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The temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea Caroline A. De Tender, Lisa Inès Devriese, Annelies Haegeman, Sara Maes, Jürgen Vangeyte, André Cattrijsse, Peter Dawyndt, and Tom Ruttink Environ. Sci. Technol., Just Accepted Manuscript • Publication Date (Web): 31 May 2017 Downloaded from http://pubs.acs.org on June 4, 2017
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The temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea
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Caroline De Tender1,2,*, Lisa I. Devriese1, Annelies Haegeman1, Sara Maes1, Jürgen Vangeyte1, André
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Cattrijsse3, Peter Dawyndt2, Tom Ruttink1
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1
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9820 Merelbeke, Belgium
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281 S9, 9000 Ghent, Belgium
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Institute of Agricultural, Fisheries and Food Research (ILVO), Burgemeester Van Gansberghelaan 92,
Ghent University, Department of Applied Mathematics, Computer Sciences and Statistics, Krijgslaan
Flanders Marine Institute, InnovOcean site, Wandelaarkaai 7, 8400 Oostende, Belgium
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*Corresponding author:
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Caroline De Tender
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Address:
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Phone number: +3292722473
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Fax:
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Email address:
[email protected] Ankerstraat 1, 8400 Oostende, Belgium
+32 (0) 9 272 24 29
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Abstract
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Despite growing evidence that biofilm formation on plastic debris in the marine environment may be
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essential for its biodegradation, the underlying processes have yet to be fully understood. Thus far,
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bacterial biofilm formation had only been studied after short-term exposure or on floating plastic, yet a
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prominent share of plastic litter accumulates on the seafloor. In this study, we explored the taxonomic
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composition of bacterial and fungal communities on polyethylene plastic sheets and dolly ropes during
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long-term exposure on the seafloor, both at a harbor and an offshore location in the Belgian part of the
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North Sea. We reconstructed the sequence of events during biofilm formation on plastic in the semi-
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enclosed harbor environment and identified a core bacteriome and subsets of bacterial indicator species
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for early, intermediate, and late stages of biofilm formation. Additionally, by implementing ITS2
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metabarcoding on plastic debris, we identified and characterized for the first time fungal genera on
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plastic debris. Surprisingly, none of the plastics exposed to offshore conditions displayed the typical
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signature of a late stage biofilm, suggesting that biofilm formation is severely hampered in the natural
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environment where most plastic debris accumulates.
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Introduction
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Global production of plastics has reached an annual production of 322 million tons in 2015.1
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Polyethylene (PE) is the most commonly produced plastic in the world and accounts for 64% of plastics
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discarded shortly after use.2 A substantial share of plastics eventually ends up in the oceans and seas,
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with an estimated 5.25 trillion plastic particles currently scattered throughout the oceans.3 In the Belgian
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part of the North Sea, PE plastic sheets and dolly ropes are the dominant types of plastic debris (PD).4
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The hard, hydrophobic surface of PD is an ideal environment for colonization by various biota, including
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diatoms, bacteria, and invertebrate species.4,5,6,7,8 Merely one week exposure to the marine environment
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is sufficient for a microbial biofilm to form on PD.9 Polymer type, environmental conditions and season
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appear to affect the composition of the microbial biofilm.10,11 Furthermore, the bacterial community of
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the biofilm significantly differs from that of the surrounding environment, e.g. seawater and sediment.4
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Based on these unique characteristics, the assemblage of taxa inhabiting the outer surface of PD is often
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referred to as the “plastisphere”.8
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As for most thermoplastics, degradation of PE is extremely slow, and it is therefore expected to persist
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in the marine ecosystem.12 Micro-organisms may contribute to the degradation of PD in the marine
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environment. So far, only a few marine bacterial strains have been identified as potential PE degraders.
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Arthrobacter sp. and Pseudomonas sp. were isolated from high-density PE (HDPE) debris from the Gulf
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of Mannar, a marine coastal area.13 Furthermore, Kocuria palustris, Bacillus pumilis and Bacillus
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subtilis strains were isolated from low-density PE debris originating from the Arabian Sea.14 Recently
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it was shown that also a marine fungus, Zalerion maritimum, has the potential to actively degrade PE.15
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Furthermore, several microbial strains, including bacteria and fungi, were isolated from PD in different
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types of soil environment, and described as potential PE degraders.16 Most studies on PE degradation
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are based on growth on medium with plastic polymers as the sole carbon source, PE mass loss and size
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reduction, and the screening of changes in functional groups by FT-IR. Only few studies described the
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actual degradation, e.g. based on enzyme production by the bacterial strain.17
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The formation of a biofilm, a structured system which facilitates metabolic interaction between cells,18
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can be important in terms of biodegradation. Biofilm formation can increase degradation efficiency of
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pollutants such as diesel oil, and destruction of the biofilm architecture can disrupt interspecies 3 ACS Paragon Plus Environment
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cooperation and interfere with degradation efficiency.19,20 To date, little is known both about the
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temporal dynamics of colonization and biofilm formation on PD and about the microbial interactions
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underlying the resulting biodegradation process. Previous studies on biofilm formation on plastics under
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controlled conditions focused on short-term processes only (< 8 weeks),9,10,21,22,23 or did not include
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taxonomic classification of the bacterial communities.24,25,26,27,28 Most of those studies focused on
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floating plastic litter, whereas the predominant part of PD is located on the seafloor.29,30 Reconstructing
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the temporal dynamics of microbial colonization based on randomly sampled PD proved difficult by
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comparing independent pieces of plastic with unknown history, travel pattern, and duration of exposure
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to the marine environment to each other.4
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We established a long-term exposure time-series experiment in which two types of PE were exposed to
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the Belgian part of the North Sea at two different locations. First, plastics were exposed and sampled at
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the harbor of Ostend, a semi-enclosed environment, with low influence of currents. The presence of
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anthropogenic activity, e.g. waste-pipes, land run-off and oil discharges, and the small median grain size
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of the sediment makes this environment more susceptible to environmental pollution.31 Second, plastics
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were exposed and sampled at the Thornton windmill park, which we will further refer to as the
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“offshore” environment. In this area, currents are stronger, but pollution is less pronounced.
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The aims of this research are fourfold. First, an in-depth study of the biofilm formed on plastic exposed
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to the marine environment on fixed locations was done. Previous research focused on bacterial
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communities,4,8,10,21,23,27 whereas the fungal community has been less intensively studied.23 Therefore,
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we used 16S rDNA and ITS2 metabarcoding in parallel to study the taxonomic composition of bacterial
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and fungal communities, respectively. To study possible factors that affect biofilm formation, we
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compared two types of plastic (sheet or dolly rope) in two different environments (harbor or offshore).
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Second, the temporal dynamics of bacterial and fungal colonization of PD are reconstructed, and we
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identified signature species for early, intermediate or late phases of long-term exposure in the harbor
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environment. This series of microbial colonization in the harbor was used to evaluate biofilm formation
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stages in the offshore samples of the exposure series. In addition, these were compared to bacterial
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communities of previously described4 randomly collected samples exposed to similar offshore
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conditions. Third, possible sources of microorganisms were studied by comparing taxonomic profiles 4 ACS Paragon Plus Environment
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of plastic to those of sediment and seawater. Fourth, we investigated if bacterial and fungal species
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previously identified as potential PE degraders were also present in the biofilm to thus assess whether
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microbial biodegradation in the marine environment may take place.
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Materials and Methods
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EXPERIMENTAL SETUP AND SAMPLE COLLECTION
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From September 2015 until July 2016, PE samples were exposed to the marine environment at two
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different locations in the Belgian part of the North Sea: at the harbor of Ostend (51°13’N, 2°56’E) and
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offshore, at the Thornton windmill park (51° 34’N, 2° 58’ E) (Supplementary (S) Figure S1). These
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locations are characterized by different features (Table S1). Environmental properties (seawater
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temperature, conductivity, pH, oxygen, salinity and density) were measured using the CTD SBE-19plus
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on each sampling date. Sediment organic matter or total organic carbon (TOC) of the upper sediment
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layer (0-5 cm) was measured at the first sampling date for both locations, using the “dichromate method”
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as previously described.32 In addition, concentrations of pollutants in the sediment were compiled from
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previous studies and are listed in Table S1.33,34
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Two types of PE, with different color and shape, were exposed to the two environments: transparent
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plastic sheets (A4 size) (RKW Hyplast, Hoogstraten, Belgium) and orange-colored dolly ropes (ø 1 cm,
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length 20 cm; ø single monofilament 1 mm). Three pieces of each type of plastic, representing three
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biological replicates, were attached to a wooden block that sinks to the seafloor, and which was secured
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in a construction (total length: +/- 60 m, weight +/- 75 kg) comprising a buoy, ropes (ø 16 mm), chains,
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an anchor and concrete weights (Figure S2). At the start of the experiment, sets of thirteen and five
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identical constructions were placed on the seafloor in the harbor and offshore, respectively. Handling of
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these constructions was always done with plastic gloves to avoid contamination. In addition, from three
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randomly chosen constructions a piece of PE sheet and dolly rope were cut with sterilized scissors to
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study the microbial load of plastics at the onset of the experiment, using metabarcoding. This was done
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for both locations.
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At the harbor, one construction per week was pulled up and removed during the first month, and from
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then on one construction per month. This led to thirteen collection dates: 1 (September 2015), 2, 3, 4, 9, 5 ACS Paragon Plus Environment
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14, 18, 22, 27, 31, 35, 40 and 44 (July 2016) weeks after placing the constructions. Offshore, the
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constructions were pulled up and removed on four collection dates: 4 (October 2015), 14, 18 and 22
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weeks (February 2016) after placing them. Upon collection of a sample, which was done with sterile
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forceps, scissors and gloves, half of the plastic was immediately stored at -80 °C for DNA extraction
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and the other half was air-dried and stored at room temperature for the biofilm assay. Offshore, three
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replicate seawater and sediment samples were collected on the same date and location as the
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constructions were sampled, as described in a previous study.4 Per replicate, 1 l seawater was filtered
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through a 0.22 μm Millipore membrane filter (Merck Millipore, Billerica, MA). After collection,
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sediment samples and the membrane filters were stored at -80 °C until further use.
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BIOFILM ASSAY
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The quantitative biofilm assay was used to measure biofilm formation on the plastic sheets.9 Briefly,
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plastic samples (4x5 cm, n=2 per time point x location x replicate, resulting in n=6 per time point x
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location) were rinsed three times with sterile water and air-dried for at least 45 min in sterile Petri dishes.
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These plastics were stained with crystal violet (1% w/v) for 45 min and washed three times with sterile
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seawater. Stained samples were air-dried for another 45 min, cut into four pieces of similar size and
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placed into a 2 ml Eppendorf tube to which 1 ml ethanol (95% v/v) was added. The ethanol was then
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diluted 100-fold in ethanol and transferred to a cuvette to measure the optical density at 595 nm using
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an UV-VIS spectrophotometer (UV-1700 Pharmaspec, Shimadzu, Brussel, Belgium). The optical
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density is directly proportional to the amount of biofilm per surface area on the plastic.
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DNA EXTRACTION
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DNA was extracted from the sediment and plastic samples using the Powersoil DNA isolation kit
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(MOBIO Laboratories, Carlsbad, CA) according to the manufacturer’s instructions. In total, 250 mg of
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sediment, a piece of 2 cm by 2.5 cm (total surface area of ± 10 cm2) of the plastic sheet, or 10 individual
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monofilaments with a length of 2.5 cm of the dolly rope (surface area of ± 10 cm2) were used for DNA
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extraction. Before extraction, plastic was rinsed three times with sterile water to remove sediment
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particles and loosely attached organisms. DNA was extracted from the Millipore filters containing the 6 ACS Paragon Plus Environment
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micro-organisms of seawater as described in a previous study.4 The DNA extracts of all samples were
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stored at -20 °C until further use for amplicon sequencing.
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16S rDNA AND ITS2 AMPLICON SEQUENCING AND SEQUENCE READ PROCESSING
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Amplicon sequencing of the V3-V4 fragment of the 16S rRNA gene and the ITS2 gene fragment using
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Illumina technology (Illumina, San Diego, CA, USA) was done to study both the bacterial and fungal
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communities on PD. DNA fragments were amplified and extended with Illumina specific index adaptors
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using an amplification PCR followed by a dual-index PCR, as previously described.32 Detailed
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information on the library preparation can be found in the supplementary materials. Each PCR reaction
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product was purified using the CleanPCR reagent kit (MAGBIO, Gaithersburg, MD, USA). Libraries
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were quality-controlled using the Qiaxcel Advanced, with the Qiaxcel DNA High Resolution kit
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(QIAGEN, Germantown, MD, USA), and concentrations were measured using the Quantus double-
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stranded DNA assay (Promega, Madison, WI, USA). The indexed libraries of each sample were diluted
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to 10 nM and pooled in a 2:1 ratio for bacterial and fungal libraries, respectively. Resulting libraries
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were sequenced using Illumina MiSeq v3 technology (2 x 300 bp) by Macrogen, South-Korea, using
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30% PhiX DNA as spike-in.
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Demultiplexing of the amplicon dataset and removal of the barcodes was done by the sequencing
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provider. The raw sequence data is available in the NCBI Sequence Read Archive under the accession
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number Bioproject ID SUB2179884 for the bacterial sequences and SRP094681 for the fungal
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sequences. Processing the sequence reads to Operational Taxonomic Unit (OTU) tables was done as
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previously described 32 and described in detail in the supplementary materials.
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The bacterial load of three PE sheets and three ropes was checked at the onset of the experiment before
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exposure to the marine environment. Samples were analyzed by metabarcoding in parallel to all other
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samples. Because the PCR product concentration was low, all DNA was used for sequencing. For each
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sample, the number of reads was < 100.
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DOWNSTREAM DATA ANALYSIS AND STATISTICS
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OTU tables of the 16S V3-V4 and ITS2 amplicon sequencing were analyzed using the QIIME software
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package (v1.9.0).35 Taxonomy was assigned with the script “assign_taxonomy.py” using the Uclust
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method considering maximum 3 database hits, with the Silva v119 97% rep set (as provided by QIIME)
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as reference for the bacterial sequences and UNITE v7 (dynamic) for fungal sequences.36,37,38
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A part of the fungal sequences could not be classified using the UNITE database. These sequences were
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extracted from the total data set and their taxonomy was assigned using BLAST for sequence
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comparison with the non-redundant nucleotide database of NCBI.39 We kept the best hit per query using
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an e-value cut-off of 1e-5 and a minimal percent identity of 98.5%.
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Rarefaction analysis was done using the “alpha_rarefaction.py” script of QIIME. A plateau was reached
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at 10,000 sequences for the bacterial and fungal OTUs (Figure S3 and Figure S4). Richness of the
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bacteria and fungi was determined on rarefied data, for which the number of sequences and the
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calculation of the Chao1 richness index was set on the plateau.
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A core microbiome was calculated separately for the bacterial and fungal communities. Only plastics
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exposed for at least four weeks were considered to calculate a core microbiome to account for some lag-
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time for biofilm build-up. This core microbiome was calculated separately for the two environments.
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OTUs were denoted as core organisms if their relative abundance contributed at least 0.1% to the total
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community of a sample in at least 90% of the plastic items per environment. Calculations for the core
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microbiome were done in R.40 In addition, the genera that were most abundant at the last time point of
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sampling were calculated by selecting those genera with a minimal mean abundance of 1% over the
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replicates for sheets or dolly ropes.
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The multivariate analysis was done using the R package vegan (version 2.0-10).41 The OTU tables of
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bacterial and fungal sequences, as generated by Usearch, were normalized by calculating relative
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abundances. Next, OTUs with a low count number were removed by only retaining the OTUs which
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had a minimal relative abundance of 0.01 % in at least three samples. The dissimilarity matrix, based on
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the Bray-Curtis dissimilarity index, was calculated from this normalized and filtered OTU table, for both
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the bacterial and fungal sequences. The homogeneity of the variances was checked on this dissimilarity
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matrix using the betadisper function. The significance of the factors environment, type of plastic, and
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time, and their various interaction effects were analyzed using PERMANOVA analysis (number of 8 ACS Paragon Plus Environment
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permutations = 1,000) using the Bray-Curtis dissimilarity index matrix as input. Factors were considered
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significantly different if p-value < 0.05.
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Results
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Bacterial and fungal colonization in the harbor environment
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From the first week of exposure onwards, a coating comprising a microbial biofilm, sediment particles,
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algae and macro-fouling (e.g. mussels) was formed on the plastic sheets at the harbor (Figure S5). Using
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a quantitative biofilm formation assay,9 at least part of the coating on the plastic sheets could be
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attributed to a microbial biofilm (Figure 1). This biofilm was already detected after one week of
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exposure, and increased slightly until week 27, followed by a period of stronger growth until week 40.
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The taxonomic composition of the bacterial and fungal community on plastic at the harbor was analyzed
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in detail by 16S and ITS2 metabarcoding. Richness of the samples was studied by estimating the number
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of observed OTUs and the Chao1 index. Both measures showed that the richness of sheets is similar to
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that of dolly ropes; both for bacterial OTUs (Figure S6 and S7 - Harbor) and for fungal OTUs (Figure
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S6 and S7 - Harbor). The number of bacterial OTUs is only slightly higher on dolly ropes compared to
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sheets in the first few weeks of exposure to the harbor environment. At each time point, the bacterial
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richness of plastic (mean around 1500 OTUs) was markedly higher than fungal richness (mean around
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500 OTUs).
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The bacterial community of plastic sheets and dolly ropes at the harbor displayed a gradual change in
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taxonomic composition during the period of exposure (Figure 2A and 2B). This temporal gradient, more
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evident on the plastic sheets, is, at least in part, caused by shifts in abundance of particular bacterial
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classes: an increase in the relative abundance of alpha- and betaproteobacteria and flavobacteria, and a
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decrease in the relative abundance of gammaproteobacteria (Figure 2A and 2B). Alpha- and
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gammaproteobacteria are characteristic for primary biofilm colonization, while bacteroidetes are known
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secondary biofilm colonizers in the marine environment.42,43,44 On the plastic sheets, a gradual decrease
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in the relative abundance of these primary colonizers and an increase in secondary colonizers was
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observed (Figure S8A), suggesting that subsequent time points reflect progressive stages of biofilm
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formation. This shift from primary to secondary colonizers, however, was not as clearly discernible for
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the dolly ropes (Figure S8B).
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Next, we defined a core bacteriome of plastic samples (see Materials and Methods). In total, 25 bacterial
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core OTUs were identified both on plastic sheets and dolly ropes (Table 1). Based on their temporal
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profile, these core members were classified into four groups: (1) OTUs without a clear period of high
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relative abundance (neutral), e.g. Arenicella, Methylotenera; (2) OTUs with higher abundance in the
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beginning (early stage; week 1-14) of the exposure period, e.g. Sulfurovum, Maritimimonas; (3) OTUs
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with higher abundance in the middle (intermediate; week 14-35) of the exposure period, e.g.
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Robiginitomaculum, and (4) OTUs with highest abundance at the end (late stage; week 35-44) of the
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exposure period, e.g. Sulfitobacter, Psychroserpens (Table 1; Figure S9; Figure S10).
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Next, the fungal community on the plastic sheets and dolly ropes in the harbor was studied. Strikingly,
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the majority of the fungal sequences (28% to 97% of the reads per sample) could not be assigned using
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the UNITE database (Figure 3A and 3B). Using NCBI Blast, some of those reads could be assigned to
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fungi, others to other members of the eukaryotes, e.g. Paramoeba permaquidensis, Paramoeba
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aestuarina, Pleurobrachia pileus, Sugiura chengshanense, Sagartia elegans, and Rhizostoma pulmo,
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but the vast majority remained unassigned (Table S2). Within the share of the fungal sequences that
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were assigned to a certain taxonomy, the Ascomycota were highly abundant, followed by a smaller
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fraction of Basidiomycota (Figure 3A and 3B). Zygomycota were also identified, but only represented
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a minor fraction. In addition, genera that were highly abundant on sheets or dolly ropes ( > 1%) sampled
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at the last time point (44 weeks) were studied (see Table S3). Especially members of the
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Lecanoromycetes, e.g. Physconia, Candelariella, and Caloplaca were abundant. No clear temporal
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profile characterized by early, intermediate, and late stage abundance peaks could be identified,
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essentially because the fungal community profile varied considerably, even between successive time
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points (Figure 3A and 3B). In addition, no core group of fungal organisms could be identified,
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illustrating the variability of the fungal community through time.
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Bacterial and fungal colonization in the offshore environment
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Biofilm formation occurred on plastic exposed to offshore conditions, but was much less pronounced
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compared to the harbor environment as described above. For instance, the biofilm layer was hardly
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visible by the naked eye even after 22 weeks of exposure (Figure S5), and the amount of biofilm that
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had accumulated after 22 weeks of exposure offshore was similar to the amount that had already
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accumulated at the harbor after 1 week of exposure (Figure 1). Until week 18 of the exposure period,
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the number of unique bacterial OTUs and the Chao1 index, both representing the richness of the samples,
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on the plastic sheets sampled offshore remained low ( < 1000 OTUs; Chao1:
