Investigating the Microbial Community Associated with Plastic Marine Debris

Investigating the Microbial Community Associated with Plastic Marine Debris:

An Experimental Colonization Study in the Coastal Waters of Woods Hole, MA, USA

A Senior Thesis Presented to The Faculty of the Department of Organismal Biology and Ecology, The Colorado College By Keven Dooley Bachelor of Arts Degree in Biology 18th day of May, 2015

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Dr. Mark Wilson Primary Thesis Advisor

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Dr. Marc Snyder Secondary Thesis Advisor Introduction

The light weight, durability, and low cost of production of plastic have made it an everyday feature of our lives. In 2013, global plastic production increased by 3.9%, from 288 to

299 million metric tons (PlasticsEurope, 2014). This increase is the continuation of a trend that has been observed since plastic was first mass produced in the 1950’s. Between 1976 and 2013, global plastic production increased by a factor of six (PlasticsEurope, 2013). Accompanying this trend in production is a corresponding trend in plastic waste generation. A 2008 review of U.S. municipal solid waste reported a nine-fold increase in plastic waste generation between 1970 and

2008 (EPA, 2009). Although no reliable estimate of plastic input to the ocean has been established, the significant increase in global plastic production and plastic waste generation suggests the amount of plastic entering the ocean has been increasing over the past several decades.

Floating plastic marine debris was first detected in the western North Atlantic Ocean in the early 1970’s (Carpenter and Smith, 1972; Carpenter et al., 1972; Colton and Knapp, 1974).

These studies reported a wide distribution of plastic fragments and pellets throughout the western

North Atlantic Ocean. However, only within the last decade have studies begun to shed light on the spatial distribution and abundance of plastic debris within the global ocean.

These studies have documented high concentrations of microplastic debris within each of the gyres of the Pacific, Atlantic, and Indian Oceans (Law et al., 2010; Cozar et al., 2014; Law et al., 2014). Ocean gyres are large systems of convergence that occur within the northern and southern regions of the world’s oceans (Dore et al., 2008). The total amount of plastic debris in global, open-ocean surface waters was conservatively estimated to be between 7,000 and 35,000 tons (Cozar et al., 2014). The plastic resins observed in surface waters are predominantly

1 polyethylene, polypropylene, and polystyrene foam (Carpenter et al., 1972; Andrady, 2011;

Oberbeckmann et al., 2014), all of which possess densities lower than that of seawater. These plastic resins are the most commonly used resins for the production of disposable packaging, one of the largest sources of plastic waste (PlasticsEurope, 2013).

In addition to documenting distribution and abundance, several studies have investigated plastic accumulation trends. In the eastern North Atlantic Ocean, Thompson et al. (2004) reported an order of magnitude increase in surface plastic concentration between the 1960s-

1970s and the 1980s-1990s, and no significant increase in abundance between the 1980s and

1990s. These results are supported by another study which found no significant increase in surface plastic abundance in the western region of the North Atlantic subtropical gyre between

1986 and 2008 (Law et al., 2010). In the eastern North Pacific, Goldstein et al. (2012) reported a

2 orders of magnitude increase in plastic abundance between 1972-1987 and 1999-2010.

However, another North Pacific study presenting a larger data set reported a more conservative 1 order of magnitude increase between 1972-1985 and 2002-2012 (Law et al., 2014). Ultimately, these studies suggest there has been less accumulation of plastic debris in the ocean surface than might be expected from the rapid increase in plastic production and waste generation since the

1970s.

The discrepancy between plastic waste generation and ocean accumulation suggests mechanisms exist that remove and sequester a large quantity of plastic marine debris. These mechanisms are likely a complex assortment of biotic and abiotic processes including: photo- oxidative degradation and fragmentation, biofouling leading to sinking (Andrady, 2011), degradation by microorganisms (Zettler et al. 2013), and ingestion (Davison and Asch, 2011;

Cozar et al., 2014). Many of the proposed processes can be mediated by interactions with marine

2 microorganisms. Apart from the roles microorganisms play in the eventual fate of plastic marine debris, interactions between microbial communities and plastic debris have the potential to cause significant effects on ocean ecosystems.

The widespread distribution of plastic debris represents an opportunity for colonizing microorganisms to have significant effects across wide areas. One such effect is the transport of non-indigenous or harmful species. The common occurrence and extensive transport of floating plastic debris gives plastic significant potential to act as a dispersal vector (Barnes, 2002; Barnes and Milner, 2004). Studies have documented the introduction and dispersal of non-indigenous bryozoan species via floating plastic debris (Gregory, 1978; Winston, 1982). Additional studies have revealed the potential for plastic to act as a vector for the dispersal of harmful species such as those that contribute to harmful algal blooms (Maso et al., 2003) and potential pathogens such as members of the bacterial genus Vibrio (Zettler et al., 2013).

Apart from the introduction and dispersal of plastic colonizers, the colonization of this novel and abundant substrate may have significant effects on open ocean ecosystems. Goldstein et al. (2012) revealed that the accumulation of plastic debris in the Northern Pacific subtropical gyre (NPSG) since the early 1970s has significantly enhanced oviposition success in the pelagic insect Halobates sericeus by increasing hard-substrate abundance. This increase in oviposition, and thus egg abundance, may cause a shift in energy transfer between pelagic and substrate associated communities as the biomass of a small number of Halobates eggs is equivalent to 9.2-

27.6% of daytime zooplankton biomass in the surface waters of the NPSG (Goldstein et al.,

2012). The rapid introduction and expansion of this novel substrate, a previously limited feature of pelagic ecosystems, may cause substantial shifts in the structure of oceanic ecosystems.

Experimental colonization studies have been employed to investigate the colonization of plastic and structure of the colonizing community. Many studies investigating the process and mechanisms of colonization have employed glass as a substrate. The surface condition of glass and plastic resins are notably different: glass is a hydrophilic, biologically inert material while plastic resins are hydrophobic and bioactive. When introduced to a marine environment, the surfaces immediately experience biochemical conditioning through the adsorption of macromolecules like polysaccharides and glycoproteins (Wahl, 1989; Van Loosdrecht et al.,

1990). Previous studies have reported initial surface conditions (e.g. hydrophobicity, charge, functional groups) influence the composition of the adsorbed chemical layer, which then plays an important role in mediating colonization (Marszalek et al., 1979; Wahl, 1989). This suggests substrates of differing surface characteristics may develop different colonizing communities.

In a recent colonization study, Oberbeckmann et al. (2014) observed significant differences in microbial community structure between glass and polyethylene terephthalate (PET) deployed in the North Sea. Understanding how microbes interact with different colonization substrates is an important part of plastic debris research. If communities observed on plastic marine debris (PMD) are substantially different from those observed on other colonization substrates, extensive colonization of plastic debris may significantly shift the structure of substrate-associated microbial communities present in pelagic systems. This may alter existing interactions and result in novel interactions between substrate-associated and pelagic communities.

The plastic itself may additionally serve as a novel source of energy within ocean ecosystems as plastic debris has been observed to host a suite of potential hydrocarbon- degrading organisms. Most studies investigating the composition of plastic colonizing marine

4 communities have observed the presence of hydrocarbon degrading taxa (Zettler et al., 2013;

Harrison et al., 2014; Oberbeckmann et al., 2014; Reisser et al., 2014). However, few studies report explicit evidence of biodegradation. Zettler et al. (2013) and Reisser et al. (2014) both observed unknown organisms referred to as “pit formers” on plastic surfaces using scanning electron microscopy. These organisms were found interacting with pits in the plastic surface, proposed to have been formed through biodegradation. A study investigating the communities present on plastic samples collected from (as well as incubated within) the North Sea employed

Fourier transform infrared (FTIR) spectroscopy to analyze the molecular structure of the plastic surface for evidence of biodegradation (Oberbeckmann et al., 2014). This study reported no chemical evidence of biodegradation on collected or incubated plastic samples; however, it did detect the presence of potential hydrocarbon-degrading organisms on incubated samples.

The resins that characterize plastic fragments found in surface waters (polyethylene, polypropylene, and polystyrene) are largely resistant to biodegradation (Singh and Sharma, 2008;

Andrady, 2011). These plastics must first experience abiotic degradation through processes such as photodegradation and thermo-oxidative degradation (Shah et al., 2008; Andrady, 2011; Sivan,

2011). These processes cleave the polymer molecules, decreasing the molecular weight of the plastic and forming functional groups that allow for subsequent degradation via the action of bacteria or fungi (Shah et al., 2008; Singh and Sharma, 2008). These processes are active on the shore but retarded in seawater, suggesting abiotic degradation of plastic debris largely occurs on shores or in transit to the ocean (Andrady, 2011). Even after significant weathering of plastic debris, the polymers still have molecular weights generally resistant to biodegradation. This may explain why the only physical evidence of biodegradation was observed on field samples, which

5 would have had more time than incubated samples for abiotic degradation and colonization by a suite of hydrocarbon degraders.

Novel interactions with plastic marine debris may significantly alter the transfer of energy within and between oceanic ecosystems, and result in changes in ecosystem structure.

Understanding what organisms are interacting with PMD is an important step in determining what ecological interactions are taking place. Most studies investigating the colonization of plastic have employed scanning electron microscopy or genetic fingerprinting techniques and limited sequencing analysis to characterize the colonizing communities (Carson et al., 2013;

Harrison et al., 2014; Oberbeckmann et al., 2014; Reisser et al., 2014).

The communities observed on collected and experimentally incubated plastic fragments are complex and diverse, with a high abundance of diatoms, as well as the presence of other eukaryotic organisms such as coccolithophores, dinoflagellates, invertebrates, and eggs of marine insects (Carson et al., 2013; Oberbeckmann et al., 2014; Reisser et al., 2014). Bacteria were also reported in high abundance, and shown by Oberbeckmann et al. (2014) to represent an assemblage primarily dominated by Cyanobacteria. These studies provide important information on the structure of colonizing communities, including similar trends in colonization between oceanic and coastal plastic samples, as well as between collected and experimentally colonized samples. However, studies to date are limited in depth and taxonomic resolution.

In-depth analysis providing highly resolved taxonomic information will further our understanding of the interactions between microbes and plastic debris, and how these interactions may influence oceanic ecosystems. This information can be obtained through next generation sequencing of entire microbial communities. Next generation sequencing of microbial

6 communities employs high-throughput sequencing of the 16S rRNA region of the bacterial genome (Figure 1). This technique is widely used for bacterial identification as the 16S region is present in almost all bacteria, the sequence has hypervariable regions, and the sequence provides enough data for identification in comparison to a database (Janda and Abbott, 2007). Bacterial groups can often be identified to the genus or species level. In addition, this technique can provide some data on eukaryotic members of the community via sequencing of mitochondrial or chloroplast DNA. Zettler et al. (2013) used this technique to analyze the V6 hypervariable region of the bacterial 16S rRNA (Huse et al., 2008), and provide the first profile of the microbial community interacting with plastic collected from the field.

The Zettler et al. (2013) study investigated communities found on plastic collected from the North Atlantic subtropical gyre. They observed communities dominated by diatoms and other phototrophic protists such as dinoflagellates and haptophytes. Heterotrophic organisms such as predatory ciliates were observed as well. Photosynthetic cyanobacteria, including the genera

Phormidium and Rivularia, were the most abundant members of the bacterial community. The bacterial community also featured potential hydrocarbon degrading organisms, identified via their association with environments featuring hydrocarbon pollution (e.g. the Deepwater Horizon oil spill). These organisms belonged to the genera Phormidium, Pseudoalteromonas,

Haliscomenobacter, Devosia, Oceaniserpentilla and members of the family Hyphomonadaceae.

In addition, their results revealed the community observed on plastic debris is significantly different from that of the water column. Their research characterized open-ocean plastic- associated communities as distinct assemblages featuring phototrophic groups and a suite of potential hydrocarbon degraders. Such specific and comprehensive information allows us to better understand how marine microbes are interacting with this novel substrate.

Figure 1. Proposed secondary structure of the 16S ribosomal component of the small-subunit of the prokaryotic ribosome (Yarza et al., 2014). The variable regions are shown in different colors. Hypervariable regions are labeled in bold. This study utilized the V6 hypervariable region.

However, their study only provided a static understanding of the plastic-associated community, with no information as to how long those plastic fragments had been within the marine environment, and limited knowledge of the environmental conditions the colonizing community had developed in. In order to further characterize the colonizing communities, understand how these communities arise and change over time, and how they are interacting with marine ecosystems, we designed an experiment investigating microbial colonization of three plastic resins and a glass reference. We hypothesized that the plastic resins and glass would each develop different microbial assemblages because of differences in surface conditions.

Methods and Materials

Our study investigated the structure of microbial communities colonizing three plastic resins over a six month period. An in situ colonization experiment consisting of polyethylene

(PE), polypropylene (PP), polystyrene (PS), and glass beads (GL) was deployed in the coastal waters of Woods Hole, MA (41°31'28.3"N 70°40'20.9"W, Figure 2). The structure of microbial communities colonizing these substrates over time was analyzed using next-generation sequencing and scanning electron microscopy (SEM).

Incubation Experiment Consumer plastic goods were used for plastic substrates, namely a one-gallon milk jug

(HDPE) and single use drinking cups (PP and expanded PS). The plastic substrates were cut into strips to facilitate sampling. Cubic glass beads were used as a control colonization substrate. All four substrates were securely suspended on nylon fishing line inside of a milk crate (Figure 2a).

Substrates were positioned to reduce contact with one another so as to preserve developing biofilms. The milk crate was covered in plastic netting with 5 mm mesh to prevent macroorganism grazing of biofilms (Figure 2b). A rock was secured to the bottom to keep the 9 experiment consistently oriented in the water column (Figure 2c). The experiment components were washed thoroughly and sterilized with 70% ethanol immediately before initial deployment to remove any biological contaminants.

PS PE

A B C

A Figure 2. Experimental set-up: (A) polyethylene (PE) jug, polypropylene (PP) cups (clear) and polystyrene (PS) cups (white), and glass beads (strung on clothes hanger) secured within the milk crate; (B) plastic netting coating milk crate; (C) rock used to maintain orientation while deployed.

Site Description The experiment was deployed in the coastal waters of Woods Hole, MA off of the Marine

Biological Laboratory dock (Figure 3). The experiment was suspended at a depth of 1-3 m below the surface. The experiment was deployed on July 17th, 2013 and sampled 9 times until January

1, 2014. Within the first three months (representing the extent of available sequence data), water temperature and salinity ranged from 18.7°C to 24°C and 25.7 to 29.6 ppt respectively. During the full six month period (representing the extent of available SEM data), temperature and salinity ranged from 3.6°C to 24°C and 25.7 to 31.2 ppt, respectively.

Figure 3. Experiment location in Woods Hole, MA.

Sample Collection and Preservation A set of time zero samples was collected prior to deployment to serve as controls for

DNA and SEM analysis. Samples of each substrate were collected weekly for one month, then monthly for six months. At each sampling time point, six replicates were collected per substrate

(three replicates for DNA and SEM analysis, respectively) and stored in individual tubes. Each replicate was a 5 mm x 5 mm square of plastic (cut from a strip) or glass bead. Nitrile gloves were worn during collections, and samples of each substrate were recovered with a set of sterilized scissors and forceps. Between substrate types, the collection scissors and forceps were sterilized with 70% ethanol and then rinsed with Milli-Q water. During the sampling process, samples were temporarily stored in 0.2 µm sterile-filtered seawater. While sampling, water temperature and salinity were measured with an YSI Model 55 handheld dissolved oxygen and temperature probe (YSI Inc., Yellow Springs, OH). Samples collected for DNA analysis were

11 placed in Puregene lysis buffer (Qiagen, Valencia, CA) and stored at -20°C. Samples collected for SEM analysis were initially fixed in a 4% paraformaldehyde solution. After 2-23 h the paraformaldehyde solution was replaced with a 50:50 phosphate buffered saline and ethanol solution, and stored at -20°C.

Scanning Electron Microscopy Samples for SEM analysis were prepared for microscopy through an ethanol dehydration series. Samples were placed in 70%, 85%, and 95% ethanol for 10 min each followed by three 15 min rounds of dehydration in 100% ethanol. Following ethanol dehydration, samples were critical point dried using a Samdri 780A (Tousimis, Rockville, MD), then sputter coated with 5 nm of platinum using a Leica EM MED020 (Leica Microsystems Inc., Buffalo Grove, IL).

Prepared samples were imaged on a Zeiss Supra 40VP SEM (Carl Zeiss Microscopy, Thornwood,

NY) and stored in a desiccator when not in the microscope chamber. Each sample was surveyed for microbial life by defining centered horizontal and vertical transects, then imaging five frames selected using a random number generator within each transect (Figure 4). Frames were imaged at a wide view of 250 µm image width and a zoomed view of 50 µm image width to provide accurate counts of larger and smaller microbes. Images were visualized and measured using the image analysis software ImageJ (Schneider et al., 2012). Images were analyzed for organism abundance, size, and type.

Figure 4. Example of horizontal and vertical transects laid over plastic fragment surfaces for diatom abundance analysis. Transects were divided into 250 µm by 250 µm frames, and then randomly selected for analysis. The overlapping central frame was ignored to prevent recounting.

DNA Extraction DNA was extracted using a Gentra Puregene Tissue Kit (Qiagen, Valencia, CA) and associated extraction protocol modified to include a bead-beating step using 0.1 mm glass beads

(MoBio Laboratories Inc., Carlsbad, CA). The presence of extracted DNA was confirmed and quantified using a Quant-iT PicoGreen dsDNA Quantitation Assay. A Nanodrop 2000 (Thermo

Fisher Scientific Inc., Waltham, MA) was used occasionally used to crosscheck measured concentrations and estimate DNA purity (based on 260/280 values).

16S rRNA PCR and Amplicon Sequencing The bacterial V6 hypervariable region of the small-subunit rRNA gene was targeted for two rounds of amplification, non-fusion and fusion, using the following primers:

Forward primer (967F), CTAACCGANGAACCTYACCCNACGCGAAGAACCTTANC CAACGCGMARAACCTTACCATACGCGARGAACCTTACC

Reverse Primer (1064R), CGACRRCCATGCANCACCT

Non-fusion PCR served only to replicate the targeted V6 region. Fusion PCR not only continued to replicate the region, but fused a barcode unique to that sample to all amplicons (Huber et al.,

2007). Amplicons from multiple samples (each representing a specific colonization substrate and time point) were pooled for high-throughput sequencing. This fusion barcoding procedure allowed sequences to be sorted by sample after sequencing.

The first 20 cycle non-fusion amplification was run in triplicate under the following program: (1) hold at 94°C for 3:00, (2) 94°C for 0:30, (3) 60°C for 0:45, (4) 72°C for 1:00, (5) repeat #2-4, 19 times, (6) hold at 72°C for 2:00, (7) hold at 4°C. Amplicons were then pooled for each sample and run in a 10 cycle fusion PCR reaction under the following program: (1) hold at

94°C for 3:00, (2) 94°C for 0:30, (3) 60°C for 0:45, (4) 72°C for 1:00, (5) repeat #2-4, 9 times, (6) hold at 72°C for 2:00, (7) hold at 4°C. The resulting barcoded amplicons were then pooled and sequenced on a HiSeq 1000 (Illumina Inc., San Diego, CA). Sequences were trimmed of adapter and primer sequences, and low quality reads were removed. Taxonomy was assigned through the global alignment for sequence taxonomy (GAST) process (Huse et al., 2008). This process employs BLAST to compare sequences to a V6 reference database derived from a 16S rRNA database. The top 100 hits were then aligned with the target sequence using MUSCLE, and the matches with the minimum pair-wise difference were selected. For each V6 reference sequence

14 selected, the source 16S rRNA sequence was selected and identified. Taxonomy was then assigned based on a consensus agreement of the 16S reference sequences.

Data Analyses and Statistical Methods The online software suite visualization and analysis of microbial population structures

(VAMPS) was used to calculate alpha diversity, beta diversity, and produce taxonomy bar graphs (Huse et al., 2014). The statistical software Minitab 17 (Minitab, Inc., State College, PA) was used to analyze alpha diversity data. Microsoft Excel (2010) was used to sort taxonomic information to identify overlap between substrates. Venn diagrams were created with Venn

Diagram Plotter (http://ncrr.pnnl.gov/).

Results and Discussion

16S rRNA sequence data and SEM micrographs were used to characterize and compare colonizing communities as they developed over time. We were particularly interested in observing patterns of development and identifying organisms representing novel ecological interactions (e.g. hydrocarbon-degrading organisms). We hypothesized that the plastic resins and glass would develop different microbial assemblages because of differences in surface conditions.

16S rRNA sequencing

A total of 51 samples were successfully sequenced. For each colonization substrate, a range of 1, 2 or 3 replicates were successfully sequenced (some replicates failed to yield sequence data) for six time points, representing three months of incubation. Taxonomic information and sequence counts for each unique sequence were used to analyze alpha diversity, community structure, and beta diversity over time.

Alpha Diversity and Community Structure

Alpha diversity was analyzed using the Shannon Diversity Index (Hill, 1973). This measure of biodiversity considers species richness as well as evenness. Mean Shannon diversity values show a difference in observed diversity between substrates and a slight increase in diversity over time (Figure 5). Over the three month period, Shannon diversity observed on polyethylene samples was on three occasions substantially higher (day 57) or lower (days 7 and

14) than that of the other resins and glass. In addition, diversity in the microbial community on polyethylene appeared to increase over time. Glass exhibited similar trends. Diversity values observed on glass at days 28 and 84 were substantially greater than values observed on plastic resins. Additionally, diversity in the microbial community observed on glass appeared to increase over time. The Shannon diversity values of communities observed on polypropylene and polystyrene were fairly constant, and did not suggest an increase over time.

Figure 5. Mean Shannon diversity (±SD) over time across all substrates. Polypropylene (PP), polyethylene (PE), polystyrene (PS), and glass beads (GL). Values without error bars are not means and represent occasions when only one replicate was successfully sequenced and available for analysis.

These trends across substrates and over time were assessed with a post-hoc repeated measures ANOVA test. The test yielded P-values of 0.423 and 0.062 for the effects of substrate and time, respectively. There was no significant difference in diversity between substrates.

Although the P-value corresponding to alpha diversity over time is statistically insignificant

(0.062), this may be due to the small sample size of this study. These results may show an ecologically relevant increase in alpha diversity over time. Ultimately, these results show the substrates are rapidly colonized by a diverse group of organisms in as little as seven days. This first week of colonization is an important subject for future research. Investigating the organisms that colonize during this early time period will enhance our understanding of how organisms initially interact with marine surfaces, and what differences in community structure may arise between substrates.

Although we observed no significant change in alpha diversity over time, this does not indicate the communities were not changing. A set of relative abundance bar graphs was used to characterize changes in community structure over time (Figure 6). Each graph displays sequence data from an individual colonization substrate replicate. Samples further down the Y-axis represent samples collected after a longer period of incubation. Each sample contains sequence data of one to three replicates. Within each replicate, every color represents a distinct taxonomic group. The relative width of each colored region represents the relative abundance of that group, normalized by percent. This is required due to variability in amplification and sequencing success for each replicate. Only taxonomic groups representing ≥1% of normalized reads appear on the bar graphs. The total number of reads sequenced and the number of reads displayed in the bar graph are listed next to the sample information.

Bacteria Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Bacteria Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Wenxinia Bacteria Bacteroidetes Sphingobacteria Sphingobacteriales Saprospiraceae Haliscomenobacter Bacteria Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Bacteria Proteobacteria Alphaproteobacteria Caulobacterales Hyphomonadaceae Maricaulis Bacteria Proteobacteria Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Bacteria Proteobacteria Alphaproteobacteria Caulobacterales Hyphomonadaceae Robiginitomaculum Bacteria Proteobacteria Alphaproteobacteria Rickettsiales SAR11 Pelagibacter Bacteria Cyanobacteria Cyanobacteria Bacteria Proteobacteria Alphaproteobacteria Rickettsiales SAR11 Bacteria Cyanobacteria Cyanobacteria SubsectionIII Unassigned Phormidium Bacteria Proteobacteria Gammaproteobacteria Vibrionales Vibrionaceae Vibrio Bacteria Cyanobacteria Cyanobacteria SubsectionI Unassigned Prochlorococcus Bacteria Proteobacteria Gammaproteobacteria Xanthomonadales Sinobacteraceae Bacteria Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Loktanella Bacteria Proteobacteria Gammaproteobacteria Bacteria Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Sulfitobacter Bacteria Verrucomicrobia Verrucomicrobiae Verrucomicrobiales Verrucomicrobiaceae Bacteria Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Organelle Chloroplast Bacillariophyta

Figure 6. Bar graphs displaying relative abundance over time of taxonomic groups representing ≥1% of reads per replicate. For each replicate, the total number of reads and number of reads represented in the bar graph are listed next to the substrate and time point information. The legend below the graphs displays the specific colors associated with each taxonomic group.

Considerable species turnover occurred over the course of the experiment (Figure 6). The communities at early time points were dominated by a few common taxonomic groups. Among these taxa, the class Bacillariophyta was especially abundant early on. This class of unicellular eukaryotes, commonly referred to as diatoms, are important primary producers estimated to contribute up to 45% of global oceanic primary production (Mann, 1999). Across all substrates, diatoms dominated the seven day communities, and continued to represent major portions of the

14 day communities observed on PP and PS. However, past these early time points, the relative abundance of diatoms steeply declined. This high abundance of diatoms supports the results observed by other studies investigating plastic colonizing communities (Carson et al., 2013;

Zettler et al., 2013; Oberbeckmann et al., 2014; Reisser et al., 2014).

As the relative abundance of diatoms decreased, the relative abundance of heterotrophic bacterial groups saw a marked increase. Of particular note is the SAR11 clade, including the genus Pelagibacter, and the families Sinobacteraceae and Verrucomicrobiaceae. These three groups exhibited a significant increase in relative abundance over the course of the experiment.

Bacteria belonging to the SAR11 clade are highly abundant oligotrophic scavengers (Morris et al., 2002). This group was observed in low abundance by Zettler et al. (2013), and thought to have become passively associated with PMD through the in-field collection process. In addition,

Oberbeckmann et al. (2014) reported no colonization by members of this clade. However, our results suggest bacteria belonging to the SAR11 clade may in fact colonize plastic surfaces. The bacteria of the family Sinobacteraceae are heterotrophs closely related to the hydrocarbon degrading genus Hydrocarboniphaga (Zhou et al., 2008). Verrucomicrobiaceae is a family of chemoheterotrophic bacteria associated with polysaccharide degradation (Cardman et al., 2014).

The increase in relative abundance of these groups represents a shift in community structure

19 from a community dominated by rapidly colonizing autotrophs to one featuring a suite of heterotrophs of similar relative abundance.

This shift in community structure and decline in diatom population may be associated with the seasonal lifecycle of diatoms. Diatoms, and most other phytoplankton, experience a minor autumnal bloom followed by a decline in population as water temperatures decrease in winter (Colebrook, 1979). However during the first month of incubation, the period of time where diatom abundance sharply declined, temperature only ranged from 22.9°C to 24°C.

Additionally, recent colonization studies spanning a variety of time frames have reported this trend of initially high diatom abundance followed by a decline (Zettler et al. unpublished data).

This suggests rapid colonization by diatoms, followed by a decline in abundance, is a common pattern of development in plastic colonizing communities. This initial colonization and subsequent decline of diatom population likely plays a role in conditioning the substrate surface for subsequent bacterial colonization by providing an ample source of degradable biomass.

Another group of particular interest is the bacterial family Rhodobacteraceae. This group was observed in considerable abundance on all substrates at every time point. Several genera belonging to this family were frequently identified: Loktanella, Sulfitobacter, Roseovarius,

Wenxinia, Phaeobacter. Previous studies have identified bacteria of this family as common colonizers of plastic (Zettler et al., 2013; Oberbeckmann et al., 2014). The bacteria belonging to this group are rapid colonizers capable of colonizing substrates with a range of surface conditions (Dang and Lovell, 2000). The activity of these bacteria is thought to play an important role in facilitating subsequent colonization by developing a primary biofilm (Dang et al., 2008).

This production of extracellular polymeric substances (EPS) and other metabolites conditions the surface for successive colonization (Wahl, 1989). Rapid colonization and surface conditioning

20 by this group and the diatoms likely facilitated the subsequent colonization and proliferation of heterotrophic groups.

Although phototrophic eukaryotic groups were observed in high abundance, phototrophic bacteria were less abundant than expected (Figure 6). Prior studies have observed plastic colonizing communities featuring high abundances of cyanobacteria, including the genera

Phormidium and Rivularia (Zettler et al., 2013), and Pseudophormidium and Stanieria

(Oberbeckmann et al., 2014). However, we infrequently detected the presence of cyanobacteria, detecting low abundances of cyanobacteria on only 13 replicates. The cyanobacterial community detected on the plastic samples was dominated by the genus Prochlorococcus, a group generally thought to be free-living. Prochlorococcus was detected on 10 of the 13 cyanobacteria harboring replicates. This genus was detected in low abundances by Zettler et al. (2013), but was not detected by Oberbeckmann et al. (2014). Our observations may indicate Prochlorococcus occasionally colonizes surfaces, or may be the result of this abundant pelagic bacteria becoming passively associated with plastic samples while collection occurred. The low level of phototrophic bacterial abundance detected in this study may be related to the light levels the experiment was exposed to. The plastic mesh enclosing the experiment was frequently fouled by algae. The algae were removed during sampling, but this fouling may have restricted light availability between longer sampling periods.

The presence of potential hydrocarbon degrading organisms is an important facet of research concerning plastic-associated microbial communities. We detected several potential hydrocarbon degraders, including the bacterial genera: Devosia, Haliscomenobacter,

Hyphomonas, Marinobacter, Oceaniserpentilla, and Phormidium. Marinobacter, a genus containing oil-degrading species (Yakimov et al., 2007), was detected on 2 replicates in

21 substantial relative abundances (greater than 1% of the total community). The genera

Phormidium and Haliscomenobacter were both detected by Zettler et al. (2013). Our study saw a low occurrence of the group Phormidium; however the genus Haliscomenobacter was detected in substantial relative abundance on all substrates throughout the course of the experiment. The genera Devosia and Oceaniserpentilla, both observed by Zettler et al. (2013), were detected in this study, but infrequently and at low abundances. The genus Hyphomonas, found in high abundance across samples analyzed by Zettler et al. (2013), was frequently detected in our study, but at consistently low abundances. The differences in observed hydrocarbon degraders between the results of Zettler et al. (2013) and this study may arise from differences in study site, as well as differences in the degree of weathering between field samples and the post-consumer plastics we used. What is important to note, is the consistency of these studies (and similar studies) in reporting a suite of potential hydrocarbon degraders associated with PMD. This consistency suggests plastic debris associated communities commonly feature biodegradation, and highlights the interaction as an important line of future research.

Beta Diversity

Beta diversity is used to measure the dissimilarity between two communities. We used the Bray-Curtis dissimilarity index, which measures the difference between two communities based on species membership and abundance (Faith et al., 1987). Beta diversity analysis was used to assess whether the communities observed on the different plastic resins were as different from each other as they were from glass. This effectively tested whether or not the plastic resins influenced the structure of the colonizing community. We hypothesized the microbial communities observed on plastic would be more similar to each other (i.e. have lower

22 dissimilarity index values) than to the glass microbial community. Additionally, we hypothesized the communities observed on the different plastic resins would be different from each other.

Figure 7. Beta diversity analysis comparing dissimilarity between a specific resin and the other resins, or that resin and glass (±SD). For each resin, each value represents the average dissimilarity between all appropriate replicates. For example, polypropylene (PP) at 7 days represents the average dissimilarity calculated from how different each PP replicate is to the other resin replicates (polyethylene (PE) and polystyrene (PS)), or to the glass replicates. Bray- Curtis dissimilarity values were used as this metric takes into account abundance data.

We assessed beta diversity between plastic and glass samples by plotting the mean dissimilarity between both a single resin and the other plastic resins, and that resin and glass

(Figure 7). For example, the top left graph displays the mean dissimilarity between the polypropylene (PP) community and the communities on the other plastic resin replicates

(polyethylene (PE) and polystyrene (PS)) over time. This graph additionally displays the mean

23 dissimilarity between the PP community and the glass (GL) community. For these comparisons of beta diversity, the difference between PP and GL is only greater than that between PP and

PE/PS at 28 and 84 days. These results are also observed in the graphs that individually plot polyethylene and polystyrene in relation to the other resins and glass. These results show the communities observed on the plastic resins are as different from one another as they are from the community observed on glass. The communities observed on plastic and glass substrates exhibited some differences, but were largely similar in structure, suggesting the plastic substrates do not substantially influence community structure.

This conclusion is further supported by the large overlap of identified taxa between substrates (Figure 8). The Venn diagrams below display taxa richness (the number of taxa detected) and overlap in taxa occurrence between substrates after one month and three months of incubation. A large overlap in community membership was observed between the different plastic substrates, as well as between the plastics and glass. These figures do not provide information on the abundance of these groups; however, relative abundance bar graphs reveal taxa shared between substrates feature similar relative abundances (Figure 6). This further demonstrates that during this period of colonization, the communities observed across the substrates were largely composed of the same taxa in similar abundances.

Based on prior research (Oberbeckmann et al., 2014) as well as the differences in surface conditions between glass and plastic, we hypothesized the communities observed on glass and plastic would exhibit distinct structures. However, colonization studies have suggested the initial adsorption of macromolecules and other processes result in a convergence of conditions across materials initially possessing different surface conditions. This in turn results in a convergence of colonizing communities. The adsorption of macromolecules on hydrophobic or hydrophilic

24 surfaces results in a convergence of surface free energy, effectively making hydrophobic surfaces less hydrophobic and hydrophilic surfaces less hydrophilic (Baier, 1981; Van

Loosdrecht et al., 1987; Wahl, 1989). In addition, both glass and plastic resins have been shown to develop slight negative charges after this initial adsorptive conditioning (Fletcher and Loeb,

1979).

Figure 8. Venn diagrams displaying overlap in community membership between substrates after one month and three months of incubation. Numbers within the differently colored regions and in parentheses outside the Venn diagrams indicate number of taxa within that group. This convergence of surface conditions is thought to allow groups of flexible early colonizers (taxa capable of colonizing substrates with different surface conditions) to abundantly colonize surfaces exhibiting slight differences in conditions (Fletcher and Loeb, 1979). This is supported by the high abundance of members of the bacterial family Rhodobacteraceae, which

25 have been shown to abundantly colonize substrates of different surface conditions (Dang and

Lovell, 2000; Dang et al., 2008). The similarity in community structure across substrates is likely due to this convergence of surface conditions and rapid colonization by flexible colonizers.

Although the communities present on the different substrates were predominantly colonized by many of the same taxa in similar relative abundances, it is important to note differences in community membership existed between the substrates. At every time point, differences in structure existed between substrates and each substrate featured some unique taxa.

When analyzing all taxa detected on each substrate over the first three months, it was found each substrate was colonized by unique taxonomic groups, with the plastic substrates featuring more than glass. Polyethylene, polypropylene, and polystyrene hosted 45, 47 and 46 unique taxa respectively, while glass only hosted 27 unique taxa. These taxa represent 4.7%, 4.6%, 4.5% and

2.9% of the total number of taxa observed on polyethylene, polypropylene, polystyrene and glass, respectively.

Preliminary analysis of the unique taxa uncovered genera of the bacterial family

Rhodobacteraceae unique to each plastic resin. Polyethylene hosted the genera Tranquillimonas and Salipiger, polypropylene hosted the genus Leisingera, and polystyrene hosted the genus

Silicibacter. Of the two species within the genus Salipiger, the type species Salipiger mucescens is notable for its production of extracellular polysaccharides (Martínez-Cánovas et al., 2004).

The genus Leisingera is closely related to the genus Phaeobacter, one associated with the production of antibiotics (Martens et al., 2006). After primary colonization by members of the family Rhodobacteraceae, both of these characteristics, the production of extracellular polysaccharides and antibiotic compounds, play integral roles in influencing subsequent colonization and community development (Wahl, 1989). These genera may play important roles

26 in recruiting other substrate-specific taxa. Understanding the presence of substrate specific groups, and what relationships they share with other organisms, is an important avenue of future research.

SEM analysis of community structure

Diatom Abundance

A total of 26 samples, collected over six months of incubation, were imaged with scanning electron microscopy (SEM). Diatom abundance was calculated for 22 of these samples.

For each substrate, abundance during the first two weeks and months 1, 3 and 6 of incubation were calculated. With the exception of the 1 month and 6 month polyethylene samples, only one replicate was analyzed for each substrate per time point. Diatom abundance was high within the first few weeks, but rapidly declined at later time points (Figure 9). This same trend was observed across all substrates. This corroborates the trend observed in the sequencing results and demonstrates the analysis of SEM micrographs can be a valuable tool in assessing overarching patterns in community structure.

Figure 9. Diatom abundance over time, across all substrates. Abundance determined via direct surveys of SEM images. PE values for 28 and 184 days (1 month and 6 months) averaged from two samples (±SD). Abundance data fit with exponential trend lines (PE R2=0.8316, Glass R2=0.4561, PP R2=0.7182, PS R2=0.4072). Community Structure and Ecology The patterns in community development observed through the analysis of SEM micrographs support those identified by the 16S rRNA sequence data. In addition, SEM micrographs revealed the presence of ecologically interesting organisms, structures and associations that our limited sequence data could not detect. These organisms and structures provide evidence of important interactions (such as predation and biodegradation) not detected via sequencing.

Early time point micrographs show the glass and plastic surfaces to be colonized by a diverse assemblage of diatoms and rod-shaped bacteria (Figure 10, A and B). Many rod-shaped bacteria are structurally favored for colonization because they approach surfaces vertically, decreasing electrical repulsion between the negatively charged substrate surface and negatively charged cell surface (Wahl, 1989). These bacteria act as early versatile colonizers, such as

28 members of the family Rhodobacteraceae. In addition, early time point micrographs commonly feature coccolithophores (Figure 10, C). This is a group of unicellular, eukaryotic phytoplankton that plays an important role in primary production and the cycling of carbon (Rost and Riebesell,

2004). These micrographs reveal early time point communities dominated by autotrophic eukaryotes, thus supporting the molecular data.

Figure 10. SEM micrographs characteristic of early time points. A. Diverse assemblage of diatoms on polyethylene at 7 days. B. Diatoms and rod-shaped bacteria on polyethylene at 7 days. C. Coccolithophore on polyethylene at 7 days.

Late time point micrographs reveal a decline in the diatom population and rise in abundance of bacterial groups and larger eukaryotic organisms. Broken or halved diatom frustules commonly featured heterotrophic bacteria or other microorganisms (Figure 11, A) suggesting the presence of microbial grazers and predators. This also supports the increase in abundance of heterotrophic bacterial groups observed in the molecular data, and suggests diatoms play an important role in facilitating subsequent colonization by providing an ample source of biomass. Larger unicellular and multicellular eukaryotes appear and increase in abundance at later time points (Figure 11, B). These organisms represent an important potential link between substrate-associated and pelagic ecosystems. As suggested by Goldstein et al.

(2012), the increase in floating substrate represented by the accumulation of PMD, and subsequent colonization of said debris, may significantly influence the transfer of energy within pelagic systems. Later time point micrographs begin to feature well developed bacterial biofilms

(Figure 11, C). These structures demonstrate complex microbial communities are developing, which may facilitate processes such as biodegradation (Wolfaardt et al., 2000; Singh et al., 2006).

The presence of developed biofilms, in concert with the suite of potential hydrocarbon degrading organisms, strongly supports the possibility of biodegradation occurring. However, no direct evidence of biodegradation (such as the “pit formers” observed by Zettler et al. (2013) and

Reisser et al. (2014)) was observed while surveying SEM micrographs. Ultimately, analysis of

SEM micrographs supported the trends in community structure development revealed by the molecular data, and provided valuable ecological insight.

Figure 11. SEM micrographs characteristic of late time points. A. Bacteria in diatom frustule on polypropylene at 1 month. B. Ciliates on polyethylene at 2 months. C. Bacterial biofilm on polypropylene at 4 months.

Conclusion By further characterizing the microbial communities that colonize plastic surfaces, and observing how these communities develop over time, we have begun to develop an in-depth understanding of how microbes interact with plastic marine debris. Plastic surfaces are quickly and diversely colonized, and experience turnover in community composition. Complex

31 communities featuring significant ecological interactions were observed. We hypothesized colonizing communities would be influenced by the plastic substrates, resulting in plastic communities exhibiting a distinct pattern of community structure. However, we saw that plastics and glass share a similar core colonizing community, and variation in community structure could not be attributed to substrate. The minor differences in community structure between substrates, and the taxa unique to each substrate, are important avenues of future research. Understanding how microbes interact with plastic surfaces in turn enhances our understanding of the ecological implications associated with PMD. The accumulation of PMD within the world’s oceans represents a rapid introduction and expansion of a novel pollutant, as well as a novel marine habitat. Its presence influences the dispersal of organisms, the transfer of energy within oceanic systems, and the health of marine populations. Understanding how organisms of all scales interact with PMD allows us to fully understand the ecological consequences of the plastic within our oceans.

Acknowledgement

I would like to thank Erik Zettler and Linda Amaral Zettler for the opportunity to contribute to their research, as well as their guidance throughout the thesis process. Mark Wilson and Marc Snyder, my advisers at Colorado College, provided invaluable help and support throughout this process. Additionally, I would like to thank the Sea Education Association for providing housing during my research, and the Marine Biological Laboratory for providing me with the opportunity and resources to participate in this research. This work was supported by an NSF Collaborative grant to Erik Zettler (OCE-1155379) and Linda Amaral Zettler (OCE-1155571).

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