegdist” function of the “vegan” library.
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Data deposition
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Raw sequence reads were submitted to the NCBI Sequence Read Archive (SRA) under the
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BioProject accession PRJN0000 (pending) and the SRA project accession SRP0000 (pending).
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Annotated metagenomes (MG) and metatranscriptomes (MT) are available in MG-RAST under
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accessions 4494020.3-4494048.3 and 4494917.3 (MG, water, project 1012) 4500695.3-
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4500711.3 (MG, sediments, project 1891), 4508873.3-4508882.3 (MT, water, project 2384) and
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4508988.3-4509004.3 (MT, sediments, project 2866).
255 256
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257
Results
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The goal of this study was to observe the effects of the DWH spill approximately one year after
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the successful capping of the well. In order to do this, water column samples from a reference
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station that was outside the spill area were compared to water column samples taken at similar
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depths at a station that was directly in the spill area. In addition, deep-sea sediment cores were
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taken in the direct vicinity of the well, and the surface, 5 cm and 10 cm sediment layers were
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compared.
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Chemical analyses and mineralization assays
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The chemical analyses of the water and sediments revealed very low concentration of alkanes
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mostly in the ng per liter of water or ng per g of sediment range (Fig. 2). At these
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concentrations, near the detection limit, variation between replicates was quite high, and the
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only significant difference between the reference and affected water column was between the
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surface water samples (t-test: t=3.53, P