It`s all about the base

Marine biofilms in the plastic age

Dissertation

Zur Erlangung der Würde des

Doktors der Naturwissenschaften

- Dr. rer.nat. –

Dem Fachbereich Biologie/Chemie der

Universität Bremen

vorgelegt von

Inga Vanessa Kirstein

Bremen

März 2019

Die vorliegende Arbeit wurde in der Zeit von Juli 2014 bis März 2019 an der Biologischen

Anstalt Helgoland, Alfred-Wegener-Institut Helmholtz Zentrum für Polar und

Meeresforschung angefertigt.

1. Gutachter: PD Dr. Bernhard Fuchs

2. Gutachter: Prof. Dr. Rudolf Amann

1. Prüfer: Prof. Dr. Ulrich Fischer

2. Prüfer: Dr. Gunnar Gerdts

Tag des Promotionskolloquiums: 3. Mai 2019

„Was immer du tun kannst oder träumst es zu können, fang damit an!

Mut hat Genie, Kraft und Zauber in sich.“

Johann Wolfgang von Goethe

TABLE OF CONTENTS

GENERAL INTRODUCTION 1

OBJECTIVES 12

OUTLINE 14

CHAPTER I 17

Mature biofilm communities on synthetic polymers in seawater - Specific or general?

CHAPTER II 39

The Plastisphere – Uncovering tightly attached plastic “specific” microorganisms

CHAPTER III 61

Dangerous Hitchhikers? Evidence for potentially pathogenic Vibrio spp. on

microplastic particles

GENERAL DISCUSSION 81

FUTURE PERSPECTIVES 95

SUMMARY 99

ZUSAMMENFASSUNG 101

SUPPLEMENT 105

REFERENCES 173

ACKNOWLEDGEMENTS 187

INTRODUCTION

1

INTRODUCTION

Living in the plastic age

Since the 1970s, plastic has become an indispensable material in industries and is present in

every aspect of modern life. Plastics are inexpensive, durable, lightweight, strong, and

corrosion-resistant (Thompson et al., 2009). The word "plastic" derivates from the Greek word

“plastikos” which means “to mold”, and refers to the malleability of a material during its

manufacture into all imaginable forms (O'Brien, 2009). Plastics are derived from organic

products, like natural materials such as crude oil, coal, and natural gas (PlasticsEurope, 2016).

Due to their better chemical and physical properties, lower costs and durability, the annual

usage of plastics in packaging has replaced cellulose-based materials and increases by

approximately 25% per year (Jayasekara et al., 2005). Plastics can be differentiated into two

main categories, thermoplastics and thermosets. The characteristics of thermoplastic, including

e.g. polyethylene (PE), polypropylene (PP), polyvinyl chloride (PVC), polyethylene

terephthalate (PET) and polystyrene (PS) are reversible, meaning that it can be heated and

reshaped repeatedly. Thermosets on the other hand, including e.g. unsaturated polyesters,

silicone and polyurethane (PUR), cannot be reformed after they were heated. The chemical

composition (e.g. polyesters, polyolefines) and physico-chemical properties of the various

plastic types within these two categories is highly diverse in order to meet the different needs

of thousands of end products (PlasticsEurope, 2018). Their broad application in packaging

technology, constructions, and other industries leads to a current global annual production of

350 million metric tons in 2017 (PlasticsEurope, 2018). Six types of synthetic polymers

including high-density polyethylene (HDPE), low-density polyethylene (LDPE), polyvinyl

chloride (PVC), polystyrene (PS), polypropylene (PP) and polyethylene terephthalate (PET)

make up 90% of the plastics produced worldwide (Andrady and Neal, 2009). Consequently,

these synthetic polymers are also among the most commonly detected plastics in the

environment (Andrady, 2011; Engler, 2012).

Plastic litter in the marine environment

Nowadays, there are multiple sources and pathways of plastic litter into the ocean (Fig 1) but

by far, improper disposal of plastics represents the most rapidly growing form of litter entering

and accumulating in the oceans (Andrady, 2011; Thiel and Gutow, 2005). In numbers, Jambeck

et al. (2015) calculated that of 192 coastal countries in 2010, 4.8 to 12.7 million MT of plastic

waste was entering the ocean. Additionally, marine plastic litter is entering the marine

INTRODUCTION

2

environment via waterway-based sources like e.g. nets from commercial fishing (Li et al.,

2016). Eriksen et al. (2014) estimated that more than 5 trillion pieces of plastic, weighing

approximately 270.000 tons, float through the oceans. The longevity of plastics in the marine

environment is a matter for debate, and estimates range from hundreds to thousands of years,

depending on the chemical and physical properties of the plastic type (Barnes et al., 2009).

Indeed, plastics remain much longer in the marine environment than most natural substrates

and are getting dispersed by wind and currents (Barnes et al., 2009), making it difficult to

determine their origin. Consequently, marine plastic litter of unknown age and origin can be

found in marine waters all over the globe.

Fig 1 Pathways of plastic litter into the ocean (Image: Alfred-Wegener-Institut / Martin Künsting (CC-BY 4.0)).

Due to their durability and the prevailing conditions in seawater (e.g. cool temperatures and

low UV radiation), most plastic types are poorly degradable in the marine environment, (Barnes

et al., 2009; Colton et al., 1974), but, rather, become brittle over time and subsequently break

down into smaller fragments, so called microplastics (Andrady, 2011; Corcoran et al., 2009).

While several size categorizations have been suggested for plastics (Gregory and Andrady,

INTRODUCTION

3

2003; Moore, 2008), microplastics generally refer to plastic fragments smaller than 5 mm

(Arthur et al., 2009; Barnes et al., 2009).

Plastic types such as PE or PP float on seawater surface, while e.g. PVC, PET and PS, are

denser than seawater (ρ ~ 1,025 g/cm3) and sink and accumulate in sediments. However, the

distribution of plastics in the marine environment is also influenced by hydrodynamic

conditions (e.g., wind and wave actions weathering and biofouling) (Ballent et al., 2013;

Browne et al., 2010; Moret-Ferguson et al., 2010). Consequently, (micro)-plastics are detected

worldwide in various marine environments (Cole et al., 2011; Eriksen et al., 2014), ranging

from surface waters (Sadri and Thompson, 2014; Thiel et al., 2003) to sediments, and from the

beach (Stolte et al., 2015) to the deep-sea (Bergmann et al., 2017). Interestingly, particularly

high concentrations of plastics were found in sea ice in remote polar regions (Peeken et al.,

2018) and in marine organisms due to ingestion (Rummel et al., 2016) (Fig 1).

Plastics represent a major threat for marine organisms, mainly due to ingestion and

entanglement of ghost nets and larger plastic items (Galgani, 2015; Gregory, 2009). The

presence and increasing accumulation of plastics in the ocean have severe implications. For

example, because of their hydrophobicity, plastics adsorb toxic metals and persistent organic

pollutants (Ashton et al., 2010; Holmes et al., 2012). Furthermore, due to its persistency plastic

serves as potential accumulation site and vector for the dispersal of pathogens (Keswani et al.,

2016; Zettler et al., 2013). The ingestion of small plastic items by marine organisms can lead

to the transport of even those, their accumulated toxins and associated pathogens, to higher

trophic levels in the food web (Keswani et al., 2016; McCormick et al., 2014). Consequently,

plastics and their associates might end up in the human gastro-intestine. The entry of plastics in

the food web is also alarming, since it has been demonstrated that even smaller fragmented

plastics, so-called nanoplastics (< 1 μm), are able to penetrate cell membranes in fish (Oryzias

latipes). Nanoplastics have been detected in the gills, intestine, blood, liver, and in the brain of

fish (Kashiwada, 2006).

Overall, in addition to aesthetic aspects, plastic pollution represents a major yet unpredictable

threat to nature and its consequences are far from being understood.

Biofilms – Sticking together for success

As any surface in the marine environment, plastics are rapidly colonized by microorganisms

(Harrison et al., 2014) and subsequently by a myriad of organisms building up complex biofilms

(Dobretsov et al., 2010). Biofilms are defined as an assemblage of microbial cells that is

irreversibly associated with a surface and enclosed in primarily extracellular polymeric material

INTRODUCTION

4

(Donlan, 2002). Biofilms are, metaphorically speaking, a “city of microbes” (Watnick and

Kolter, 2000). Extracellular polymeric substances (EPS) represent the “house of the biofilm

cells” (Flemming et al., 2007). Although every biofilm is unique in composition and

functionality, biofilm development follows a general pattern (Artham et al., 2009; Bravo et al.,

2011) that determines the final characteristics of a biofilm (Boland et al., 2000; Gottenbos et

al., 2002; Lobelle and Cunliffe, 2011). At the onset of the biofilm formation, the substrate

surface is covered by a conditioning layer created by the adsorption of dissolved organic

molecules. Since the first colonizers adhere to the conditioning layer and not to the substrate

itself, the structure and composition of this layer define the strength of the initial biofilm. Then,

the attachment of bacterial cells, followed by the excretion of EPS, make the reversible adhesion

irreversible (Boland et al., 2000). Subsequently, the initial biofilm expands, forming a habitat

(Fig 2). Finally, unicellular eukaryotes attach, followed by larvae and spores (Dobretsov, 2010).

This biological assembly is kept together by the biofilm matrix. Complex biofilms include a

heterogeneity in form of organisms with various metabolic capacities and physiologies which

generates on the one hand competition but also provides on the other hand opportunities for

cooperation within the biofilm habitat (Fig 2) (Flemming et al., 2016). Bacteria in biofilms are

known to exhibit enhanced resistance to antibiotics and other types of stress compared to their

planktonic forms (Salta et al., 2013), underlining biofilms as a successful strategy of life

(Flemming and Wingender, 2010).

Fig 2 Emergent properties of biofilms and habitat formation adapted from (Flemming et al., 2016)

The biofilm Matrix serves as the “cement” of the biofilm enclosing cells, water, ions and

soluble low- and high-molecular mass products. This matrix holds functions such as protection

(Oliveira et al., 1994) which is ensured by maintaining a highly hydrated layer around the

INTRODUCTION

5

biofilm, hence preventing lethal desiccation (Sutherland, 2001). Further properties include

localized gradients (e.g. oxygen, pH) that provide habitat diversity within the biofilm and

resource capture by sorption of nutrients. The EPS connects the cells and acts as an external

digestive system by keeping the extracellular enzymes in close proximity to the cells (Flemming

et al., 2016). This enables the cells to metabolize both, dissolved and solid biopolymers

(Flemming and Wingender, 2010).

Marine biofilm formation on artificial surfaces is commonly considered as problematic. The

practical consequence of colonisation by marine organisms is biofouling. Biofouling refers to

the unwanted accumulation of biological material on man-made surfaces (Flemming et al.,

2009) leading to impairment or biological degradation, consequently resulting in high costs of

maintenance of even those materials (Callow and Callow, 2002). The most diverse and

important microorganisms within marine biofilms, in terms of composition, dynamics, and

function are bacteria (Dang and Lovell, 2016). The composition and dynamics of mature

biofilm communities may be already defined in the very early stage of biofilm development,

by pioneer microbes sensing the surface of a substrate (Dang and Lovell, 2016). Marine bacteria

are known to prefer either a free-living or a surface-associated lifestyle, although some species

may switch their preference under certain environmental circumstances or life stages (Dang and

Lovell, 2016; Salta et al., 2013). Several groups of bacteria are known to be frequently surface

associated in marine environments, like Rhodobacteraceae (Alphaproteobacteria),

Alteromonadaceae and Vibrionaceae (Gammaproteobacteria), as well as Bacteroidetes

(mainly Flavobacteria) (Dang and Lovell, 2016) representing “general” surface colonizers.

The “Plastisphere”

Because they are physically and chemically distinct from naturally occurring substrates, plastics

offer a unique type of substrate to the microbial community. Zettler et al. (2013) coined out the

term “Plastisphere”, showing that these microbial communities on marine plastics differ

consistently from the surrounding seawater communities of the North Atlantic Ocean. At the

onset of this PhD thesis in 2014, the work of Zettler et al. (2013) was the first study published

using a culture-independent next generation sequencing approach in order to explore microbial

communities on marine plastic litter. In the following years, there has been a growing concern

about the ecological impact of plastics and its Plastisphere on the marine environment and

researchers all over the globe started exploring the Plastisphere in various locations (Amaral-

Zettler et al., 2015; Bryant et al., 2016; De Tender et al., 2017; De Tender et al., 2015; Debroas

et al., 2017; Oberbeckmann et al., 2014; Oberbeckmann et al., 2016). Oberbeckmann et al.

INTRODUCTION

6

(2014) found that the composition of biofilm communities present on plastics in marine habitats

is driven by spatial and seasonal effects, but also varies with the plastic type of randomly

sampled plastics in the North Sea. Amaral-Zettler and colleagues (2015) reported that

Plastisphere communities of the Atlantic and Pacific Ocean clustered to a greater extend by

geography than by plastic type. Bryant et al. (2016) described taxonomically distinct plastic

communities compared to their planktonic counterparts in the North Pacific Subtropical Gyre,

which confirms previous findings that marine bacteria prefer either a free-living or a surface-

associated lifestyle (Dang and Lovell, 2016; Salta et al., 2013).

Although a growing body of research has analysed marine plastic biofilms, using culture-

independent approaches (Amaral-Zettler et al., 2015; Bryant et al., 2016; De Tender et al., 2017;

De Tender et al., 2015; Debroas et al., 2017; Oberbeckmann et al., 2014; Oberbeckmann et al.,

2016; Zettler et al., 2013), little is known on the specificity of marine biofilms on chemically

distinct (e.g. polyesters, polyolefines) plastic types under comparable conditions. Many studies

conducted so far lack in systematic and statistically robust analysis of distinct plastic types

because they focussed on the comparisons of randomly collected diverse marine plastics of

unknown exposure time and origin (Amaral-Zettler et al., 2015; De Tender et al., 2015;

Oberbeckmann et al., 2014; Zettler et al., 2013) which impede a proper evaluation of substrate

specificity. A few studies were conducted under comparable conditions over short time scales

(Kettner et al., 2017; Oberbeckmann et al., 2018; Oberbeckmann et al., 2016). For example, in

a study located in the North Sea no apparent differences could be perceived between glass and

PET associated communities (Oberbeckmann et al., 2014; Oberbeckmann et al., 2016).

Recently, Oberbeckmann et al. (2018) investigated wood, HDPE and PS associated

communities in a short term experiment (14 days) and found no significant differences

comparing both plastic types. Kettner et al. (2017) investigated fungal communities in the same

short term experiment but also found no differences comparing PE and PS communities

(Kettner et al., 2017).

To date, it is well established that marine biofilms colonizing different plastic types contain

several families in common. These include e.g. Flavobacteriaceae, Erythrobacteraceae,

Hyphomonadaceae and Rhodobacteraceae detected in the North Sea, the coastal Baltic Sea,

multiple locations in the North Atlantic, and freshwater systems (De Tender et al., 2017;

Oberbeckmann et al., 2018; Zettler et al., 2013). Researchers investigating the Plastisphere have

discussed the potential of plastic “specific” organisms/assemblages to be possibly involved in

biological degradation (Amaral-Zettler et al., 2015; Bryant et al., 2016; De Tender et al., 2017;

De Tender et al., 2015; Oberbeckmann et al., 2018; Oberbeckmann et al., 2014; Oberbeckmann

INTRODUCTION

7

et al., 2016; Zettler et al., 2013). For instance, De Tender et al. (2017) identified a core group

of 25 single OTUs, belonging to the phylum Proteobacteria, Bacteroidetes and

Verrucomicrobia on PE. However, it remains unclear whether these “core organisms” are

specific for an environment or whether they are also found on other types of plastics, natural

surfaces or other hard substrates.

Several physicochemical factors, such as hydrophobic surface properties (Oliveira et al., 2001)

and surface rugosity (Bravo et al., 2011; Carson et al., 2013; Characklis, 1991), influence

microbial colonization. The hydrophobic nature of plastics themselves, as opposed to the inert

hydrophilic surfaces (e.g. glass), may result in dissimilarities in community composition, as it

has been already found that microorganisms attach more rapidly to hydrophobic than to

hydrophilic substrates (Bendinger et al., 1993; Fletcher and Loeb, 1979; Pringle and Fletcher,

1983). By comparing three polyolefins (HDPE, LDPE and PP) Artham et al. (2009) showed

that hydrophobicity can favour biofouling. Bravo et al. (2011) observed, in early stage biofilm

formation, fewer taxa on plastic jar surfaces than on Styrofoam pieces and volcanic pumice,

indicating that substrate surface rugosity facilitates initial colonization of floating objects.

Several microorganisms of diverse environments were reported, including bacteria and fungi,

to have a degradative effect on specific plastic types (Crawford and Quinn, 2017; Restrepo-

Flórez et al., 2014). In fact, the biological degradation of plastics is known to be slow and

plastics remain therefore in marine environments for years to centuries (O’Brine and

Thompson, 2010). With all the broad metabolic abilities of microbes, including the ability to

use complex carbon sources the question is raising, why significant differences between diverse

plastics and other inert substrates could not be detected comparing marine biofilms (Kettner et

al., 2017; Oberbeckmann et al., 2018; Oberbeckmann et al., 2016). On the other hand, it needs

to be clarified if organisms repeatedly detected on plastic surfaces reflecting rather a general

biofilm community or a plastic specific one. Moreover, in order to understand the impacts on

plastics as a substrate and potential carbon source in the marine environment, plastic “specific”

microorganisms or assemblages need to be identified.

Most synthetic polymers are rapidly colonized by plethora of organisms. Masó et al. (2003)

detected potential harmful dinoflagellates such as Ostreopsis sp. and Coolia sp., resting cysts

of unidentified dinoflagellates and Alexandrium taylori on floating plastics along the Catalan

coast. Hence, in marine environments plastics can not only serve as an appropriate substrate but

also could function as a vector for the dispersal of alien species including harmful or even

pathogenic species (Barnes, 2002; Masó et al., 2003; Zettler et al., 2013). Also the family of

Vibrionaceae was detected being part of the Plastisphere (Zettler et al., 2013). In this context,

INTRODUCTION

8

Vibrionaceae are of particular interest since this family is known to contain several pathogenic

species. Vibrio spp. are known as animal pathogens invading e.g. coral species (Ben-Haim et

al., 2003), others as human pathogens causing serious infections (Morris, 2003). Especially V.

parahaemolyticus, V. vulnificus and V. cholerae are known as water-related human pathogens

which cause wound infections associated with recreational bathing, septicemia or diarrhea after

ingestion of contaminated foods (Thompson et al., 2004a). For the first time Zettler et al. (2013)

reported the presence of potentially pathogenic Vibrio spp. attached to plastic particles of the

North Atlantic. However, a conclusive identification of Vibrio spp. on the species level was

not provided - (Zettler et al., 2013). Favoured by global warming and the increase of plastics

in the marine environment, it is presumed that potential pathogens could propagate and spread

(Baker-Austin et al., 2016; Baker‐Austin and Oliver, 2018; Zettler et al., 2013).

Exploring the “Plastisphere” – Methodological and experimental approaches

One of the first references from Carpenter and Smith (1972) reported about visually identified

marine organisms, including hydroids and diatoms, associated to plastics surfaces sampled in

the Sargasso Sea. As already mentioned above, in the past years there has been a growing

concern about the Plastisphere and researchers all over the globe started exploring the

Plastisphere at various locations applying a large number of methods with reference to

Plastisphere specific questions. This section focusses on research that has been carried out on

the basis of culture-independent techniques. Comparing the methodological and experimental

approaches of former studies, limitations and research gaps regarding the Plastisphere in natural

marine environments have been identified and linked to the methodological and experimental

approaches used within the frame of this PhD project contributed to fill these gaps.

Various molecular based techniques like cloning, metagenomics, 16S rRNA gene tag

sequencing and denaturing gradient gel electrophoresis (DGGE) have been applied to document

the Plastisphere diversity and variation in natural marine environments (Table 1).

Fingerprint methods like DGGE, used by Oberbeckmann et al. (2014), allow the observation of

the whole prokaryotic community of the Plastisphere by amplification of specific molecular

markers in the environmental DNA. A major advantage of these fingerprinting methods is the

fast and simultaneous analysis of multiple samples, which enables a high comparability

between these samples. However, DGGE alone does not provide taxonomic information, and

the recovering of single bands for direct sequencing is challenging.

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Another approach to gain more detailed information, not only in community structure, but also

in the taxonomic composition of the Plastisphere is the preparation of 16S ribosomal clone

libraries, as used by Dang et al. (2008); Dang and Lovell (2000); Viršek et al. (2017). However,

the preparation of clone libraries requires a strong effort in both, working time and cost, since

every sample can result in hundreds of clones, which are all sequenced separately.

Table 1 List of studies on the marine Plastisphere diversity in natural marine environments based on culture-

independent techniques. Exp. = Experimental; E = exposure experiment, R = random sampling, N.i. = Not

identified/unknown age, Bac. = Bacteria, Prok. = Pokaryotes, Euk. = Eukaryotes, Fun. = Fungi; PVA = Polyvinyl

acetate, PVC = Polyvinyl chloride, PMMA = Polymethyl methacrylate, PE = Polyethylene, PP = Polypropylene,

PA = Polyamide, PS = Polystyrene, PET = Polyethylene terephthalate.

Method Target Study site Habitat Exp. Biofilm

age

Plastic

size

Plastic

type References

Clone libraries

Bac. Salt marsh system,S.C.

USA Marine coastal E

1, 3

days >5mm

PVA,

PVC

Dang and

Lovell (2000)

Bac. Western Pacific Ocean,

CHN Marine coastal E

1, 3

days >5mm

PMMA,

PVC

Dang et al.

(2008)

Bac. North Adriatic Sea, SVN Marine coastal R N.i. <5mm &

>5mm

PE, PP,

PA, PS

Viršek et al.

(2017)

DGGE Bac. North Sea, UK Marine coastal &

offshore E & R

6weeks/

N.i.

<5mm &

>5mm

PET, PS,

PE, PP

Oberbeckmann

et al. (2014)

Amplicon

sequencing

Bac. North Atlantic Ocean Marine offshore R N.i. <5mm PE, PP Zettler et al.

(2013)

Bac. North Sea, BE Marine coastal &

offshore R N.i. >5mm PE, PP

De Tender et al.

(2015)

Bac. North Pacific & North

Atlantic Ocean

Marine coastal &

offshore R N.i. <5mm PE, PP

Amaral-Zettler

et al. (2015)

Bac. Bay of Brest, FRA Marine coastal R N.i. <5mm PE, PP, PS Frère et al.

(2018)

Bac. River Warnow & Baltic

Sea, DE

Marine coastal &

River E 2 weeks <5mm PE, PS

Oberbeckmann

et al. (2018)

Prok.,

Euk. North Sea, UK Marine offshore E 6 weeks >5mm PET

Oberbeckmann

et al. (2016)

Prok.,

Euk.

North Atlantic,

subtropical gyre Marine offshore R N.i.

<5mm &

>5mm

PE, PET,

PS

Debroas et al.

(2017)

Bac.,

Fun. North Sea, BE

Marine coastal &

offshore E 1 year >5mm PE

De Tender et al.

(2017)

Fun. River Warnow & Baltic

Sea, DE

Marine coastal &

River E 2 weeks <5mm PE, PS

Kettner et al.

(2017)

Shotgun

metagenomics

Prok.,

Euk.

North Pacific,

Subtropical Gyre Marine offshore R N.i.

<5mm &

>5mm N.i.

Bryant et al.

(2016)

These limitations were overcome with the introduction of high-throughput sequencing

technologies, like e.g. Roche 454 pyrosequencing, which largely replaced the conservative

Sanger Sequencing. Nowadays, high-throughput sequencing platforms, like “Illumina MiSeq”

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intended for targeted amplicon sequencing and “Illumina HiSeq” for high-throughput

applications as e.g. shotgun metagenomics, allow extensive microbial ecological studies

(Reuter et al., 2015). High-throughput sequencing techniques have the advantage to enable the

processing of a large number of samples simultaneously (>100). Amplicon gene tag sequencing

targets a genomic locus for amplification, e.g. the 16S rRNA gene for prokaryotes or 18S rRNA

for eukaryotes. Therefore, the genomic locus is amplified with specific primers and individual

barcode sequences (tags), which are added to each sample. After sequencing, sequence data can

be differentiated and well-sorted based on the assigned tags. To date MiSeq sequencers can

generate approximately 25 million read clusters with up to 2x300 basepairs (bp) during a single

Illumina run. These large data sets are currently used to explore the vast biodiversity in marine

environments, like e.g. here the Plastisphere (Table 1).

Nevertheless, also amplicon gene tag sequencing confronts limitations. The extractions of

environmental DNA include detritus also in the form of dead organisms and it is therefore

possible that detected highly abundant organisms, are not the most abundant living organisms

in the environment (Taberlet et al., 2012). Microscopic methods like SEM and catalyzed

reporter deposition fluorescence in situ hybridisation (CARD-FISH) had been used previously

to demonstrate the bacterial attachement onto LDPE, and to target specific genera following to

bacterial 16S rRNA gene sequencing analysis (Harrison et al., 2014). Microscopic methods are

also commonly used for the identification of eukaryotic organisms (Salta et al., 2013), which

underlines the need of complementary techniques like e.g. SEM to verify the presence/absence

of e.g. eukaryotic organisms, which are detected by rRNA gene tag sequencing.

Furthermore, due to short read length, a conclusive identification on the species level of the

detected taxa is often not possible. Zettler et al. (2013), using a culture-independent approach,

detected sequences affiliated to Vibrio spp. on marine plastics. Also, De Tender et al. (2015)

reported Vibrionaceae on marine plastics, by using next-generation amplicon sequencing.

Some Vibrio species are known as human pathogens, but within both studies, a conclusive

identification on the species level could not be provided. Thus, this specific Plastisphere related

question if human pathogenic Vibrio spp. are part of the Plastisphere remains unresolved by the

solely use of culture-independent techniques, but can be complemented by the use of rather

conservative culture-dependent approaches.

Considering the impact of geography, season, exposure time and substrate type on the

community composition on marine plastics a proper comparison of different studies is

challenging. Beside that and the additional fact of the use of different methodological

approaches, several further points attract attention, comparing studies addressing the

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Plastisphere with culture-independent techniques (Table 1). The majority of studies focussed

on bacteria. Just three analysed both, prokaryotes and eukaryotes, and only two studies

investigated fungi associated to plastics. The interactions between various groups of organisms

within a biofilm are highly complex. Within this PhD project, prokaryotes and eukaryotes

associated to plastics were investigated to create a more complete picture of the Plastisphere.

Next, polyethylene (PE), followed by polystyrene (PS) and polypropylene (PP) are by far the

most studied substrates. This is not surprising since these plastic types account to the most

produced plastics and consequently represent the most frequently detected plastic particles in

marine environments (Andrady, 2011). Nevertheless, the chemical composition of synthetic

polymers is highly diverse and, as already mentioned above, several plastic types exist which

are also introduced in the oceans. In the frame of this thesis, the Plastisphere communities

associated to nine chemically distinct plastic types were investigated and compared to the inert

control substrate glass. Furthermore, approximately half of the so far conducted studies relies

on randomly collected plastics of unknown exposure time and origin which impede a proper

evaluation of e.g. substrate specificity. Here, a statistically robust analysis of the substrate

specificity of the Plastisphere attached to diverse plastic types was realized. Also, biofilms

investigated were predominantly “young” (weeks), only De Tender et al. (2017) carried out an

annual exposure experiment of PE. Considering that, plastics remain over long time periods in

natural marine environments, incubation over longer timescales allows mimicking more

realistic conditions. Therefore, 15 months old mature biofilms were analysed within this study.

In summary, within this PhD project culture dependent, culture-independent molecular (18S

and 16S rRNA gene tag sequencing) and visual tools (SEM) were applied to investigate the

Plastisphere to provide detailed description of the eukaryotic and prokaryotic marine biofilm

community composition, to further analyse substrate dependent specificities and the

relationships of single bacterial OTUs to various chemically distinct plastic types. To identify

weather potentially pathogenic Vibrio spp. being part of the Plastisphere, a culture-dependent

approach was applied.

OBJECTIVES

12

OBJECTIVES

Since the middle of last century, the global production of plastics was accompanied by an

accumulation of plastic litter in the marine environment. Persistent plastic items are rarely

degraded but become fragmented over time and are dispersed by currents and wind.

Consequently, marine plastic litter can be found in marine waters all over the globe and is

rapidly colonized by marine microorganisms which form dense biofilms on the plastic surface,

the so called Plastisphere (Zettler et al., 2013). However, the number of studies addressing

Plastisphere related questions remains limited. Hence, the ecological impacts of the Plastisphere

and the overall consequences are far from understood. The scope of this thesis was to

comprehensively describe the Plastisphere of a variety of chemically distinct plastics. Hence,

the current thesis provides in-depth insights of the Plastisphere structure gained through culture-

dependent and culture-independent high-resolution methods at community and species levels.

The title and objective of each chapter are listed below:

I. Mature biofilm communities on synthetic polymers in seawater - Specific or general?

Is the Plastisphere a substrate specific or rather a general marine biofilm? How different are

communities attached to diverse plastics and other inert substrates, and which organisms

discriminate the diverse substrates? The substrate specificity of microbial communities on

plastics remains under debate as many studies conducted so far lack systematic and statistically

robust analyses of chemically distinct plastics. Former studies focussed on the comparisons of

randomly collected marine plastics of unknown exposure time and origin which impede a

proper evaluation of substrate specificity. A few studies were conducted over short time scales

in order to address substrate specificity. Considering that plastics remain over long time periods

in natural marine environments, incubation over longer timescales allows mimicking more

realistic conditions. In this study, we examined the specificity of mature (15 months) microbial

communities attached to nine chemically distinct plastic types as well as glass slides as a control

substrate. In this long-term experiment, the different substrates were incubated in a natural

seawater flow-through system allowing colonisation by close to natural biofilm communities.

The composition of both prokaryotic and eukaryotic communities on the different substrate

types was determined by 16S and 18S rRNA gene tag sequencing.

OBJECTIVES

13

II. The Plastisphere – Uncovering tightly attached plastic “specific” microorganisms

Which microorganisms are preferentially able to colonize and interact with plastic surfaces, as

opposed to generalists that colonize also other surfaces? Previous investigations (Chapter I)

indicated that the shared core of the various mature Plastisphere biofilms is rather substrate

unspecific, pointing towards the importance of rather rare species in plastic associated marine

biofilms. Considering that the competition pressure in mature biofilms can be colossal (e.g. for

space or nutrients), uncovering those rare species might be the necessary first step to identify

microbes that are preferentially able to interact with plastics surfaces. Hence, it was

hypothesized that i.) plastic “specific” microorganisms are tightly attached to the polymeric

surface and ii.) that the specificity of plastics biofilms is rather related to members of the rare

biosphere. To test these hypotheses, a three-phase stepwise experiment was conducted. In Phase

1, nine chemically distinct plastic films, and glass for control, were incubated in situ for 21

months in a natural seawater flow through system. In Phase 2, a self-developed high-pressure

water jet treatment technique was used to remove the upper biofilm layers. In Phase 3,

recolonization of a plastic “specific” community was allowed. To verify whether microbes

colonizing different plastics are distinct from each other and from other inert hard substrates,

16S rRNA gene tag sequencing was performed.

III. Dangerous Hitchhikers? Evidence for potentially pathogenic Vibrio spp. on

microplastic particles

Are plastic surfaces a potential spot for the accumulation of pathogens? More specifically, are

potentially human pathogenic Vibrio spp. part of the “Plastisphere”? Previous studies indicated

that potentially pathogenic Vibrio spp. might be present on floating microplastics and therefore

could be transported over long distances in marine environments. Due to short read lengths, a

conclusive identification on the species level was not provided so far. To test the occurrence of

potentially pathogenic Vibrio spp. on marine plastics, plastics and corresponding water samples

of the North and Baltic Sea were analysed with respect to potentially human pathogenic Vibrio

spp. by using cultivation-dependent methods (alkaline peptone water (APW),

CHROMagar™Vibrio), followed by state of the art identification of bacteria on the species

level by MALDI-TOF MS.

OUTLINE

14

OUTLINE

The present thesis consists of a general introduction, three chapters representing one

manuscript each, a general discussion and future perspectives.

Manuscript I (published in Marine Environmental Research)

Kirstein IV, Krohne G, Wichels A and Gerdts G Mature biofilm communities on synthetic

polymers in seawater - Specific or general?

This manuscript describes the specificity of prokaryotic and eukaryotic communities attached

to nine chemically distinct types of plastics and glass as an inert control substrate. The main

outcome is that biofilm communities attached to synthetic polymers are distinct from glass

associated biofilms; apparently a more general marine biofilm core community serves as shared

core among all synthetic polymers rather than a specific synthetic polymer community.

Furthermore, results suggest that synthetic polymer “specialists” might be represented by rather

rare species. Sampling and laboratory investigations were accomplished by Inga Vanessa

Kirstein. 16S rRNA gene tag sequencing was done at LGC Genomics GmbH (Berlin,

Germany). Analysis of sequencing data was done by Inga Vanessa Kirstein. SEM imaging was

carried out by Prof. Dr. Georg Krohne (University Würzburg, Germany). The planning,

statistical analysis, evaluation and writing were carried out by Inga Vanessa Kirstein under the

guidance of Dr. Antje Wichels and Dr. Gunnar Gerdts.

Manuscript II (under review in PLOS ONE)

Kirstein IV, Wichels A, Gullans E, Krohne G and Gerdts G The Plastisphere – Uncovering

tightly attached plastic “specific” microorganisms

This manuscript demonstrates the uncovering of marine plastic “specific”

microbes/assamblages of nine distinct plastic types. It is shown that tightly attached

microorganisms might account rather to the rare biosphere in mature biofilms and furthermore

suggest the presence of plastic “specific” microorganisms/assemblages. The planning,

statistical analysis, evaluation and writing were carried out by Inga Vanessa Kirstein under the

guidance of Dr. Antje Wichels and Dr. Gunnar Gerdts. Laboratory work and DNA extraction

was done by Inga Vanessa Kirstein. 16S rRNA gene tag sequencing was done at LGC Genomics

GmbH (Berlin, Germany). SEM imaging was carried out by Inga Vanessa Kirstein under the

guidance of Prof. Dr. Georg Krohne (University Würzburg, Germany). Inga Vanessa Kirstein

OUTLINE

15

together with the master student Elisabeth Gullans developed the “high pressure treatment”

technique.

Manuscript III (published in Marine Environmental Research)

Kirstein IV, Kirmizi S, Wichels A, Garin-Fernandez A, Erler R, Löder M, and Gerdts G

Dangerous Hitchhikers? Evidence for potentially pathogenic Vibrio spp. on microplastic

particles

This manuscript demonstrates the occurrence of potentially pathogenic Vibrio spp. on floating

microplastics. It is shown that the potentially pathogenic Vibrio parahaemolyticus was part of

the Plastisphere on a number of polyethylene, polypropylene and polystyrene particles from

North and Baltic Sea. Two data sets of two years (2013 and 2014) were combined for this

publication. The master student Sidika Kirmizi collected and analysed samples from 2013, Inga

Vanessa Kirstein collected and analysed samples from 2014. Data evaluation and manuscript

writing was carried out by Inga Kirstein and Sidika Kirmizi under the guidance of Dr. Antje

Wichels and Dr. Gunnar Gerdts. Alexa Garin-Fernandez (2014) and Dr. Rene Erler (2013)

assisted during MALDI TOF analysis. Micro-plastic identification by ATR FTIR was carried

out under the guidance of Dr. Martin Löder.

CHAPTER I

Mature biofilm communities on synthetic polymers in seawater -

Specific or general?

Inga V. Kirsteina*, Antje Wichels a, Georg Krohne b and Gunnar Gerdts a

aAlfred-Wegener-Institute Helmholtz Centre for Polar and Marine Research, Biologische

Anstalt Helgoland, Helgoland, Germany

bUniversity of Würzburg, Biocenter, Imaging Core Facility, Würzburg, Germany

*Corresponding author: Inga Kirstein, Alfred-Wegener-Institute Helmholtz Centre for Polar

and Marine Research, Biologische Anstalt Helgoland, Postbox 180, 27483 Helgoland,

Germany, Tel.: +49 (4725)819-3233; fax: +49 (4725)819-3283; e-mail: inga.kirstein@awi.de

CHAPTER I

18

Abstract

To understand the ecological impacts of the ”Plastisphere”, those microbes need to be identified

that preferentially colonize and interact with synthetic polymer surfaces, as opposed to general

surface colonizers. It was hypothesized that the microbial biofilm composition varies distinctly

between different substrates. A long-term incubation experiment was conducted (15 month)

with nine different synthetic polymer films as substrate as well as glass using a natural seawater

flow-through system. To identify colonizing microorganisms, 16S and 18SrRNA gene tag

sequencing was performed. The microbial biofilms of these diverse artificial surfaces were

visualized via scanning electron microscopy. Biofilm communities attached to synthetic

polymers are distinct from glass associated biofilms; apparently a more general marine biofilm

core community serves as shared core among all synthetic polymers rather than a specific

synthetic polymer community. Nevertheless, characteristic and discriminatory taxa of

significantly different biofilm communities were identified, indicating their specificity to a

given substrate.

CHAPTER I

19

Introduction

During the last decade, there has been a growing concern about the ecological impact of plastics

in the marine environment. The longevity of plastics in the marine environment is a matter for

debate, and estimates range from hundreds to thousands of years depending on the chemical

and physical properties of the synthetic polymer (Barnes et al., 2009). Indeed, plastics remain

much longer in the marine environment than most natural substrates; they represent a new

microbial habitat and due to floating characteristics, they could function as a vector for the

dispersal of pathogenic species (Kirstein et al., 2016; Zettler et al., 2013).

Because synthetic polymers are physically and chemically distinct from naturally occurring

substrates, they offer a new type of substrate to the microbial community. As any surface in the

marine environment, synthetic polymers are rapidly colonized by microorganisms (Harrison et

al., 2014) and subsequently by a myriad of organisms building up complex biofilms (Dobretsov

et al., 2010). Using a culture-independent approach, Zettler et al. (2013) explored for the first

time microbial communities on marine plastic litter. They showed that microbial communities

on marine plastic debris differ consistently from the surrounding seawater communities and

coined these specific biofilms “Plastisphere”. Amaral-Zettler and colleagues (2015) reported

that “Plastisphere” communities of the Atlantic and Pacific Ocean clustered to a greater extend

by geography than by synthetic polymer type. Also, Oberbeckmann et al. (2014) found that the

composition of biofilm communities present on synthetic polymers in marine habitats is driven

by spatial and seasonal effects, but also varies with the plastic substrate type of randomly

sampled plastics. However, in a short-term exposure experiment located in the North Sea they

could not perceive significant differences between glass and PET associated communities

(Oberbeckmann et al., 2014; Oberbeckmann et al., 2016). Despite the increasing research effort

in analysing and understanding the spatial, seasonal, habitational, or substrate parameters

influencing the “Plastisphere”, there is still no consistency concerning the specificity of

microbial communities on different synthetic polymers and other surfaces.

Although some studies have analysed marine plastic biofilms, using a culture-independent

approach (Amaral-Zettler et al., 2015; Bryant et al., 2016; De Tender et al., 2017; De Tender et

al., 2015; Debroas et al., 2017; Oberbeckmann et al., 2014; Oberbeckmann et al., 2016; Zettler

et al., 2013), little is known on the specificity of marine biofilms on chemically distinct (e.g.

polyesters, polyolefines) synthetic polymers under comparable conditions. Recently,

Oberbeckmann et al. (2018) investigated wood, HDPE and PS associated communities in a

short term experiment (14 days) and found no significant differences comparing both polymers.

CHAPTER I

20

Ogonowski et al. (2018) incubated cellulose, glass, PE, PP and PS for two weeks in pre-filtered

seawater and found significant differences between plastic and non‐plastic substrates, but the

specificity of marine biofilms on the respective chemically distinct substrates remains unclear.

Furthermore, in order to understand the ecological impacts of the ”Plastisphere”, those

microbes that preferentially colonize and interact with synthetic polymer surfaces, as opposed

to generalists that colonize other surfaces, need to be identified (Harrison et al., 2014). Recently,

De Tender et al. (2017) identified a core group of 25 single OTUs, belonging to the phylum

Proteobacteria, Bacteriodetes and Verrucomicroboa, on polyethylene (PE), but it remains

unproved whether these “core organisms” are specific for an environment or whether they are

also found on other types of synthetic polymers.

In the present study, it was hypothesized that the composition of marine biofilm communities

varies significantly depending on the substrate type. A long-term experiment was designed in

which nine different synthetic polymers as foils as well as glass slides were incubated in a

natural seawater flow-through system. Previous studies focused essentially on the prokaryotic

or bacterial community composition (Amaral-Zettler et al., 2015; De Tender et al., 2015;

Harrison et al., 2014; Oberbeckmann et al., 2014; Zettler et al., 2013), whereas only a few

studies addressed the complete eukaryotic, or fungal, communities of synthetic polymer

biofilms (Bryant et al., 2016; De Tender et al., 2017; Kettner et al., 2017; Oberbeckmann et al.,

2016). The composition of both prokaryotic and eukaryotic communities on the different

substrate types was determined by 16S and 18S rRNA gene tag sequencing and substrate

specificity assessed. Furthermore, characteristic and discriminatory genera of synthetic polymer

and glass biofilms were identified, and compared those to previously described synthetic

polymer associated biofilms.

CHAPTER I

21

Materials and Method

Experimental design and sample preparation

Synthetic polymers were incubated from August 2013 to November 2014 in the dark (max.

light intensity 0.1033 µmol/m2/s) in a natural seawater flow-through system (Fig S1a) in

conventional slide frames (5 x 5 cm) (Fig S1b) located at the “Biologische Anstalt Helgoland”

(North Sea, Germany, Latitude 54.18286, and Longitude 7.888838) approximately 60 km off

the German coastline. North Sea water was directly pumped through the system (flow rate of

approx. 5800 l/day). The experimental setup simulates sunken plastic, which is largely

protected from photochemical degradation, enabling a well-defined interaction between the

different synthetic polymers and the microbial community. The different exposed synthetic

polymers represent the most frequent polymer types in the marine environment and were

provided by various suppliers: high-density polyethylene (HDPE) (ORBITA-FILM GmbH),

low-density polyethylene (LDPE) (ORBITA-FILM GmbH), polypropylene (PP) (ORBITA-

FILM GmbH), polystyrene (PS) (Ergo.fol norflex GmbH), polyethylene- terephthalate (PET)

(Mitsubishi Polyester Film), polylactic acid (PLA) (Folienwerk Wolfen GmbH), styrene-

acrylonitryle (SAN) (Ergo.fol norflex GmbH), polyurethane prepolymer (PESTUR) (Bayer),

polyvinyl chloride (PVC) (Leitz) (Table S1). As control substrate, glass slides were incubated

in parallel. Glass is inert opposed to most natural surfaces and therefore enables the

development of a general marine biofilm community. Using foils allowed us to 1. Separately

incubate each piece without touching each other, so that even biofilms can develop. 2. It enables

us of taking subsamples of the same piece of foil/biofilm for different approaches (e.g. future

FISH studies). After 15 months of incubation, five replicates of each synthetic polymer with

the associated microbial biofilm were taken (Fig S1c). Environmental data including salinity

(S), water temperature (T) and chlorophyll a (Chl a) were recorded in parallel as part of the

Helgoland Roads time series (Wiltshire et al., 2008) (Fig S1d). Each foil was cut into strips and

glass was broken into fragments of ̴ 1 cm2 using ethanol sterilised forceps, scalpels and scissors.

To remove the unspecific loosely attached part of the biofilm, each polymer strip was washed

in 1 mL 0.2 µm filtered and autoclaved sterile seawater three times for 30 s (vortex) with

transferring the strip after each washing step in a new 1.5 mL tube. Synthetic polymer strips

and glass fragments were stored at -20°C for further analysis.

CHAPTER I

22

SEM

Strips or fragments of subsamples of two replicates (out of five) of each synthetic polymer and

glass were fixed at 4°C in sterile sea water containing 2.5% glutaraldehyde and 50 mM sodium

cacodylate (pH 7.2) and stored at 4°C (4-10 days) until processing. Before, one subsample of

each replicate (n = 2) was washed to remove the unspecific loosely attached part of the biofilm

as described above; the other one remained untreated to visualize the whole community.

Samples were stepwise dehydrated in ethanol, critical point dried (BAL-TEC CPD 030;

Balzers, Liechtenstein) and sputter coated (BAL-TEC SCD 005; Balzers, Liechtenstein) with

gold-palladium before SEM analysis (JEOL JSM-7500F; Freising, Germany).

DNA extraction

DNA of microbial biofilms was extracted using a modified protocol from Sapp et al. (2006).

Each replicate of each substrate (n = 5) was individually transferred into 2 mL screw cap

reaction tubes containing a mixture of 100 µm Zircona/-Silica beads, 700 µL Sodium Chloride

–Tris – EDTA (STE) - Buffer was added before mechanically pulped (FastPrep® FP 120,

ThermoSavant,Qbiogene, United States) for 40 seconds on level 4.0. DNA concentrations were

quantified with a PicoGreen assay (Invitrogen, Waltham, MA) using a Tecan Infinite M200

NanoQuant microplate reader (Tecan, Switzerland).

16S & 18S rRNA gene tag sequencing of biofilm communities

16S and 18S rRNA gene tag sequencing was performed at LGC Genomics GmbH (Berlin,

Germany). Community DNA samples were sent to LGC for generation of 16S V3 / V4 and 18S

V4 rRNA amplicon libraries for Illumina sequencing. Community DNA was amplified using

amplification primers targeting the V3 / V4 region of the 16S rRNA gene using 341F (5’-

CCTACGGGNGGCWGCAG-3’) and 785R (5’-GACTACHVGGGTATCTAATCC-3’)

(Klindworth et al., 2013). Eukaryotic community DNA was amplified using amplification

primers targeting the V4 region of the 18S rRNA gene using Eu565F (5`-

CCAGCASCYGCGGTAATTCC-3`) and Eu981R (5`-ACTTTCGTTCTTGATYRATGA-3`)

(Piredda et al., 2017). The amplicons were paired-end sequenced 2 x 300 bp on an Illumina

MiSeq platform. The paired-end reads were merged using BBMerge 34.48 software

(http://bbmap.sourceforge.net/) and processed through the SILVAngs pipeline (Quast et al.,

2013). All sequences were de-replicated at 100% identity and further clustered with 98%

sequence identity to each other. Representative sequences from operational taxonomic unit

CHAPTER I

23

clusters (OTUs) were classified up to genus level against the SILVA v123 database using

BLAST as first described by Ionescu et al. (2012). Sequences having an average BLAST

alignment coverage and alignment identity of less than 93% were considered as unclassified

and assigned to the virtual taxonomical group “No Relative" (Quast et al., 2013). Finally,

3,517,422 (99.37%) classified sequences were obtained for bacteria and archaea, and 5,163,443

(86.49%) classified sequences were obtained for eukaryotes. For following downstream

analyses, classifications on the genus-level were used to generate the final abundance matrixes.

All classifications contained the sum of all sequences represented by OTUs with the equal

taxonomic path. Sequence data was deposited in the European Nucleotide Archive (Toribio et

al., 2017) under the accession number PRJEB22051, using the data brokerage service of the

German Federation for Biological Data (Diepenbroek et al., 2014), in compliance with the

Minimal Information about any (X) Sequence (MIxS) standard (Yilmaz et al., 2011).

Statistics and Downstream Data Analysis

All multivariate analyses were carried out with the Primer 6 software package plus the add-on

package PERMANOVA+ (PRIMER-E Ltd, UK). The entire prokaryotic and eukaryotic

communities were analysed separately. The virtual taxonomical group “No Relative” was

removed from the analysis. Subsequently, counts per classification were normalized by

calculating their relative abundances to the total number of SSU rRNA gene reads per sample.

For prokaryotes OTUs with a minimal mean relative abundance of 0.1% (n=5) in at least one

substrate type were considered for further analysis. Beta diversity analysis and related

hypothesis testing of the complete eukaryotic community was carried out on the basis of

presence-absence metrics. OTUs with a total abundance of 1 read were excluded from

downstream analyses. To visualize patterns in community composition, principal coordinates

analysis (PCO) was performed using Hellinger distance (D17; (Legendre and Legendre, 1998))

or Jaccard index for eukaryotes. Binary (presence/absence) or square root transformed relative

abundances of sequence read numbers were used for distance matrix calculation. To test for

statistically significant variance among the biofilm communities attached to the different

substrates, PERMANOVA with fixed factors and 9999 permutations at a significance level of

p<0.05 was performed. Tests of significant differences in the within-group dispersion among

the substrate groups were accomplished by performing tests of homogeneity of dispersions

(PERMDISP) using 9999 permutations at a significance level of p<0.05. Similarity percentage

analysis (SIMPER) allowed us to calculate the total similarity within and dissimilarity between

CHAPTER I

24

the different groups of substrates, and to determine characteristic and discriminatory OTUs.

SIMPER analysis was performed using Bray Curtis similarity (S17) by the use of binary

(presence/absence) or fourth root transformed relative abundances (Clarke, 1993).

CHAPTER I

25

Results

Prokaryotic and eukaryotic biofilm composition of nine synthetic polymers & glass

After 15 month of exposition in the natural sea water flow through system, a dense microbial

biofilm colonized all provided substrates (Fig S1 (c)). SEM was used to examine the biofilm in

addition to DNA based techniques. The synthetic polymer and glass associated biofilm

communities analysed by 16S and 18S rRNA gene tag sequencing contained in total 1479

prokaryotic and 692 eukaryotic different operational taxonomic units (OTUs). SEM confirmed

a highly diverse biofilm community growing on all substrate types (Fig 1A(a-k)) consisting of

prokaryotic and eukaryotic microorganisms of different morphologies. Different flagellates

were observed being part of the biofilm community. Exemplarily Fig 1A (i) shows a flagellate

cell having a substantial covering or pellicle. Mature loricae of Acanthoeca spectabilis

(Leadbeater et al., 2008) belonging to the detected class Acanthoecida (Fig 1C) were often

observed by SEM being part of the biofilm community (Fig 1A (d)). Fig 1A (k) shows a striking

specimen what appear to be a surface arrangement of scales and a peripheral array of long

flexuous spines with obconical meshwork bases. The most closely similar specimens are

attributable to the genus Luffisphaera spp. (VØRS, 1993).

Prokaryotic biofilm communities of all substrates were dominated (mean relative abundance

>1% in at least one substrate type) by OTUs assigned to 20 classes (Fig 1B). All biofilms

consisted of a high proportion of Proteobacteria (42–47%) with most abundant classes of

Alpha- (11–15%), Delta- (11–13%) and Gammaproteobacteria (13–16%). Beside the high

proportion of Proteobacteria the taxonomic classes of Nitrospira (7–12%), Planctomycetacia

(5–8%), Caldilineae (4–7%), Acidimicrobiia (4–7%), Sphingobacteria (3–7%) and an

unclassified OTU of Planktomycetes OM190 (2–4%) were more abundant in all biofilm

communities (Fig 1B). Interestingly, the biofilms on glass displayed clear differences in

community composition compared to all synthetic polymers. For example, an unclassified

Latescibacteria and the unclassified Proteobacteria AEGEAN-245 were more abundant on

glass (Fig 1B).

CHAPTER I

26

Fig 1 Biofilm community composition on different synthetic polymers and glass. A: Scanning electron

microscopy images of the biofilm community attached to synthetic polymers and glass. Scale bar = 1 µm. (a)

Region of the highly diverse marine biofilm observed on PVC. (b) Spirochete embedded in EPS (HDPE). (c)

Organized rod-shaped bacteria embedded in EPS (glass). (d) Acanthoeca spectabilis showing left-handed helical

arrangement of costae in stalk and vase (PESTUR). (e) Box-shaped bacteria (LDPE). (f) Stalked Salpingoeca sp.

(PS). (g) Belike cyanobacteria (PP). (h) Region of a biofilm with rod- and spiral shaped bacteria (PET) (i)

Flagellate (PET). (j) Belike fungi spores and hyphae (HDPE). (k) Luffisphaera sp. (PESTUR). Images a), c), e)

and i) show biofilms without, images b), d), f), g), h), j) and k) show biofilms after excessive washing. B:

Abundance profiles of prokaryotic and C: eukaryotic classes on different synthetic polymers and glass. OTUs with

a mean relative abundance of at least 0.1% in one substrate type (n = 5) were analysed. Displayed are prokaryotic

taxonomic classes with abundances of > 0.1% and eukaryotic classes of > 1% in at least one substrate type. The

group `others` was made up of classes with abundances < 1%. A * indicates the term `unclassified class`. Numbers

indicate highly abundant prokaryotic (1-9) and eukaryotic (10-14) classes. Arrows indicate differences in glass

biofilms (B) and the most abundant class of fungi (C).

CHAPTER I

27

In contrast to the relative homogenous prokaryotic community composition among all synthetic

polymers, the eukaryotic biofilm communities were highly heterogeneous (Fig 1C).

Intramacronucleata, belonging to the SAR clade, was one of the most abundant eukaryotic

classes (4–25%) within the biofilm communities of both synthetic polymers and glass. The

diverse class of crustaceans Maxillopoda had a mean relative abundance between 0.8–22%. An

unclassified OTU belonging to Gastrotricha made up a portion of between 0.2 up to 24% of

the eukaryotic biofilm community. Demospongiae, a highly diverse class of the phylum

Porifera, appeared with abundances in between 3–21% and Chromadorea, belonging to the

phylum Nematoda, appeared with abundances between 0.8–23% within the eukaryotic biofilm

communities. Interestingly, animals like Maxillopoda or Nematoda were not observed by SEM

as opposed to regularly seen Diatomea and Sponges (data not shown). Considering the

proportion of Fungi within the eukaryotic community, Chytridiomycetes represented the highest

abundances among biofilms of all substrates with 3% on PET and 1.2% on glass (Fig 1c).

Substrate specificity of the prokaryotic biofilm communities

To determine whether microbial communities colonizing the different substrates are distinct

from each other, the community structure on the genus level of biofilms attached to nine

different synthetic polymers and those colonizing glass was compared. Samples of synthetic

polymers and the control substrate glass clustered clearly in bisection (Fig 2a). The 16S rRNA

gene sequence comparisons showed significant differences between the glass associated

biofilm communities and those associated with synthetic polymers (p<0.05; pairwise

PERMANOVA, Table S3). A separate test of dispersion using PERMDISP revealed that the

differences among the specific synthetic polymers to glass were at least partially driven by

different within-system heterogeneities in five cases (Table S4). Significant differences were

also observed in 15 out of 36 possible synthetic polymer-pair combinations, between different

polymer-colonizing communities (Table S3). PLA communities were significantly different

from seven other synthetic polymer communities, followed by PESTUR and PVC communities

that significantly differed from five and four further synthetic polymer communities. HDPE,

PS, PET and SAN communities differed significantly from three, PP and LDPE communities

differed significantly from one other synthetic polymer communities (Table S3).

CHAPTER I

28

Fig 2 Principle Coordinate Ordination (PCO) relating variation in microbial community composition

between different synthetic polymers and glass biofilm communities. PCOs representing similarity of biofilm

communities based on relative abundances (prokaryotes) and presence/absence (eukaryotes) of OTUs across

samples. Displayed are comparisons of (a) prokaryotic and (b) eukaryotic communities of synthetic polymer

attached and glass attached 15 month old biofilm communities.

Prokaryotic biofilm communities associated with different synthetic polymers differed between

3.9–5.5% from each other, and between 5.5–7.6% from the control substrate glass (Table S5).

Considering the relative abundances of single OTUs, nine OTUs appeared with relative

abundances >3% of the total community composition including e.g. Nitrospira (OTU 576), the

unclassified Deltaproteobacteria SH765B-TzT-29 (OTU 1123) and an uncultured unclassified

Caldilineacea (OTU 359) (Fig 3).

Five OTUs were predominantly discriminating the biofilm on glass from synthetic polymer

biofilm communities: the unclassified genus Acidobacteria AT-s3-28 (OTU 13), Halophagae

Sva0725 of the subgroup 10 (OTU 37), the genus Gilvibacter (OTU 231), Leptobacterium

(OTU 240), and the Candidatus Entotheonella (OTU 1058) (Fig 3). The unclassified

Halophagae Sva0725 and Gilvibacter were more characteristic for synthetic polymer

communities (Table S7), with relative abundances of >1%, respectively. The unclassified genus

Acidobacteria AT-s3-28 contributed to the total dissimilarity between glass and all synthetic

polymers, and was always more characteristic for glass biofilm communities, with relative

abundances <1% (Fig 3, Table S7). The Candidatus Entotheonella, with relative abundances

of >3%, contributed more to total similarity of glass biofilm communities (Fig 3, Table S7).

Beside the detected differences of glass and synthetic polymer communities, PLA associated

communities showed significant differences to seven synthetic polymer community groups

(Table S3). The largest dissimilarities between PLA and all other substrates was caused by an

OTU belonging to the genus Leptobacterium (OTU 240), with overall relative abundances <1%

(Fig 3). While the genus Leptobacterium was characteristic for PLA communities, the

CHAPTER I

29

unclassified Acidobacteria AT-s3-28 also contributed to the total dissimilarities of PLA by

being characteristic of glass communities (Table S7). Further, five OTUs contributed explicitly

to the total dissimilarities between PLA and the other synthetic polymer associated biofilm

communities. Genera contributing explicitly to the total dissimilarities between PLA and the

other synthetic polymers were an unclassified Holophagae CA002 of the Subgroup 10 (OTU

35), Ardenticatenales (OTU 355), an unclassified Oligosphaeria (565), Nitrospira (OTU 576)

and Nitrospina (OTU 1059). The unclassified Holophagae CA002 was most characteristic for

PLA (Table S7). The unclassified Oligosphaeria contributed least to the total similarity of PLA.

Nitrospira clearly discriminated PLA from PESTUR communities. The unclassified genus

Ardenticatenales contributed highly to the total dissimilarities, explained by relative

abundances of 0.9% for PLA and 1.1% for PVC communities, compared to relatively low

contributions of 0.2% for HDPE communities (Fig 3).

Fig 3 Most abundant and discriminative prokaryotic OTUs of the nine different synthetic polymers and

glass (n=5). OTUs with a mean relative abundance of at least 0.1% (n=5) in at least one substrate type were

analysed. Displayed are OTUs with a mean relative abundance of at least 3% or jointly contributing, with a

minimum of 2%, to the total dissimilarity between different statistically significant (PERMANOVA p<0.05) glass

and synthetic polymer groups. Groups showing both, PERMANOVA and PERMDISP significant p values were

rejected. The amount of contribution is indicated by the colour of cells, darker colours represent higher

contributions. Bold lines indicate OTUs contributing to the same phylum. A * indicates the term “unclassified”.

With exception of Nitrospira (OTU 576) and Candidatus Entotheonella (OTU 1058), the OTUs

contributing most to the total dissimilarity between substrates were not the most abundant ones.

Instead, less abundant OTUs like the unclassified Acidobacteria AT-s3-28, being more

characteristic for glass communities, contributed strongly to the total dissimilarity between

glass and synthetic polymer biofilm communities (Fig 3, Fig S3, Table S7).

CHAPTER I

30

Substrate specificity of the eukaryotic biofilm communities

Considering the possible bias due to preferential amplification of primers resulting variation in

copy numbers which might affect the relative abundance estimates of all species in the sample

by over-representation of specific taxa, Beta diversity and related hypothesis testing of the

general eukaryotic community was carried out on basis of presence-absence metrics. In contrast

to the prokaryotic communities, for eukaryotes no clear clustering between the different

synthetic polymers or the control substrate glass was observed (Fig 2b). Eukaryotic biofilm

communities differed between 44.1–56.3% from each other (Table S6). Furthermore, there was

a significant difference between the HDPE-, LDPE-, PESTUR-, PP-, PS-, PET-, and PLA to

glass associated eukaryotic communities. However, a separate test of dispersion using

PERMDISP revealed that these differences among substrates were most likely driven by

different within-system heterogeneities (Table S4). Significant differences, devoid of within-

system heterogeneities, were also observed in synthetic polymer-pair combinations. Eukaryotic

communities colonizing PLA significantly differed to PP-, PVC and PESTUR associated

communities (p<0.05; pairwise PERMANOVA, Table S3). Furthermore, communities

colonizing PS significantly differed to PESTUR. LDPE communities differed significantly to

PET (p<0.05; pairwise PERMANOVA, Table S3).

Explicitly discriminant of the PLA communities as compared to communities on PP-, PVC and

PESTUR was an OTU belonging to the genus Hatena (Cryptophyceae, OTU 71) and Gyromitus

(Rhizaria, OUT 499) both absent on PLA. An OTU belonging to the class of Asteroidea

(Metazoa, OUT 144) contributed to the total dissimilarities between PLA, PVC and PS.

Another genus discriminating PLA from PP communities was the dinoflagellate Prorocentrum

(OTU 442). The overall variation between synthetic polymer eukaryotic communities was in

total not driven by fungal OTUs (Fig S4).

Biofilm vs. free living communities

To demonstrate the distinctness of microbial biofilm communities, commonly found marine

prokaryotic microbial seawater communities of weekly collected samples of a one year time

series at Helgoland Roads (March 2012 – February 2013, (Lucas et al., 2015) were compared

to the pooled microbial biofilm communities (Fig 4, Table S9) on the class level.

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31

Fig 4 Venn diagram showing prokaryotic taxonomic class overlap for pooled biofilm samples (n=50, incubated

in Helgoland seawater from August 2013 – November 2014, OTUs with a mean relative abundance of at least

0.1% (n=5) in at least one substrate type were analysed.) associated to nine different synthetic polymers and glass,

and seawater samples (n=42, collected weekly from March 2012 – February 2013 OTUs with a mean relative

abundance of at least 0.1% (n=42)) at Helgoland Roads (Lucas et al., 2015); n = number of OTUs per group.

Numbers inside the circles represent the number of shared or unique classes for the given environment. Images

were generated using Venny 2.1 (http://bioinfogp.cnb.csic.es/tools/venny/index.html).

The percentage of shared classes across the two habitats (Fig 4, Table S9) reflects the

distinctness of seawater and biofilm communities. More classes were detected in biofilm

samples than in seawater samples, the former were partly consisting of single OTUs that could

not be assigned to a taxonomic class (Table S9). Seven classes (14%) were exclusively detected

within seawater communities including i.e. Actinobacteria, Cyanobacteria, Deferribacteres

and Thermoplasmata (Table S9). Further 26 classes (52%) were exclusively detected within

biofilm communities, including i.e. Acidobacteria, Ardenticatenia, Caldilineae, Caldilineae,

Deinococci, Holophagae, Melainabacteria, Nitrospira, Oligosphaeria and Phycisphaerae

(Table S9). Overall, 34% of the classes were common to biofilm and seawater communities and

included members of Acidimicrobiia, Alphaproteobacteria, Betaproteobacteria, Cytophagia,

Deltaproteobacteria, Epsilonproteobacteria, Flavobacteria, Gammaproteobacteria and

Gemmatimonadetes.

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32

Discussion

The substrate specificity of microbial communities on synthetic polymer remains under debate

as many studies conducted so far lack in systematic and statistically robust analysis of distinct

synthetic polymers. Former studies focussed on the comparisons of randomly collected diverse

marine synthetic polymers of unknown exposure time and origin (Amaral-Zettler et al., 2015;

De Tender et al., 2015; Oberbeckmann et al., 2014; Zettler et al., 2013) which impede a proper

evaluation of substrate specificity. A few studies were conducted over short time scales (Kettner

et al., 2017; Oberbeckmann et al., 2018; Oberbeckmann et al., 2016), considering that synthetic

polymers remain over long time periods in natural marine environments, incubation over longer

timescales allows mimicking more realistic conditions. Here, a thorough analysis of substrate

specificity of prokaryotic and eukaryotic North Sea biofilms with regard to the taxonomic

structure and composition of 15 month old microbial biofilms as compared on different

synthetic polymer types in a natural seawater flow-through system was carried out.

Comparison of biofilm and seawater communities showed that, despite possessing classes in

common, both communities are generally distinct. This finding supports several previous

studies (Amaral-Zettler et al., 2015; Bryant et al., 2016; De Tender et al., 2017; De Tender et

al., 2015; Oberbeckmann et al., 2014; Oberbeckmann et al., 2016; Zettler et al., 2013) pointing

toward a consensus that free-living seawater communities are different from synthetic polymer

attached ones. A possible explanation might be the much higher cell density in biofilms as

compared to seawater; hence higher cell density may support the development of matrix-

stabilized, synergistic micro-consortia.

Synthetic polymer associated prokaryotic biofilm communities were different from glass

biofilm communities. Furthermore, significant differences between the prokaryotic and

eukaryotic community composition of different synthetic polymers communities were found.

In contrast to clearly distinct prokaryotic seawater communities, differences between substrates

were generally low (3.9–7.6%). A few notable OTUs uniquely discriminated the biofilm

communities across the diverse substrates, suggesting that physicochemical properties of the

substrate shape synthetic polymer communities. Complex biofilms include a diversity of

organisms with different metabolic capacities and physiologies which generates on the one

hand competition but also provides on the other hand opportunities for cooperation (Flemming

et al., 2016).

In contrast to the homogenous prokaryotic communities analysed here, substantial

heterogeneity between eukaryotic communities on the diverse substrates was observed.

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33

Statistical analyses of eukaryotic communities revealed significant differences between diverse

substrates, surprisingly mainly due to OTUs predominantly assigned to mobile organisms e.g.

Dinoflagellata or starfish (Asteroida). Chesson and Kuang (2008) assumed that competition

dynamics at lower trophic levels (bacteria and microflagellates) might have consequences for

protists` dynamics. Thereby, the polymer characteristics may select for microorganisms and

they, in turn, might attract different grazers. However, this mobile organism may not be specific

for a substrate and may not be found as discriminating organisms in other studies. For

clarification, the polymer strips were washed excessively in that loosely attached biofilm parts

were removed. This suggests that reads assigned to mobile organisms could also originate from

detritus or eggs strongly embedded in the EPS, this is also an explanation why, beside others,

starfish have been identified only by molecular tools but not by SEM. Furthermore, based on

the general heterogeneity of eukaryotic communities it can be assumed that this observation

may be coincidental.

Analysing the eukaryotic community composition, the class of Chytridiomycetes

(Chytridiomycota) was found with highest abundances across all detected fungal classes.

Recently, Kettner et al. (2017) investigated fungal communities attached to PE and PS from the

River Warnow to the Baltic Sea but found no significant differences comparing both substrates

communities. Interestingly, in the study of Kettner et al. (2017), the majority of fungal 18S

rRNA reads were assigned to Chytridiomycota, which is consistent with our findings. Since

fungi are of particular interest in their role as potential plastic degraders in the environment

(Grossart and Rojas-Jimenez, 2016; Krueger et al., 2015), the repetitive detection of highest

abundances of Chytridiomycota associated to marine plastics in both studies suggests that

further investigations on their role in plastic biofilms are required.

In general, differences in the biofilm community composition are related to different factors,

for example the substratum physicochemical properties e.g. hydrophobicity, roughness,

vulnerability to weather but also surface chemodynamics like surface conditioning or nutrient

enrichment (Dang and Lovell, 2016). Particularly primary colonizers, sensing the synthetic

polymer surface, impact community formation, dynamics, and function (Dang et al., 2008). In

respect of PLA, which is known to be biodegradable when composted, the degradation

mechanism start with chemical hydrolysis in the presence of water at elevated temperatures

(60°C and above), followed by biological degradation (Shah et al., 2008). Since North Sea

water temperatures were never above 18°C during the 15 month of our experiment, biotic

degradation is unlikely.

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34

Beside physicochemical surface properties, it has been shown that the composition of biofilm

communities associated to synthetic polymers differed distinctly with respect to different ocean

basins (Amaral-Zettler et al., 2015) and underlies both seasonal and spatial effects e.g. in North

Sea waters (Oberbeckmann et al., 2014). Biofilms in this study were sampled at one time point,

thus seasonal and temporal changes in the taxonomic composition were not investigated.

However, these biofilms were exposed to seasonal variation of several environmental factors

in the North Sea such as temperature or nutrient variation within the seawater flow-through

system. To delineate the effects of seasonal variation on the community composition biofilms

should be monitored at close intervals best over more than one seasonal cycle. The incubation

conditions applied in this setting of a natural seawater flow-through system with e.g. less shear

forces and lack of light, in contrast to incubation in the open sea, may have influenced the

establishment of a synthetic polymer specific community. It is known that biofilm community

composition is strongly driven by the factor environment (Salta et al., 2013). Recently, in a

long-term exposure experiment of PE in two different environments, harbour and offshore, De

Tender and colleagues (2017) demonstrated a shift toward more secondary colonizers of PE

biofilms at later stages, interestingly, only in the harbour environment, an environment which

is less exposed to shear and current forces. To the best of our knowledge, the only other study

which compared PET with glass- communities, after exposure in the open sea (i.e. high shear

stress), found no distinct communities (Oberbeckmann et al., 2016). In contrast, in the present

study clear differences were observed between prokaryotic communities on synthetic polymers

as compared to glass after exposure in a seawater flow-through system with low shear stress.

However, the time of exposure in our experiment was much longer than in the study of

Oberbeckmann et al. (2016), thus the latter synthetic surfaces (i.e. glass vs. PET bottles) were

colonized by a relatively “young” biofilm community after exposure of 5 to 6 weeks as opposed

to the 15 month “old” biofilm, investigated in the present study. Hence it can be presumed that

early colonizers might be more generalists than specialists and specific biofilm communities

evolve over a longer period of time or/and in semi enclosed environments.

OTUs with a mean relative abundance of at least >0.1% in one substrate type were analysed,

and found that along these, even if sometimes rare (<0.1%) all prokaryotic OTUs were detected

on synthetic polymers and glass. Hence, the dissimilarities in the prokaryotic community

composition observed as a function of the synthetic polymers investigated resulted from

variable relative abundance profiles of dominant OTUs. Recently, De Tender et al. (2017)

identified a core group of 25 single OTUs based on their abundance profiles on PE in the

Belgian North Sea. Comparison with our data revealed that four of the reported genera were

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35

also present, with relative abundances >0.1%, in the 15 month old biofilm communities

analysed in the present study, belonging to Anderseniella, an uncultured Rhodobacteraceae,

Sulfurovum, and the unclassified OTU belonging to Proteobacteria of the marine benthic group

JTB255 (Fig S3, Table S7). It remained unclear whether these indicator organisms are specific

for the environment or whether they are commonly found more generally on different types of

hard substrates. First, these organisms seem to be rather unspecific for the tested environment

and may be therefore useful as indicator organisms for biofilm development in several parts of

the North Sea. Second, with the exception of Sulfurovum, the above-mentioned genera were

present on all substrate types without notably discriminating the different biofilm communities,

suggesting that these organisms are common members of North Sea biofilms. Third, the overall

dissimilarities between the analysed prokaryotic communities were generally low, which

indicates that the shared core of the various biofilms is rather substrate unspecific. Fourth, the

strongest contribution to the total dissimilarity between the diverse substrates was often given

by less abundant OTUs (<1%). Consequently, identification of a core group of indicator

organisms of polymer specific biofilms based on the dominant OTUs is limited, because it

illustrates a more general marine biofilm core community rather than a synthetic polymer

specific one.

Significant differences between various substrates for prokaryotes and eukaryotes were

detected but also substantial heterogeneity between eukaryotic biofilms. The present study, as

well as other research about the composition and function of eukaryotes in marine biofilms,

suffers from a gap in current taxonomic reference databases. Only 86.49% of the sequences

obtained for eukaryotes were classified (coverage and alignment identity of min. 93%). This

illustrates the current need to combine molecular based techniques and visual tools like SEM.

Luffisphaera (VØRS, 1993) probably represents one of those taxa which probably counted

among the unclassified sequences (13.5%). Even though the genus Luffisphaera has been

described, and comprises several species, tag sequence data is not available yet and the

phylogeny of this protist is still unresolved. Furthermore, visual inspection by SEM enables to

identify species, e.g. Acanthoeca spectabilis, verify the presence/absence of mobile organisms,

e.g. starfish (Asteroida), which were detected only by rRNA gene tag sequencing. Concerning

the repetitive detection of highest abundances of Chytridiomycota associated to marine plastics,

the use of fungi specific primers in upcoming studies needs to be considered, to gain detailed

insights in their taxonomy. To date due to short read lengths, a conclusive identification of

discriminative biofilm members on the species level is not reliable. However, synthetic polymer

“specialists” might be represented by rather rare species, thus they would have been missed

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36

them since the sequencing approach was not deep enough for analyses of the rare biosphere.

Since phylogenetic assignment based on rRNA gene tag sequencing is not linked to specific

functions or metabolic activity, specific roles of the discriminating members related to the

synthetic polymers remain theoretical. To gain insights into the function and activity of

microbial biofilm communities, including the rare biosphere, attached to synthetic polymers

further experiments including “omics” need to be conducted. To identify those specialised

microbes that are preferentially able to colonize and interact with synthetic polymer surfaces,

those organisms need to be selected and enriched from the shared core biofilm community and

to test their potential degradation ability.

Conclusion

Our study represents a systematic and statistically robust analysis of 15 month old biofilms

associated to distinct synthetic polymers, and therefore enrich our knowledge on the substrate

specificity of the “Plastisphere”. First and foremost, it has been proofed that mature biofilms

attached to synthetic polymers are significantly different from glass biofilms. Although

differences of prokaryotic communities between synthetic polymers were generally low (3.9–

5.5%), significant differences between biofilms on diverse polymers were observed.

Furthermore, it was shown that a more general prokaryotic marine biofilm core community

serves as shared core among all synthetic polymers rather than a specific synthetic polymer

community. However, the general heterogeneity of eukaryotic communities was much higher,

concluding that observations of significant differences may be coincidental. These findings

indicate that the term “Plastisphere” is valid for mature prokaryotic but may not be for

eukaryotic biofilm communities.

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37

Acknowledgments

We would like to thank Dr. Barry Leadbeater, University of Birmingham, for helping us with

the identification of eukaryotic organisms visualized by SEM. We thank also Maike

Timmermann for her assistance in the laboratory. Furthermore, we would like to thank Dr.

Marlies Reich, University of Bremen, for her support. We thank Dr. Cédric Meunier for fruitful

discussions. We are grateful for kind provision of environmental data by Prof. Dr. Karen

Wiltshire. This work was founded by the Alfred Wegener Institute, Helmholtz Centre for Polar

and Marine Research and supported by the German Federal Ministry of Education and Research

(Project BASEMAN - Defining the baselines and standards for microplastics analyses in

European waters; BMBF grant 03F0734A).

CHAPTER II

The Plastisphere –

Uncovering tightly attached plastic “specific” microorganisms

Inga V. Kirsteina*, Antje Wichelsa, Elisabeth Gullansa, Georg Krohneb, Gunnar Gerdtsa

aAlfred-Wegener-Institute Helmholtz Centre for Polar and Marine Research, Biologische

Anstalt Helgoland, Helgoland, Germany

bUniversity of Würzburg, Biocenter, Imaging Core Facility, Würzburg, Germany

*Corresponding author: Inga Kirstein, Alfred-Wegener-Institute Helmholtz Centre for Polar

and Marine Research, Biologische Anstalt Helgoland, Postbox 180, 27483 Helgoland,

Germany, Tel.: +49 (4725)819-3233; fax: +49 (4725)819-3283; e-mail: inga.kirstein@awi.de

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Abstract

In order to understand the degradation potential of plastics in the marine environment,

microorganisms that preferentially colonize and interact with plastic surfaces, as opposed to

generalists potentially colonising everything, need to be identified. Accordingly, it was

hypothesized that i.) plastic “specific” microorganisms are closely attached to the polymeric

surface and ii.) that specificity of plastics biofilms are rather related to members of the rare

biosphere. To answer these hypotheses, a three phased experiment to stepwise uncover closely

attached microbes was conducted. In Phase 1, nine chemically distinct plastic films and glass

were incubated in situ for 21 months in a seawater flow through system. In Phase 2, a high-

pressure water jet treatment technique was used to remove the upper biofilm layers to further,

in Phase 3, enrich a plastic “specific” community. To proof whether microbes colonizing

different plastics are distinct from each other and from other inert hard substrates, the bacterial

communities of these different substrates were analysed using 16S rRNA gene tag sequencing.

Our findings indicate that tightly attached microorganisms account to the rare biosphere and

suggest the presence of plastic “specific” microorganisms/assemblages which could benefit

from the given plastic properties or at least grow under limited carbon resources.

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41

Introduction

Since the middle of last century the increase of global plastics production is accompanied by

an accumulation of plastic litter in the marine environment (Andrady, 2011; Thiel and Gutow,

2005). Persistent plastic items are rarely degraded but become fragmented over time and are

dispersed by currents and wind (Andrady, 2011; Barnes et al., 2009; Corcoran et al., 2009).

Consequently, marine plastic litter can be found in marine waters all over the globe.

In contrast to interactions of larger organisms with plastics, which are mainly characterised by

the consequences of ingestion or entanglement, the interaction of microorganisms and plastics

are of completely different nature. Plastics function as habitats and are rapidly colonized by

marine microorganisms which form dense biofilms on the plastic surface, the so called

“Plastisphere” (Zettler et al., 2013). Therefore, plastic litter is a substrate which can serve as a

vector for the widespread distribution of a variety of organisms, including harmful algae

species, barnacles, bryozoans (Barnes, 2002; Masó et al., 2003) as well as potentially

pathogenic Vibrio species (Kirstein et al., 2016; Zettler et al., 2013). The persistence of plastics

in marine environments is a matter for debate, and estimates range from hundreds to thousands

of years depending on the chemico-physical properties of the plastic type (Barnes et al., 2009).

“Biofouling refers to the undesirable accumulation of a biotic deposit on a surface” (Characklis,

1991) and can play a major role in controlling plastic buoyancy (Lobelle and Cunliffe, 2011).

Additionally, biofouling also lead to deterioration resulting in fragmentation of larger plastic

items and may also result in degradation of the polymers (Flemming, 1998; Flemming, 2010).

Based on culture-independent approaches, the current state of knowledge regarding the

“Plastisphere” is as follows; microbial communities on marine plastic debris differ consistently

from the surrounding seawater communities (Amaral-Zettler et al., 2015; Oberbeckmann et al.,

2014; Oberbeckmann et al., 2016; Zettler et al., 2013), the plastics community composition is

driven by spatial and seasonal effects (Amaral-Zettler et al., 2015), the community composition

varies with the substrate type (Kirstein et al., 2018; Oberbeckmann et al., 2014), and plastics

biofilm composition is dependent on the habitational conditions, e.g. harbour vs. offshore (De

Tender et al., 2017). Overall, the composition of marine plastics biofilms is probably resulting

from a unique interaction of various factors such as the substrate type, the surrounding

environment, the geographical location and the seasonal variation of environmental parameters.

However, it is well established that several prokaryotic families build the general plastic biofilm

community. These include Flavobacteriaceae, Erythrobacteraceae, Hyphomonadaceae and

Rhodobacteraceae found in the North Sea, the coastal Baltic Sea, multiple locations in the

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42

North Atlantic, and freshwater systems (De Tender et al., 2017; Oberbeckmann et al., 2018;

Zettler et al., 2013).

Recent studies investigated the specificities of plastics communities comparing different types

of plastics with other substrates such as wood or glass (Kettner et al., 2017; Kirstein et al., 2018;

Oberbeckmann et al., 2018; Oberbeckmann et al., 2016). Comparing the PET and glass

associated microbiome, Oberbeckmann et al. (2016) could not detect significant differences in

community composition after 5 to 6 weeks of incubation. In contrast, Kirstein et al. (2018)

found significant differences between the community composition associated to diverse plastics

and glass investigating mature biofilms (15 month). However, the differences in community

composition were generally low, indicating that the shared core of the various biofilms is rather

substrate unspecific. Furthermore, the strongest contribution to the total dissimilarity between

the diverse substrates was often given by less abundant operational taxonomic units (OTUs).

All this points towards the importance of rather rare species in plastic associated marine

biofilms (Kirstein et al., 2018). Considering that the competition pressure in mature biofilms

can be particularly high (e.g. for space or nutrients), uncovering those rare species is a necessary

first step to identify microbes that are closely associated/interact with the polymeric surface,

which will select for species able to survive better when the competition pressure decreases.

To date, researchers of the “Plastisphere” have discussed the potential of plastic “specific”

organisms/assemblages to be involved in biodegradation (Amaral-Zettler et al., 2015; Bryant et

al., 2016; De Tender et al., 2017; De Tender et al., 2015; Oberbeckmann et al., 2018;

Oberbeckmann et al., 2014; Oberbeckmann et al., 2016; Zettler et al., 2013). Here, a plastic

“specific” organism/assemblage is discriminating a respective plastic type from another

substrate type. Several microorganisms, including bacteria and fungi, were isolated from

various environments and were reported to have a degradative effect on specific plastic types

(Crawford and Quinn, 2017; Restrepo-Flórez et al., 2014). Regarding assemblages, recently

Syranidou and colleagues developed tailored micro-consortia suggesting that those are capable

of degrading weathered polystyrene (PS) and polyethylene (PE) fragments, respectively

(Syranidou et al., 2017a; Syranidou et al., 2017b).

Microbes generally have the potential to degrade complex organic compounds in various

environments. This is raising the question, why significant differences between diverse plastics

and other inert substrates could not be detected comparing young marine biofilms (Kettner et

al., 2017; Oberbeckmann et al., 2018; Oberbeckmann et al., 2016) or were found to be generally

low between mature marine glass and diverse plastic biofilms (Kirstein et al., 2018). Kirstein

et al. (2018) has evidence for a general marine biofilm core community of abundant bacterial

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43

taxa, which serve as shared core among diverse substrates, indicating that plastic “specific”

microorganisms might be represented by rather rare species. Assuming that these specificities

of plastic biofilms are referring to microbes of the rather rare biosphere and that plastic

“specific” microorganisms are closely attached to the polymeric surface; a three phase stepwise

uncovering experiment was conducted. In Phase 1, nine distinct plastic films and glass as

control were incubated in situ for 21 months in a natural seawater flow-through system. In

Phase 2, a high-pressure water jet treatment technique was applied to remove the upper loosely

attached biofilm layers, to unveil potential plastic “specific” microorganisms. Thereafter, in

Phase 3, those treated films were used as a source for colonisation of the same type of sterile

plastic strips. Illumina sequencing of the hypervariable V3/V4 region of the 16S rRNA gene

was applied to analyse and compare the prokaryotic communities attached to the various

substrates. In addition attached cells were visualized via Scanning Electron microscopy.

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44

Materials and Methods

Phase 1 - Biofilm formation

A three phased experiment to stepwise uncover closely attached rare microbes was conducted

(Fig 1). In Phase 1, biofilm formation was performed on 9 distinct plastic types such as high-

density polyethylene (HDPE), low-density polyethylene (LDPE), polypropylene (PP),

polystyrene (PS), polyethylene-tereaphthalate (PET), polylactic acid (PLA), styrene-

acrylonitrite plastics (SAN), polyurethane prepolymer (PESTUR) and polyvinyl chloride

(PVC) (Table S1) highly abundant in the marine environment and on glass slides as a neutral

control for 21 month in the dark (max. light intensity 0.1033 µmol/m2/s) in a natural seawater

flow-through system located at the “Biologische Anstalt Helgoland” (North Sea, Germany,

Latitude 54.18286, and Longitude 7.888838) approximately 60 km off the German coastline.

North Seawater was directly pumped through the system (flow rate of approx. 5800 l/day).

Fig 1 Experimental design. Schematic presentation of the three phased stepwise uncovering experiment of

potential plastic “specific” bacteria.

Phase 2 - Removal of the “upper” biofilm layers by high pressure treatment

In order to remove the upper biofilm layers in Phase 2 of our stepwise experiment (Fig 1), a

high-pressure treatment technique was developed to remove the loosely attached biofilm layers.

This was performed with a mini high-pressure cleaning device (Lico-Tec; Arnstorf, Germany)

established to shot (Fig 2a) sterile seawater (0.2 µm filtered and autoclaved) vertically onto the

biofilm associated to the different substrates. Seawater was shot with a working distance of 1

cm for 2 minutes at 4 bar. Next, to evaluate and compare how many cells were still attached on

each substrate after the high-pressure treatment, cell counting of all samples was performed.

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45

Therein, staining with propidium iodide (PI) and SYBR® Green allowed distinguishing

between membrane intact and membrane damaged cells (Figs 2g, h). The treatment was

repeated 9 times on each plastic foil with every sample in triplicates. Fluorescence microscopy

was investigated with the optical microscope Axioplan2, imaging (Zeiss; Oberkochen,

Germany). Detection of the total cell number stained with fluorescent dye SYBR® Green was

performed with the filter set 09 (Zeiss; Oberkochen, Germany). To evaluate the proportion of

damaged cells, the filter set 20 has been applied (Zeiss; Oberkochen, Germany). Detailed

information on the development of the high-pressure treatment technique, staining, and

visualization can be found in the supplement. ImageJ has been used for cell counting (Collins,

2007).

Phase 3 - Selective enrichment on distinct plastics

In order to enrich the uncovered potential plastic “specific” microorganisms a re-colonization

experiment was designed. Therefore a strip of ̴ 1 cm2 with associated 21 month old biofilm of

each of the substrates were treated for 2 minutes and 4 bars with the high-pressure device by

moving the strip slowly under the stream. These strips with the remaining closely surface

attached microorganisms were transferred into sterile glass Petri dishes with 40 ml sterile

filtered and autoclaved North Seawater. For each of the nine plastic types and glass, new ethanol

sterilised strips of the same size were added to these Petri dishes and incubated at 18°C in the

dark. All different substrate strips were sterilized in 70% ethanol and air dried before being

placed in the Petri dishes. After six weeks the re-colonization source was removed (short-term).

Fresh sterile seawater was provided every four weeks. After 60 days one strip of each substrate

was taken for visualization via SEM. After five months of incubation five replicates of each

long-term incubated substrate was taken for DNA extraction followed by 16S gene tag

sequencing.

Scanning Electron Microscopy

Scanning electron microscopy was used to visualize the colonized plastics. Strips of each re-

colonized substrate of about 0.5 cm2 with the attached cells were fixed at 4 °C in sterile seawater

containing 2.5% glutaraldehyde and 50 mM sodium cacodylate (pH 7.2). Samples were stored

in the fixative at 4 °C (4-10 days) until processing for scanning electron microscopy. The

samples were stepwise dehydrated with ethanol bath series of 10 min each at concentrations of

30%, 50%, 70%, 90%, followed by 3 baths of 10 min in 100% ethanol. Samples were

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46

immediately critical point dried (BAL-TEC CPD 030). All samples were sputter coated (BAL-

TEC SCD 005) with gold-palladium before observing with a field emission scanning electron

microscope (JEOL JSM-7500F) with the in-lens detector (SEI-detector) at 5kV and a working

distance of 8 mm.

DNA extraction & 16S Illumina tag sequencing

After five month of selective enrichment, the DNA of microbial biofilms of the nine different

short- and long-term incubated substrates was extracted using the PowerBiofilm® DNA

Isolation Kit (MOBIO Laboratories, Carlsbad, CA) according to the manufacturer's protocol,

including mechanical pulping (FastPrep® FP 120, ThermoSavant,Qbiogene, United States) for

40 seconds on level 4.0. DNA quantity was determined photometrically with a PicoGreen assay

(Invitrogen, Waltham, MA) in duplicates using a Tecan Infinite M200 NanoQuant microplate

reader (Tecan, Switzerland).

16S rRNA gene tag sequencing of the V3 / V4 fragment of the 16S rRNA was performed at

LGC Genomics GmbH (Berlin, Germany). DNA fragments were amplified using amplification

primers 341F (5’-CCTACGGGNGGCWGCAG-3’) and 785R (5’-

GACTACHVGGGTATCTAATCC-3’) (Klindworth et al., 2013). Primers also contained the

Illumina sequencing adapter sequence and a unique barcode index. Resulting amplicons were

paired-end sequenced 2 x 300 bp on an Illumina MiSeq platform. Paired-end reads were merged

using BBMerge 34.48 software (http://bbmap.sourceforge.net/) and processed through the

SILVAngs pipeline (Quast et al., 2013). Sequences were de-replicated at 100% identity and

further clustered with 98% sequence identity to each other. Representative sequences from

operational taxonomic unit clusters (OTUs) were classified up to genus level against the SILVA

v128 database using BLAST as first described by Ionescu et al. (Ionescu et al., 2012).

Sequences having an average BLAST alignment coverage and alignment identity of less than

93% were considered as unclassified and assigned to the virtual taxonomical group “No

Relative" (Quast et al., 2013). Finally, 1,307,882 (99.77%) classified sequences were obtained.

For following downstream analyses, classifications on the genus-level were used to generate

the final abundance matrixes. All classifications contained the sum of all sequences represented

by OTUs with the equal taxonomic path. The raw sequence data is available in the European

Nucleotide Archive (Toribio et al., 2017) under the accession number PRJEB30284, using the

data brokerage service of the German Federation for Biological Data (Diepenbroek et al., 2014),

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47

in compliance with the Minimal Information about any (X) Sequence (MIxS) standard (Yilmaz

et al., 2011).

Statistics and Downstream Data Analysis

To see whether the data of the cell counts were normally distributed R statistical software with

the nlme package has been used. Generalized linear models (GLM) were used to explain the

variability of the attached cells during the establishment of the final LicoJet treatment as well

as in the viability assay. GLM are used in statistics to generalize linear regression with variables

that have an error distribution not normally distributed (McCullagh and Nelder, 1989).

Species richness (S) of the bacterial communities on different short- and long-term incubated

substrates was calculated based on read counts of operational taxonomic units (OTUs).

For beta diversity analysis, first the virtual taxonomical group “No Relative” was removed from

further analysis. Next, counts per classification were normalized by calculating their relative

abundances to the total number of SSU rRNA gene reads per sample. OTUs with a minimal

mean relative abundance of less than 0.1% in at least one substrate type were excluded.

Permutational multivariate analysis of variance (PERMANOVA) was used to test for

statistically significant variance among the source and re-colonized communities attached to

the different substrates. PERMANOVA was carried out with fixed factors and 9999

permutations at a significance level of p < 0.05. Homogeneity of dispersion (PERMDISP) was

applied, to test whether data in significant PERMANOVA results were not over dispersed,

using 9999 permutations at a significance level of p < 0.05. To visualize patterns of samples

regarding various substrates, source and re-colonized communities, principal coordinates

analysis (PCO) using Hellinger distance (D17; (Legendre and Legendre, 1998)) was performed.

To determine OTUs that discriminated the various re-colonized substrates from each other

similarity percentage analysis (SIMPER) was applied. SIMPER was performed using Bray

Curtis similarity (S17) with fourth root transformed relative abundances.

For shade plot creation of unveiled plastic “specific” taxa, first all OTUs with a mean relative

abundance of at least 0.1% present on both, plastics and glass, were rejected. Next, OTUs

contributing most (> 3%) to the total dissimilarity between different plastic groups (SIMPER

analysis) were subjected into cluster analysis. This trimmed data set resulted in 23 OTUs to that

the moderate square root transformation was applied. To determine which groups of plastics

cluster together in respect of plastic “specific” taxa, hierarchical cluster analysis was performed

using Bray Curtis similarity (S17) using square root transformed relative abundances. To test

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48

our hypothesis that specificities of plastics biofilms might be related to members of the rather

rare biosphere the plastic “specific” OTUs were compared with a former dataset of 15 month

old biofilms origin of the same experimental set up (Sequence data deposited in the European

Nucleotide Archive under the accession number PRJEB22051).

Alpha diversity, PERMANOVA, PERMDISP, PCO, SIMPER and CLUSTER analysis were

carried out with the Primer 7 software package plus the add-on package PERMANOVA+

(PRIMER-E Ltd, UK).

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49

Results

Evaluation of adherent cells

Cell counts revealed that after the high-pressure treatment both, cells with intact and damaged

membranes were still attached to the different plastics (Fig 2b). A total cell count of the adhesive

cells on each substrate revealed that attachment occurred to the largest extent on PP followed

in the range of LDPE, PS, HDPE, PESTUR, PVC, SAN, PLA, PET and at least on glass.

Furthermore it is noticeable that mostly the mean of membrane damaged cells exceed the mean

membrane intact cells except for PP, PET and PLA. The mean cell numbers of PP by far

outnumbered the cell counts of all other substrates (Fig 2b). Both states, of membrane damaged

and intact cells were significantly dependent on the substrates (Table S3).

Fig 2 High-pressure water Jet treatment with the a) high pressure treatment device. b) Barplot of the enumerated

mean of adherent membrane intact (green) and membrane damaged (red) cells after a high pressure treatment at 4

bar for 2 minutes, vertical bars denote the Standard Error. Photograph of the 21 month old biofilm attached to c)

Polylactic acid and d) Low density polyethylene. Resulting spots e; f) in respective biofilms after high pressure

treatment. Double stained (SYBR Green & PI) cells on respective substrate g; h) after high pressure treatment

with, scale bars are 10 µm.

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Scanning electron microscopy of colonized plastics

To prove successful colonization, after 60 days of incubation in sterile seawater, plastic strips

were visualized by SEM (Fig 3). Examination of the plastic strips by SEM confirmed re-

colonization of all substrates and provided a closer picture of the microbes attached to the

diverse substrate surfaces (Fig 3).

Fig 3 SEM images of colonized plastics. a) Meshwork of morphological diverse cells embedded in EPS attached

to PS. b) Colony attached to PESTUR c) Single cells of rods and cocci on HDPE d) Consortia of rods and cocci

embedded in EPS on PS e) Rod with spore, comma and spiral cells on PVC.

Various microbial species of different morphologies connected through a network of EPS or

solely distributed across the surface without visible adhesive structures were observed (Fig 3).

Exemplarily, Figures 3a), b) and d) are showing morphological diverse bacteria embedded in

EPS building colonies on the polymeric surfaces of PS and PESTUR. Fig 3c) shows rods and

cocci attached to HDPE and on Fig 3e) three single cells of different morphologies present on

PVC are shown. These organisms were not identified but 16S rRNA gene tag sequencing data

provided evidence that communities varied distinctly between the different substrates.

Selective enrichment & community analysis

For selective enrichment the high-pressure treated plastics (comprising the attached source

community) were incubated with newly provided strips of the same polymer kind. The source

community strips were removed after six weeks (short-term) of incubation and after further five

month (long-term) of selective enrichment the taxonomic composition of the bacterial

communities on the diverse substrates were analysed in detail by 16S rRNA gene tag

sequencing. The species richness of the different samples, analysed by calculating the number

of observed OTUs (number of species (S)) and Margalef`s species richness (d) (Fig 4, Table

S6), showed that the short-term communities had a higher richness compared to the long-term

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51

communities on all substrates but glass (Fig 4), what point towards a selection of “specific”

microbes on the respective plastic type.

Fig 4 Richness of the bacterial communities attached to the diverse substrates based on the number of observed

OTUs. Vertical bars denote the standard deviation (nshort-term=1; nlong-term=5).

Principle coordinate analysis was used to visualize the similarities and dissimilarities between

the various short- and long-term communities (Fig 5). First, all samples of all substrates were

clearly divided (Fig 5). Second, the short-term communities of HDPE, LDPE, PP, PS and PVC

clustered nearby their related long-term communities whereas the short-term community of

glass, PLA, PESTUR, SAN and PET clustered more distant to their long-term communities

(Fig 5). However, the first two axes merely represent 38.8% of the total variation within the

analysed communities. PERMANOVA analysis confirmed that the selective enriched long-

term communities differed significantly between all colonized substrate types (p<0.05; pairwise

PERMANOVA, Table S5).

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Fig 5 Principle Coordinate Ordination relating variation in the community composition between different short-

and long-term incubated substrates. PCOs representing similarity of biofilm communities based on relative

abundances of OTUs across samples. OTUs with a mean relative abundance of at least 0.1% in one substrate type

(nshort=1; nlong=5) were analysed. The different colours indicate the respective substrate, filled symbols represent

short-term samples, open symbols long-term samples. Arrows connect short- and long-term samples of the

respective substrate.

The bacterial community of short- and respective long-term incubated substrates displayed a

change in community composition during the time of selective enrichment. Overall, Alpha- (18-

53%) and Gammaproteobacteria (20-75%) displayed the highest relative abundances in all

samples of all substrates (Fig 6). Some classes were abundant in the short-term communities

but nearly disappeared over the time of selective enrichment e.g. the class of

Epsilonproteobacteria on PLA or Cytophagia on SAN (Fig 6). Vice versa, some classes showed

lower abundances in the short- than in the long-term samples e.g. Flavobacteria on PP, PET

and glass. The class of Sphingobacteria appear to be characteristic for PS as this class was

nearly equally abundant in the short- and in the long-term samples (Fig 6).

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Fig 6 Biofilm community composition based on abundance profiles of the short- and long-term communities on

the class level on different plastics and glass. OTUs with a mean relative abundance of at least 0.1% in one substrate

type (nshort=1; nlong=5) were analysed. A * indicates the term “unclassified”, a # indicates the term “Incertae

Sedis”.

Uncovered plastic “specific” bacteria

Long-term enriched communities associated with different substrates differed between 35-66%

from each other (Table S7). For hierarchical clustering OTUs with a mean relative abundance

of at least 0.1% present on both, plastics and glass were rejected, resulting in 68 OTUs (Fig

S3). To visualize patterns of mostly discriminating members, OTUs jointly contributing with a

minimum of 3% (max. dissimilarity between plastics = 6.07%), to the total dissimilarity

between different plastic groups (SIMPER analysis) were subjected into cluster analysis.

Accordingly, the trimmed data set resulted in 23 mostly discriminating and therefore potential

plastic “specific” OTUs (Fig 7). The hierarchical clustering of the potential plastic “specific”

OTUs indicated closest relatedness of HDPE and LDPE (polyolefins) as well as of PS and SAN

(styrenes), whereas e.g. PVC cluster clearly away from all other plastics (Fig 7). This

differences or similarities are caused by the presence or absence of particular OTUs, or related

to differences in relative abundances of OTUs in common. The main reason for the distinctness

of PVC is an OUT assigned to the genus Flexithrix, with relative abundances of >5% on PVC

(Fig 7, S3). The genus Hirschia and Erythrobacter contributed to the dissimilarity between

PESTUR and all other plastics (Fig 7, S3). Whereas an OUT assigned to the uncultured

Phyllobacteriaceae contributed to the similarity between the polyolefins HDPE, LDPE and PP.

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54

Fig 7 Shade Plot of plastic “specific” OTUs (indicated by numbers) on different long-term plastics and

comparison of their relative abundance in untreated mature biofilms of the same experimental set up after 15

month. Abundant OTUs (mean relative abundance <0.1%; n = 50) are indicated in turquoise, rather rare OTUs

(mean relative abundance >0.1%; n = 50) are indicated in black. Shade Plot creation was based on square root

transformed relative abundances. OTUs with a mean relative abundance of at least 0.1% in one substrate type

(n=5) were analysed. Displayed are OTUs jointly contributing, with a minimum of 3%, to the total dissimilarity

between different plastic groups (SIMPER analysis). OTUs with a mean relative abundance of at least 0.1% present

on both, plastics and glass, were rejected. The amount of contribution is indicated by the colour of cells, lighter

colours represent higher contributions. A * indicates the term “unclassified”, # indicates the term “uncultured”.

Comparison of the resulting 23 OTUs with a former dataset of 15 month old biofilms attached

to the same substrates (Kirstein et al., 2018) revealed that 16 out of the 23 OTUs related to the

rather rare biosphere (relative abundance <0.1%) including Oceanococcus (OUT 1112),

Nannocystaceae (OUT 799), Polycyclovorans (OUT 1045), Phyllobactereacea (OUT 524),

Labrenzia (OUT 572), Maricaulis (OUT 463), Simiduia (OUT 885), Winogradskyella (OUT

198), Dokdonia (OUT 156), Spongiibacter (OUT 905), Roseovarius (OUT 611),

Congregibacter (OUT 889), Planctomycetes SPG12-401-411-B72 (OUT 442), Hirschia (OUT

460), Erythrobacter (OUT 687) and Flexithrix (OUT 120). Seven OTUs assigned to Aquibacter

(OUT 145), Ulvibacter (OUT 197), Planctomycetes OM190 (OUT 406), Planctomycetes BD7-

11 (OUT 405), Parvularcula (OUT 477) Saprospiraceae (OUT 230) and Rhizobiales OCS 116

(OUT 510) showed relative abundances >0.1% in the mature biofilms (Fig 7).

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55

Discussion

Identification of microbes that preferentially colonize and interact with plastics surfaces

remains challenging as the differences in community composition of mature biofilms are

generally low (Kirstein et al., 2018). Furthermore, young biofilms (2-6 weeks) appear to be

rather unspecific between different plastic types or other inert substrates like glass (Kettner et

al., 2017; Oberbeckmann et al., 2018; Oberbeckmann et al., 2016). Here, we present a three

phase experimental approach to uncover potential plastic “specific” microbes. Our findings

indicate that tightly attached microorganisms might account to the rather rare biosphere and

suggest the presence of plastic “specific” microorganisms/assemblages which could possibly

benefit from the given plastic properties.

Water Jet treatment & Selective enrichment

As the main hypothesis of this study was that plastic “specific” microorganisms are tightly

attached to the polymeric surface, a technique to remove the upper loosely attached part is the

first step to facilitate further analysis. There are numerous studies trying to achieve a complete

sanitation of biofilms but, to the best of our knowledge, no method can successfully achieve

entire detachment (Meyer, 2003). The persistency of biofilms towards removal techniques, as

inauspiciously as it may be for sanitation issues, is of great advantage to investigate these

strongly adherent cells on the substrate. Techniques to remove the cohesive layers of the

biofilm, while leaving the adhesive layer attached to the substrate on purpose is not published.

Since chemical or enzymatic action can break adherent bonds, removal of the coherent biofilm

layers requires mechanical action which does not seem to have much influence on the biofilms

integrity (Simoes et al., 2004; Simoes et al., 2010). Microscopic investigations revealed that

strongly attached microbes were able to survive the high-pressure water Jet treatment on all

plastics with the largest extent of adhesive cells on PP followed by LDPE, PS, HDPE, PESTUR,

PVC, SAN, PLA, PET and at least on glass. Already in 1979, Fletcher and Loeb (1979)

examined substrates with a hydrophilic and positive to neutral surface charge, revealing a

moderate number of cells, while only very few cells stayed attached to hydrophilic and

negatively charged surfaces such as glass. This might explain the variation in cell numbers

between the diverse substrates as well as the low cell numbers found on glass compared to those

on the nine different plastic types after the high pressure removal in this study.

Differentiated communities (short-term vs. long-term) developed within the third phase of the

experiment after five month under nutrient limited conditions in the sterile seawater incubation.

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56

PERMANOVA pairwise comparison indicated that all microbial communities on their

respective substrate differed significantly to each other. Although all substrates were treated

similarly, it should be noted that differences in community profiles could be induced by the

considerable difference in remaining cell numbers on diverse substrates after the high-pressure

water Jet treatment. However, the detected differences still imply that the substrate shaped the

community as a result of the adherence strength of the biofilm to the respective substrate

surface. Comparing short-term (six weeks) and long-term (five month) incubated communities

revealed shifts towards communities with lower richness over time for all plastic types but

glass, which points towards a selection of microbes, that are either specialised to low nutrient

conditions or the respective plastic type. On the class level, three different changes were

observed between short- and long-term incubated communities; a shift from high to low

abundant classes, and vice versa, but also classes being characteristic for a plastic type,

implying that the plastic type is responsible for shaping the community composition. Biofilm

communities include a heterogeneity in form of organisms with various metabolic capacities

and different physiological properties which generates on the one hand competition but also

provides on the other hand opportunities for cooperation (Flemming et al., 2016). Hence, some

of the observed changes in community composition might be related to organisms playing a

specific role in interspecies interactions (cooperation) in plastic-degrading microbial

assemblages.

Potentially plastic “specific” microbes

The three phases stepwise uncovering of potential plastic “specific” bacteria resulted in 23 final

OTUs contributing highly to the total dissimilarity between the nine plastic types. Generally,

the chemical composition (e.g. polyesters, polyolefines) and physico-chemical properties of

different plastic types, including the ones used in this study, are highly diverse in order to meet

the different needs of thousands of end products (PlasticsEurope, 2018). The plastic foils used

as substrate in the present study, are commonly produced for e.g. packaging and construction.

It was hypothesized that plastic “specific” microorganisms are tightly attached to the polymeric

surface and might be represented by rare but active species, since differences of mature biofilms

between distinct plastic types were found to be generally low (Kirstein et al., 2018). Comparing

our two datasets revealed that 70% of the uncovered potential plastic “specific” OTUs of the

present study, were assigned to the rather rare biosphere (<0.1%) of the biofilms investigated

six month earlier (15 month old biofilm (Kirstein et al., 2018)). Former research reports that

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57

rare phylotypes tend to stay rare (Galand et al., 2009; Kirchman et al., 2010). Other studies

suggested that rare but active populations might be controlled by top-down forces (e.g.

predation) or competition (e.g. space, nutrients) within the biofilm and to underlie

environmental controls (e.g. temperature) (Andersson et al., 2010). Having the potential to

increase in abundance (Besemer et al., 2012), our findings clearly support the idea that potential

plastic “specific” species are, at least partly, controlled by competitive interactions in mature

dense biofilms.

Several studies have investigated microbial communities on marine plastics under various

conditions (Amaral-Zettler et al., 2015; Bryant et al., 2016; De Tender et al., 2017; De Tender

et al., 2015; Oberbeckmann et al., 2018; Oberbeckmann et al., 2016; Viršek et al., 2017; Zettler

et al., 2013). After subtracting OTUs also abundant on glass, in total 68 OTUs were found

specifically associated with the different plastics and, out of those, 23 mostly discriminating the

chemically distinct plastics. Several researchers, reported about multiple families in common

on a variety of marine plastics in different locations e.g. Nannocystaceae, Flavobacteriaceae,

Planctomycetes, Saprospiraceae, Erythrobacteraceae, Hyphomonadaceae and

Rhodobacteraceae (Bryant et al., 2016; De Tender et al., 2017; Kirstein et al., 2018;

Oberbeckmann et al., 2018; Oberbeckmann et al., 2016; Viršek et al., 2017; Zettler et al., 2013).

Members of these families were also present within the 23 most discriminating OTUs in this

study. Two OTUs were discriminating PESTUR from all other plastics assigned to Hirschia

(Hyphomonadaceae) and Erythrobacter (Erythrobacteraceae). Several studies have previously

reported about the abundance of these two families associated to diverse plastics in different

experimental approaches and locations (De Tender et al., 2017; Oberbeckmann et al., 2018;

Zettler et al., 2013). Recently, Oberbeckmann et al. (2018) reported about the two families

Hyphomonadaceae (mostly Hyphomonas) and Erythrobacteraceae (mostly Erythrobacter),

being exclusively abundant in two weeks old biofilms on PE and PS. The genera Erythrobacter

and Parvularcula were reported to be part of plastic biofilms in the North Atlantic and North

Adriatic Sea (Viršek et al., 2017; Zettler et al., 2013). In our study one OTU belonging to the

family Saprospiraceae was highly discriminating PS from the other plastics. Oberbeckmann et

al. (2018) also detected members of this family on diverse substrates, PE and PS just being one

of them. Phyllobacteriaceae were found to be significantly more abundant on plastics, despite

showing overall high relative abundances in the study of Oberbeckmann et al. (2018). In our

study Phyllobacteriaceae contributed to the similarity between the polyolefins HDPE, LDPE

and PP.

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58

Since bacterial families are large, the uncovered plastic “specific” genera were compared with

former studies and found four genera already recognized on marine plastics as Roseovarius

(Viršek et al., 2017), Erythrobacter (Oberbeckmann et al., 2018; Viršek et al., 2017; Zettler et

al., 2013) Ulvibacter (Oberbeckmann et al., 2016) and Parvularcula (Viršek et al., 2017; Zettler

et al., 2013). The repetitive detection of these genera associated to marine plastics in various

approaches suggests that further investigations on their role in plastic biofilms are required.

However, since our experimental design focused on the enrichment of tightly attached and

rather rare taxa, they might have been present but not recognized in previous research. For

example, in the study of Kirstein et al. (2018) the genera Roseovarius and Erythrobacter

accounted to the rare biosphere (<0.1%) in the mature biofilms (15 month) and were therefore

further not considered. Interestingly, in other studies Roseovarius and Erythrobacter were

detected in relatively young (2 weeks) or in biofilms of unknown age (Oberbeckmann et al.,

2018; Viršek et al., 2017; Zettler et al., 2013).

Sensing the surface – plastic properties

Sensing of a non-soluble surface followed by the successful colonization are the first steps for

marine bacteria to develop a community, potentially leading to plastic biodegradation (Dang

and Lovell, 2016; Sivan, 2011). Beside surface properties like hydrophobicity and roughness,

surface chemodynamics like surface conditioning or nutrient enrichment also play a role in

forming distinct biofilm communities (Dang and Lovell, 2016). This questions whether we, and

other researcher, were detecting “plastic specific” organisms or “plastic specific coatings”

organisms needs to be addressed in future studies. In the present study, bacterial taxa able to

survive on glass likely used dissolved organic carbon present in the sterile seawater as carbon

source, and consequently did not benefit from plastics surface properties or chemical

composition. All other OTUs detected on the various plastic types were therefore potentially

plastic “specific”. Due to short read lengths of 16S rRNA gene tag sequencing, a conclusive

identification on the species level of the unveiled plastic “specific” OTUs was not possible so

far. Since successful surface colonisation does not prove a special role as e.g. plastic

degradation, the next step must be the systematic isolation and identification of those plastic

“specific” organisms and to further test for the potential of one species or consortium to degrade

the respective plastic type. On the community level, the next steps should be the disclosure of

the mechanisms that allow the plastic “specific” assemblages to survive, their possible

metabolic pathways and enzymes involved.

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Conclusion

This study represents a systematic and robust experimental approach uncovering potential

plastic “specific” microbes and is therefore a step forward in understanding the substrate

specificity of the “Plastisphere”. For the first time, a high-pressure water Jet treatment technique

was used to remove the cohesive layer of mature biofilms, while leaving the adhesive layer on

the plastics surface. Our results indicate the presence of plastic “specific”

microorganisms/assemblages which could possibly benefit from the given plastics properties.

Furthermore, our findings clearly indicate that plastic “specific” microorganisms might account

to the rather rare biosphere and are tightly surface attached. Underrepresentation, due to low

read counts, might be an explanation why specificities between plastics biofilms in natural

marine environments were not detected so far in young biofilms or seem to be generally low in

mature biofilms.

Acknowledgments

We would like to thank Dr. Cédric Meunier for fruitful discussions. This work was founded by

the Alfred Wegener Institute, Helmholtz Centre for Polar and Marine Research and supported

by the German Federal Ministry of Education and Research (Project BASEMAN - Defining the

baselines and standards for microplastics analyses in European waters; BMBF grant

03F0734A).

CHAPTER III

Dangerous Hitchhikers?

Evidence for potentially pathogenic Vibrio spp. on microplastic particles

Inga V. Kirsteina*+, Sidika Kirmizia+, Antje Wichelsa, Alexa Garin-Fernandez a, Rene Erlera,

Martin Löder a, b, Gunnar Gerdts a

aAlfred-Wegener-Institute Helmholtz Centre for Polar and Marine Research, Biological Station

Helgoland, Helgoland, Germany

bAnimal Ecology I, University of Bayreuth, NWI 5.0.01.43.1, Bayreuth, Germany

*Corresponding author: Inga Kirstein, Alfred-Wegener-Institute Helmholtz Centre for Polar

and Marine Research, Biologische Anstalt Helgoland, Postbox 180, 27483 Helgoland,

Germany, Tel.: +49 (4725)819-3233; fax: +49 (4725)819-3283 e-mail: inga.kirstein@awi.de

+ These authors contributed equally to the manuscript

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Abstract

The taxonomic composition of biofilms on marine microplastics is widely unknown. Recent

sequencing results indicate that potentially pathogenic Vibrio spp. might be present on floating

microplastics. Hence, these particles might function as vectors for the dispersal of pathogens.

Microplastics and water samples collected in the North and Baltic Sea were subjected to

selective enrichment for pathogenic Vibrio species. Bacterial colonies were isolated from

CHROMagarTMVibrio and assigned to Vibrio spp. on the species level by MALDI-TOF MS

(Matrix Assisted Laser Desorption / Ionisation - Time of Flight Mass Spectrometry). Respective

polymers were identified by ATR FT-IR (Attenuated Total Reflectance Fourier Transform -

Infrared Spectroscopy). We discovered potentially pathogenic Vibrio parahaemolyticus on a

number of microplastic particles, e.g. polyethylene, polypropylene and polystyrene from North

/ Baltic Sea. This study confirms the indicated occurrence of potentially pathogenic bacteria on

marine microplastics and highlights the urgent need for detailed biogeographical analyses of

marine microplastics.

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Introduction

The production of synthetic polymers started over 100 years ago and meanwhile the worldwide

production reached up to 311 million tons per year (PlasticsEurope, 2015). As a consequence

of improper disposal synthetic polymers represent the most rapidly growing form of

anthropogenic debris entering and accumulating in the oceans (Andrady, 2011; Thiel and

Gutow, 2005).

Due to their durability most synthetic polymers are poorly degradable in the marine

environment but become brittle and subsequently break down in small particles, so called

microplastics (Andrady, 2011; Corcoran et al., 2009). Several size categorizations of plastics

have been suggested by various researchers (Gregory and Andrady, 2003; Moore, 2008) while

plastic fragments smaller than 5 mm are categorized as microplastics by Barnes et al. (2009).

Once floating on seawater, plastic debris can be transported over long distances by wind,

currents and wave action (Barnes et al., 2009).

As all surfaces in the marine environment microplastic is rapidly colonized by bacteria

(Harrison et al., 2014) and subsequently by a plethora of organisms building up complex

biofilms (Dobretsov, 2010). Harrison et al. (2014) detected bacterial colonization of low density

polyethylene microplastics already after 7 days exposure in marine sediments. Also (Lobelle

and Cunliffe, 2011) proved biofilm formation on plastics after 1 week of incubation in seawater

via quantitative biofilm assays. Prior studies evidenced that even harmful algal species were

detected in biofilms on plastic debris (Masó et al., 2003). Being highly heterogeneous

environments, biofilms offer important ecological advantages such as the accumulation of

nutrients, as protective barrier, for mechanical stability (Flemming, 2002) or the formation of

micro-consortia of different species that orchestrate the degradation of complex substrates

(Wimpenny, 2000).

Zettler et al. (2013) showed that microbial communities on marine plastic debris differ

consistently from the surrounding seawater communities and coined the term “Plastisphere” for

this habitat. Furthermore, Amaral-Zettler et al. (2015) reported that “Plastisphere” communities

are genetically unique from the free marine water communities that envelop them and possess

dominant taxa that are highly variable and diverse. Moreover, the composition of biofilm

communities on plastic in marine habitats varies with season, geographical location and plastic

substrate type (Oberbeckmann et al., 2014).

Zettler et al. (2013) have suggested that plastic particles may serve as vectors for the dispersal

of human pathogens (Vibrio spp.). Using a culture-independent approach, the author’s detected

sequences affiliated to Vibrio spp. on marine plastic debris (Zettler et al., 2013), i.e. on plastic

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64

particles in the North Atlantic by using molecular tools (Amplicon Pyrotag Sequencing).

Furthermore, De Tender et al. (2015) recently detected Vibrionaceae on marine plastics from

the Belgian North Sea, by using next-generation amplicon sequencing. However, due to short

read lengths, a conclusive identification on the species level was not provided so far (De Tender

et al., 2015; Zettler et al., 2013).

Species of the genus Vibrio belong to the class Gammaproteobacteria and are highly abundant

in sediments, estuaries and marine coastal waters (Barbieri et al., 1999). Vibrios are gram-

negative, rod-shaped chemoorganotrophic and facultatively anaerobic organisms. Besides

occurring free-living in aquatic environments, Vibrio spp. are known to colonize a variety of

marine organisms, utilizing released nutrients on these living surfaces (Huq et al., 1983; Visick,

2009) or living in symbiosis (McFall-Ngai and Ruby, 1991; McFall-Ngai, 2002; McFall-Ngai

and Ruby, 1998). Some Vibrio species are known as animal pathogens invading coral species

and causing coral bleaching (Ben-Haim et al., 2003) and others are classified as human

pathogens causing serious infections (Morris, 2003). Especially V. parahaemolyticus, V.

vulnificus and V. cholerae are known as water-related human pathogens which cause wound

infections associated with recreational bathing, septicemia or diarrhea after ingestion of

contaminated foods (Thompson et al., 2004a).

Although Vibrio infections are common in tropical areas, the last decade showed a significant

increase in documented cases also in European regions, such as in the Mediterranean Sea (Gras-

Rouzet et al., 1996; Martinez-Urtaza et al., 2005) or in the more temperate Northern waters

(Eiler et al., 2006). Prior studies reported that the number of Vibrio infections correspond

closely with the sea surface temperature pointing to a possible link to climate change related

phenomena (e.g. global warming, heat waves) (Baker-Austin et al., 2010; Baker-Austin et al.,

2013).

Böer et al. (2013) reported that V. alginolyticus, V. parahaemolyticus, V. vulnificus and V.

cholerae occurred in water and sediments in the central Wadden Sea and in the estuaries of the

rivers Ems and Weser. The most prevalent species were V. alginolyticus followed by V.

parahaemolyticus, V. vulnificus and V. cholera (Böer et al., 2013), reflecting earlier findings on

the composition of Vibrio communities in other parts of the North Sea (Bauer et al., 2006;

Collin and Rehnstam-Holm, 2011; Hervio-Heath et al., 2002; Schets et al., 2011). While V.

vulnificus and V. cholerae were detected mainly in the Baltic Sea, V. parahaemolyticus occurred

as the main potential pathogenic Vibrio spp. in the North Sea (Böer et al., 2013; Oberbeckmann

et al., 2011b; Ruppert et al., 2004; Schets et al., 2010).

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65

As already mentioned most synthetic polymers are poorly degradable and are rapidly colonized

by microorganisms. Microplastics could be transported over long distances in marine

environments, as compared to naturally occurring polymers, and therefore function as a vector

for the dispersal of harmful or even human pathogenic species. To verify or falsify the

occurrence of potentially pathogenic Vibrio spp. on marine plastics, we analysed plastics and

corresponding water samples of the North and Baltic Sea with respect to potentially human

pathogenic Vibrio spp. by using cultivation-dependent methods (alkaline peptone water (APW),

CHROMagar™Vibrio), followed by state of the art identification of bacteria on the species

level by MALDI-TOF MS (Erler et al., 2015). The main focus of the study was on detecting

the main potentially human pathogenic species V. cholerae, V. parahaemolyticus and V.

vulnificus. Polymers were identified by ATR FT-IR (Attenuated Total Reflectance Fourier

Transform - Infrared Spectroscopy).

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Materials and Methods

Sampling

To detect Vibrio spp. attached to microplastics, neustonic particles were collected during two

research cruises in 2013 and 2014 at 62 sampling stations in the North and Baltic Sea (see Table

S1). Neuston samples were taken with a Neuston Catamaran equipped with a 300 µm net. The

Catamaran was towed alongside the vessel for about 30 to 45 min per station. The volume

passing the Neuston net was recorded by use of a mechanical flowmeter (Table S2). Further

samples were taken at the drift line of the south port beach at the island Helgoland at low tide

in August 2013 (station 63). Particles recovered in the cod end of the Neuston net or sampled

at the drift line of Helgoland were sorted by stereo microscopy and using a Bogoroff chamber

and finally transferred to Petri dishes containing sterile seawater. Single particles identified

visually according to the definition by Barnes et al. (2009) in a size range of 0.5 – 5 mm and to

colour and texture as being synthetic polymers were picked with sterile forceps and washed

three times with 10 ml of sterile seawater, to remove loosely attached organisms.

For comparison of microplastic-attached and waterborne Vibrio spp., additional surface

seawater samples were taken on both research cruises with a thoroughly flushed bucket or

rosette sampler (SBE 911 plus, Sea-Bird Electronics, US) and a maximal volume of 1 l was

filtered onto 0.45 µm sterile membrane filters (Sartorius stedim biotech, US). Environmental

parameters (temperature, salinity) were recorded by a ship-based thermosalinograph (SBE

21SeaCAT, Sea-Bird Electronics, US) or by the sensors of the rosette sampler. The temperature

of Helgoland was measured manually with a thermometer and the salinity was recorded with a

salinometer (Autosal, GUILDLINE, Canada) (Table S3).

Enrichment & isolation of Vibrio spp.

All particles and membrane filters (seawater samples) were immediately transferred

individually into sterile glass tubes with alkaline peptone water (15 ml APW) and incubated in

a rotating incubator at 37 °C for 48 h in the dark for the growth of a broad spectrum of

mesophilic and potentially pathogenic Vibrio spp., enabling their selective enrichment.

After APW incubation the tubes were visually checked for growth and turbid samples were

plated by using an inoculation loop or Spiral-plater (easySpiral® Dilute; Interscience, France)

on selective CHROMagar™Vibrio (MAST Diagnostica GmbH, Germany) (Di Pinto et al.,

2011). All inoculated CHROMagar™Vibrio were incubated at 37 °C for 24 h in the dark. The

appearing colonies were checked with respect to distinct colony colorations typical for V.

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parahaemolyticus, V. vulnificus and V. cholerae according to the manufacturers’ instruction.

Representative colonies for each coloration were picked and differentially streaked out on

marine broth agar (Oppenheimer and ZoBell, 1952) with reduced salinity (MB-50%=16PSU).

Incubation was performed at 37 °C for 24 h in the dark.

Even though CHROMagar™Vibrio is a selective medium for the isolation of V. cholerae, V.

vulnificus and V. parahaemolyticus, other species have the ability to grow on these media

appearing with the same colony colorations. For instance, V. fluvialis occurred in mauve

coloured colonies distinct from V. parahaemolyticus and V. mimicus in turquoise coloured

colonies distinct for V. vulnificus and V. cholerae. Hence for a conclusive identification all

presumptive V. cholerae, V. vulnificus and V. parahaemolyticus strains were further analysed

by MALDI-TOF MS.

MALDI-TOF MS

For MALDI-TOF analysis, all isolates were grown overnight on MB-50% agar plates as

described above. To create high quality mass spectra, proteins of the strains isolated during the

cruise in 2013 were extracted using a previously described formic acid/acetonitrile extraction

method (Mellmann et al., 2008). For fast identification, all other strains (cruise 2014 and

Helgoland samples 2013) were analysed via the direct transfer procedure according to

manufacturers` recommendations (Bruker Daltonics Inc., Germany, Bremen). This involved

picking colonies after 24 hours of cultivation with sterile toothpicks and directly transferring

onto the MALDI-TOF MS target plate (MSP 96 target polished steel) as thin layer. Each sample

spot was first overlaid with 1 µl formic acid (70% v/v) followed by an overlay with 1 µl matrix

solution (saturated solution of α-cyano-4-hydroxycinnamic acid in 50% acetonitrile and 2.5%

trifluoroacetic acid) and directly screened. All spectra were acquired using the microflex LT/SH

system (Bruker Daltonics Inc., Germany, Bremen). Species identification was done by using

the BiotyperTM software (version 3.1) according to the manufacturer’s instructions, where 70

most prominent mass peaks were compared to the mass spectra of the Bruker library as well as

the “VibrioBase” library (Erler et al., 2015).

In order to check the reliability of the species assignment via MALDI-TOF MS all V. cholerae,

V. vulnificus and V. parahaemolyticus were verified by PCR amplification of species-specific

genes and additionally screened for virulence-associated genes (section 2.4).

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PCR of regulatory and virulence-related genes

As described previously (Oberbeckmann et al., 2011a), DNA extraction of Vibrio strains

identified by MALDI-TOF MS was carried out using lysozyme/SDS lysis and

phenol/chloroform extraction, followed by isopropanol precipitation. Prior to PCR

experiments, DNA quantity and quality was determined photometrically (TECAN infinite

M200, Switzerland). Species-specific PCR for toxR genes was performed with all V.

parahaemolyticus, V. vulnificus and V.cholerae strains respectively using the universal forward

primer UtoxF together with the species specific primers VptoxR, VvtoxR and VctoxR,

respectively (Bauer and Rorvik, 2007; Di Pinto et al., 2005). Specific PCRs targeting

thermostable direct haemolysin (tdh) (Nishibuchi and Kaper, 1985) and the tdh related

haemolysin (trh) (Honda et al., 1991; Honda and Iida, 1993) genes were performed with the

primer sets tdhD3F/tdhD1R and trhFR2/trhRR6 to strains assigned to V. parahaemolyticus

(Bauer and Rorvik, 2007; Tada et al., 1992). To test V. cholerae strains for the presence of a

unique chromosomal region indicating the serotypes O139 (Albert et al., 1997) and O1

(Katsuaki Hoshino 1998) and the cholera toxin gene ctxA (Singh et al., 2002) a multiplex PCR

was performed with the primer sets O139F/O139R, O1F/O1R and ctxA1/ctxA2 (Bauer and

Rorvik, 2007; Mantri et al., 2006; Nandi et al., 2000). All reactions were performed in duplicate.

In case of discordant results, a third PCR was carried out. The PCRs were performed as

described by Böer et al. (2013) with the exception that 20 ng of template DNA was used. The

following reference strains were used as positive controls: V. vulnificus ATCC 27562 (VvtoxR)

(The Federal Institute for Risk Assessment, BfR), V. parahaemolyticus RIMD 2210633

(VptoxR; tdh) (German Collection of Microorganisms and Cell Cultures, DSMZ), V.

parahaemolyticus CM12 (tdh; trh), V. parahaemolyticus CM24 (trh) (provided by Carsten

Matz, HZI), V. cholerae CH 111 (VctoxR; O1), V. cholerae CH 187 (VctoxR; O139; ctxA) and

V. cholerae CH 258 (VctoxR; ctxA; O1) (BfR). V. harveyi ATCC 25919 (DSMZ) was used as

negative control in each PCR. PCR products were confirmed to be of the expected size by a

MultiNA Microchip electrophoresis system (MCE-202 MultiNA, Shimadzu Biotech).

FT-IR analyses of particles

After incubation in APW, all particles were rinsed using deionized water and dried at 60°C

overnight. Prior to analysis, particles were rinsed with ethanol (70% v/v) and the surface was

scraped with a scalpel to avoid organic contamination interfering with FT-IR analysis. The FT-

IR spectra of particles were recorded by the attenuated total reflectance (ATR) technique using

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a Tensor 27 spectrometer with a Platinum ATR unit (Bruker, Germany). For each analysis 16

scans in the range 4000-400 cm-1 with a resolution of 4 cm-1 and 6 mm aperture were performed

and averaged. The obtained IR spectra were compared to reference-spectra of an in-house

database covering 143 spectra of different synthetic polymers and the IR Library from Bruker

Optics containing 350 entries. Spectra processing and database comparisons were performed

by using OPUS 7.2. (Bruker, Germany).

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Results

Occurrence and characterization of microplastics

Particles were collected from 39 stations in the North Sea and 5 stations in the Baltic Sea. In

total, 170 particles were collected in the North Sea and 15 particles in the Baltic Sea, mostly

abundant at stations 17, 56, 58 and 61, with ≥ 10 particles from each station, respectively (Table

S4). Almost all particles showed signs of weathering, including cracks and pitting. Most

particles were covered at least partially with dense biofilms on their surface, indicating

colonization by various biota. Polymer identification of presumptive synthetic polymer

particles, (ATR FT-IR (Table S3)) confirmed 141 as synthetic polymers, 14 particles were non-

plastics such as chitin or keratin, and 30 could not be further identified. All of the 15

presumptive microplastics of Helgoland drift line were identified as synthetic polymers. The

most abundant synthetic polymer throughout all sampling sites was polyethylene, comprising

over 40 % of the collected particles at all sites. Polypropylene and polystyrene were also

frequently found at all sites, representing 14-20 % and 5-7 % of all particles, respectively (Fig.

1).

Fig 1: Proportions of synthetic polymers and other particles collected during research cruises in the North and

Baltic Sea and the drift line of Helgoland. Sampling took place in September 2013 (left), July/August 2014

(middle) and July 2013 (right). Particles were characterized using ATR FT-IR spectroscopy. Also given are

numbers of total particles (N) and percentages of polyethylene, polypropylene and polystyrene particles.

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Identification and geographic distribution of Vibrio spp. in water samples

Water samples were taken from all stations in the North and Baltic Sea with the exception of

Helgoland drift line (station 63) resulting in 326 APW enrichment cultures. Out of these, 323

displayed growth and were subjected to further isolation of bacteria on selective

CHROMagar™Vibrio agar plates, with respect to V. cholerae, V. vulnificus and V.

parahaemolyticus.

From all water samples, 151 pure cultures of representative mauve and turquoise blue colonies

were grown on marine broth agar and subjected to MALDI-TOF MS. Out of these, 104 were

identified as Vibrio spp. by MALDI-TOF MS.

With the exception of three isolates, all Vibrio water strains could be identified by MALDI-

TOF MS on a conclusive species level. We identified 38 % out of all Vibrio water isolates (104)

as V. parahaemolyticus, 16 % as V. vulnificus and 11 % as V. cholerae. Further on, 21 % of the

strains were classified as V. fluvialis, 7 % as V. mimicus, 5 % as V. diazotrophicus, 1 % as V.

metschnikovii (Table S6).

A single V. parahaemolyticus strain (VN-4212) isolated from water (station 3) carried the

virulence-associated gene tdh, while trh was not detected in any strain (Table S6). No V.

cholerae strain belonged to the O1/O139 type or carried the ctxA gene.

In general, V. parahaemolyticus was detected only in North Sea waters (Fig. 2) in a temperature

range of 14.9 to 21.1 °C and at salinities between 16.9 to 32.4 PSU (Table S3). The potentially

pathogenic species V. cholerae, V. vulnificus and V. parahaemolyticus occurred mainly in

coastal and estuarine regions of the North Sea. Vibrio fluvialis was the only species that was

detected in open waters in the North Sea (Fig. 2 a, c).

In the Baltic Sea both species, V vulnificus and V. cholerae appeared close to the Polish border

at 14.5 to 14.9 °C and 5.7 - 7.3 PSU (station 36, 37, 38). V. cholerae occurred also nearby to

Rostock at 14.1 °C and 11.7 PSU (station 31) (Fig. 2 b; Table S3). Vibrio fluvialis was detected

once in Baltic surface water inside Germany and Denmark (station 32).

Identification and geographic distribution of Vibrio spp. on microplastics

All collected particles of North Sea, Baltic Sea and Helgoland drift line were subjected to

selective APW enrichment resulting in 200 APW cultures. Out of these 161 displayed growth

and were processed as described previously. From 15 microplastic particles from the North and

Baltic Sea, in total 37 putative (according to the colony colorations) V. cholerae, V. vulnificus

or V. parahaemolyticus strains were isolated. At the drift line of Helgoland 4 putative V.

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parahaemolyticus strains from 4 different microplastic particles were isolated. Of these 41

strains, 22 were identified as Vibrio spp. by MALDI-TOF MS. Thirteen strains were identified

as V. parahaemolyticus (59 %), six as V. fluvialis (27 %) and one as V. alginolyticus (5 %)

(Table S5). Even though we isolated representative coloured colonies neither V. vulnificus nor

V. cholerae could be detected on microplastic particles.

Fig 2: Geographical occurrence of Vibrio spp. On microplastics and surface water of a) the North Sea from

research cruise HE409 on RV Heincke in September 2013 b) the Baltic Sea from research cruise HE409 on RV

Heincke in September 2013 and c) North Sea from research cruise HE430 on RV Heincke in July/August 2014

and the drift line of Helgoland (station 63). ( ) species detected from surrounding seawater ( ) species detected on

microplastic particles.

V. parahaemolyticus was isolated from three polyethylene fibres and four polyethylene

fragments during the cruises in the North Sea at temperatures between 14.8 and 21.1°C and

salinities between 12.6 - 32.4 PSU (Table S3). These were collected in the Ems estuary (station

5), near the uninhabited island Mellum (station 9), the Elbe estuary (station 21), and close to

the Frisian islands (stations 39 and 41) (Fig. 2 a, c). Additionally V. parahaemolyticus was

isolated from two polyethylene films and two polypropylene fragments of Helgoland drift line

at a water temperature of 16.6°C and a salinity of 30.2 PSU (station 63) (Fig. 2 c). V. fluvialis

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was detected on four non-identified particles collected between the UK and the Netherlands

(stations 58, 59) and on a polyethylene fragment of the Weser estuary (Germany, station 11).

V. alginolyticus was detected on one polyethylene fragment close to the Frisian island Juist

(station 41). In the English Channel (station 55) an unspecified Vibrio spp. was detected on a

polyethylene fragment (Fig. 2 c).

One polypropylene film (station 30; Fig. 2b) collected close to the coastal regions of Wismar

in the Baltic Sea at 14.8 °C and 12.6 PSU (Table S3) was colonized by both species, V.

parahaemolyticus and V. fluvialis. Vibrio parahaemolyticus was detected only once on this

single microplastic particle in the Baltic Sea (Fig. 2b).

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Discussion

Although the microbial colonization of marine plastic particles was reported already in the

1970s (Carpenter et al., 1972; Carpenter and Smith, 1972), this issue received increasing

attention in the last years due to the discovery of the large oceanic garbage patches (Kaiser,

2010; Ryan, 2014) and the general perception of microplastics being an emerging

environmental topic of concern. In this context, it was also hypothesized, that microplastics

may function as a vector for dispersion of invasive species including toxic algae but also

pathogenic organisms (Masó et al., 2003; Zettler et al., 2013).

Recently the microbial community on marine plastics was targeted in several studies,

highlighting the composition and diversity of plastic-attached microorganisms (Amaral-Zettler

et al., 2015; Carson et al., 2013; De Tender et al., 2015; Oberbeckmann et al., 2014; Reisser et

al., 2014; Zettler et al., 2013). Within the microbial community on the “Plastisphere” (Zettler

et al., 2013) sequences related to the genus Vibrio, a group of bacteria also containing serious

pathogens, were found (De Tender et al., 2015; Zettler et al., 2013). However, in both studies a

conclusive identification on the species level could not be provided so far due to the usage of

next-generation amplicon sequencing and the short read lengths inherent to the methodology.

In our study we were able to prove the presence of potentially pathogenic V. parahaemolyticus

on twelve floating microplastics for the first time by a selective cultivation approach and

identification on species level by MALDI-TOF MS.

Microplastics in the North and Baltic Sea

In the present study, we observed more microplastic particles in North Sea waters compared to

the Baltic Sea. Up to now, information on the abundance of microplastics in coastal waters of

the North and Baltic Sea is scarce, and a comparison of the findings is problematic due to

missing standard operational procedures (SOP) for sampling, extraction and analysis of

microplastics (Löder and Gerdts, 2015).

During both cruises in 2013 and 2014, 77 % of all collected and identified microplastics as well

as all collected microplastics at the drift line of Helgoland, occurred as fragments with rough

and uneven edges clearly indicating a breakdown of larger plastics (Thompson et al., 2004c).

Brittleness of particles including cracks and pitting could be detected on collected microplastics

which might be the result of degradation processes or wind and wave actions (Andrady and

Neal, 2009). Thus it could be suggested that most of the collected microplastics were exposed

long enough to the marine environment to get brittle and be transported over long distances.

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Thiel et al. (2011) reported hotspots of accumulating microplastics in the North Sea and a rapid

transport through the German Bight due to strong westerly winds. In contrast, based on the

relationship between litter accumulation on Helgoland beaches and southerly winds, Vauk and

Schrey (1987) suggested that these winds might push anthropogenic debris from source regions

which results in accumulation on local beaches. Galgani et al. (2000) proposed that the

predominant northward currents in the eastern part of the German Bight transport floating

debris and accumulate it in an area to the west of Denmark. However due to the focus of our

study (Vibrio spp.), these findings should be interpreted with care since we were not aiming at

monitoring microplastics explicitly and in a systematic way.

By far the majority of microplastics from the North and Baltic Sea as well as from the Helgoland

drift line was identified as polyethylene, followed by polypropylene and polystyrene (Fig. 1).

Prior studies already reported high portions of these three polymers in the course of various

samplings in marine and coastal environments which mirrors our results (Browne et al., 2010;

Moret-Ferguson et al., 2010; Oberbeckmann et al., 2014) and furthermore reflect the usage of

these polymers in the worldwide economy. In the United States polyethylene, polystyrene,

polypropylene and polyethylene terephthalate are the most widely produced and disposed

synthetic polymers (Barnes et al., 2009). In Europe polyethylene and polypropylene are the

synthetic polymers with the highest demand in various application segments, especially in

packaging (PlasticsEurope, 2015).

Vibrio hitchhikers

Biofilm communities on environmental plastic samples were recently characterized in several

studies applying molecular tools. The diverse microbial communities on marine plastic debris

differed clearly from the surrounding seawater (Amaral-Zettler et al., 2015; De Tender et al.,

2015; Oberbeckmann et al., 2014; Zettler et al., 2013).

The herein described presence of potentially human pathogenic Vibrio spp. on microplastics

has to be discussed in the light of these latter studies. The first indication of the presence of

Vibrio spp. on marine microplastics was published by Zettler et al. (2013), who reported the

dominance of this genus that constituted nearly 24 % of the whole biofilm community on a

single polypropylene particle collected from the North Atlantic. In 2015, De Tender et al.

(2015) reported the detection of members of the family Vibrionaceae on marine plastics from

the Belgian North Sea. Recently a review of Keswani et al. (2016) highlights the lack of

knowledge about the persistence of potentially pathogenic Vibrio spp. on plastic debris. Our

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study clearly confirmed the presence of cultivable Vibrio spp. on 13 % of all marine collected

microplastic particles. Amongst others, potentially pathogenic V. parahaemolyticus strains

were detected on 12 microplastic particles. Only collected polyethylene, polypropylene and

polystyrene fragments were colonized by Vibrio spp.

In general Vibrio spp. tends to colonize marine biotic surfaces like corals or zooplankton /

phytoplankton surfaces. V. cholerae strains, both O1 and non-O1 serovars, as well as V.

parahaemolyticus strains were found to be attached to the surfaces of copepods in natural waters

(Huq et al., 1983). In comparison to naturally occurring polymers like chitin, synthetic polymers

are poorly degradable and could therefore function as a mechanism for the transport and

persistence of Vibrio species. (Pruzzo et al., 2008) reviewed substrate-specificity of V. cholerae

on the naturally occurring polymer chitin. They reported close interactions between V. cholerae

and chitin surfaces in the environment including cell metabolic and physiological responses e.g.

chemotaxis, cell multiplication, biofilm formation, and pathogenicity. With respect to plastic

microbial communities, Oberbeckmann et al. (2014) found that the structure and taxonomic

composition of these plastic associated communities vary with plastic type, but also with

geographical location and season. Moreover, Amaral-Zettler et al. (2015) found that

“Plastisphere” communities of the Atlantic and Pacific Ocean clustered more by geography

than by polymer type, with exception of polystyrene that showed significant differences to

polyethylene and polypropylene.

The substrate specificity of Vibrio spp. on synthetic polymers is still not investigated. However,

since polyethylene, polypropylene, polystyrene and polyethylene terephthalate are the most

widely disposed synthetic polymers globally (Barnes et al., 2009), it can be supposed that our

results are biased due to the high accumulation of these specific synthetic polymers in our

oceans.

Potentially pathogenic V. parahaemolyticus as well as V. fluvialis occurred in water as well as

on microplastic particles. Recent studies report that V. parahaemolyticus and V. alginolyticus

are prevailing inhabitants of North Sea waters (Böer et al., 2013; Oberbeckmann et al., 2011b).

In contrast, V. vulnificus and V. cholerae are more abundant in the Baltic Sea (Böer et al., 2012),

which is also reflected by our findings. As already shown elsewhere, free-living bacterial

communities in general differ significantly from plastic-attached ones (Amaral-Zettler et al.,

2015; De Tender et al., 2015; Oberbeckmann et al., 2014; Zettler et al., 2013), which holds also

for microplastics investigated here. With respect to potentially pathogenic Vibrio spp., the

species V. vulnificus and V. cholerae were only isolated from seawater samples but not

identified on microplastics in the framework of our study. In contrast, V. parahaemolyticus was

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detected in both, water and on microplastic particles (Fig. 2). Additionally, V. parahaemolyticus

was detected once in the Baltic Sea and only on a microplastic particle throughout the entire

cruise.

Plastic is a persistent material and may serve as a reservoir and vector for potentially pathogenic

microorganisms. The drift of potentially harmful algae species, barnacles and bryozoans on

plastic debris (Barnes, 2002; Masó et al., 2003) is already well documented. Our results fuel

the evidence for potentially pathogenic bacteria being dispersed on microplastic particles by

wind or currents. However, although we identified V. parahaemolyticus on microplastics to

species level, due to the high intra-species diversity information on the geographical origin of

these hitchhikers or the microplastics is not possible, since the assignment of Vibrio species

down to specific ecotypes was not successful.

Vibrio spp. on microplastics were detected mainly close to the coast and only occasionally

offshore. However, microplastics and seawater samples carrying V. parahaemolyticus were

located exclusively in estuarine and coastal areas of the North and Baltic Sea. V.

parahaemolyticus occurrences in seawater were already addressed in several studies in

Northern European waters (Bauer et al., 2006; Böer et al., 2013; Collin and Rehnstam-Holm,

2011; Ellingsen et al., 2008; Lhafi and Kühne, 2007; Oberbeckmann et al., 2011b; Schets et al.,

2010) (Schets et al., 2011). Environmental parameters, such as temperature, salinity or plankton

abundance have an effect on Vibrio spp. communities and abundances (Blackwell and Oliver,

2008; Caburlotto et al., 2010; Drake et al., 2007; Martinez-Urtaza et al., 2008; Thompson et al.,

2004b; Turner et al., 2009; Vezzulli et al., 2009). Vezzulli et al. (2010) and Schets et al. (2010)

identified seawater temperature as a key factor influencing the presence of Vibrio spp., for

instance it is well documented that V. parahaemolyticus favours warmer water temperatures

(Sobrinho et al., 2010). Recently, pathogenic V. parahaemolyticus was detected even in

temperate European waters (Baker-Austin et al., 2010; Martinez-Urtaza et al., 2005). Martinez-

Urtaza et al. (2008) observed higher occurrence of this taxon during periods of lower salinity

and in general this taxon was primarily detected in areas of reduced salinity close to freshwater

discharge runoff, which is also in agreement with our findings.

In our study V. parahaemolyticus occurred also on microplastics collected from the drift line at

Helgoland. Oberbeckmann et al. (2011b) detected V. parahaemolyticus during summer months

and reported that the abundance of Vibrio spp. was influenced by specific environmental

conditions like the decrease in salinity due to an inflow of coastal water at Helgoland Roads

(North Sea, Germany). Each Vibrio group was influenced by different combinations of

environmental parameters but no single environmental parameter could explain the whole

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community structure of V. alginolyticus and V. parahaemolyticus populations in the German

Bight (Oberbeckmann et al., 2011b). The authors also reported that free-living and plankton-

attached Vibrio spp. abundances were mainly driven by the same environmental parameters

(Oberbeckmann et al., 2011b). This suggests that the potentially pathogenic V.

parahaemolyticus detected both on North Sea microplastics and in seawater samples of one

station were influenced equally by environmental conditions.

Conclusion

This study successfully evidences the occurrence of potentially pathogenic Vibrio spp. on the

species level on marine microplastics by use of MALDI-TOF MS for the first time. In most of

the cases, these species co-occurred also in surrounding seawater, suggesting that seawater

serves as a possible source for Vibrio colonization on microplastics. The fact that we for the

first time detected V. parahaemolyticus exclusively on polyethylene, polypropylene and

polystyrene particles, points to the urgent need to further address the biogeography and

persistence of these hitchhikers on marine microplastics. Studies on the co-occurrence of

specific V. parahaemolyticus genotypes on microplastic and surface water from the North Sea

are particularly important specifically with reference to the potential health impacts of

microplastic-colonizing microbial assemblages.

Acknowledgments

We would like to thank the team of the RV Heincke (AWI) for technical support. The authors

thank also for the modified maps provided by Dr. Mirco Scharfe (AWI, Helgoland).

This work was partially founded by the Alfred Wegener Institute for Polar and Marine

Research. We thank for the support of MikrOMIK a SWL/PAKT project by the Leibniz

Association (SAW-2014-IOW-2). We thank three anonymous reviewers for their helpful

comments.

GENERAL DISCUSSION

81

GENERAL DISCUSSION

After recognizing the potential threat posed by plastic pollution to humans and nature, scientists

started to study the ecological impacts of plastics and the Plastisphere in various habitats (e.g.

soil, fresh water and marine environments). Originally, the term “Plastisphere” was reffered to

a diverse microbial community of heterotrophs, autotrophs, predators, and symbionts which

was detected on diverse plasic samples (Zettler et al., 2013). In this thesis, the term Plastisphere

is reffering to the on plastics formed biofilm habitat with all its emergent properties. So far,

most studies adressing the Plastisphere focussed on randomly sampled plastics, and substrate

specificity had not been specifically addressed. Moreover, despite a growing number of

investigations on Plastisphere communities, most studies conducted so far focussed on bacterial

communities, neglecting the specificity of eukaryotic community associated with marine

plastics. Also the lack of knowledge about plastic surfaces as a potential site for the

accumulation of pathogenic microorganisms was highlighted by the scientific community

(Keswani et al., 2016; Osborn and Stojkovic, 2014). This thesis aimed at filling these three

knowledge gaps, and increases our understanding of the diversity and interactions within the

Plastisphere. The following sections discuss in a general context how the outputs of Chapters

I, II, and III contribute in answering those knowledge gaps and how the surrounding

environment, age of the biofilm, and the substrate specificity may determine the composition

of the Plastisphere. Furthermore, the role of plastic as accumulation site for pathogens is

discussed. Finally, new research questions emerging from this PhD work and avenues for future

studies are highlighted.

The Plastisphere, a unique microbial habitat

Zettler et al. (2013) for the first time coined out the term “Plastisphere”, referring to microbial

communities colonizing plastic substrates. This definition was based on differences in the

composition of microbial communities present on diverse, randomly collected, floating plastic

particles in contrast to their surrounding seawater communities. It is well documented, that

marine microbes mostly appear to prefer either a free-living or a surface-associated lifestyle,

although some species may switch their preference under certain environmental conditions or

life stages (Dang and Lovell, 2016; DeLong et al., 1993; Salta et al., 2013). However, several

subsequent studies confirmed the distinctness of plastic-associated microbial communities

compared to their planktonic counterparts (Amaral-Zettler et al., 2015; Bryant et al., 2016; De

Tender et al., 2017; De Tender et al., 2015; Oberbeckmann et al., 2014; Oberbeckmann et al.,

GENERAL DISCUSSION

82

2016). These findings are also supported by the outcome of this thesis indicating that, despite

possessing classes in common, biofilm and seawater communities are generally distinct

(Chapter I). Consequently, the results of this thesis (Chapter I) together with other studies,

clearly point towards the consensus that free-living seawater communities are different from

plastic-attached ones.

Within this thesis, Plastisphere communities were compared to biofilm communities attached

to glass. Furthermore, a thorough analysis of substrate specificity of microbial communities on

nine chemically distinct plastic types was carried out (Chapter I & II). The insights gained,

comparing Plastisphere and glass communities allow to conclude that, in marine environments,

the microbial core community of the Plastisphere is rather general than specific and, that

specificities for particular plastic types are rather related to the rare biosphere. Furthermore, the

composition of the Plastisphere also results from various interactions (1) between marine

biofilms and the surrounding environment, (2) in different age and between diverse organisms

within the biofilm, and (3) between marine biofilms and the substrate. These interactions are

discussed in more detail in the following sections. Up until now, the substrate specificity of

microbial communities present on chemically distinct plastic types was under debate, as many

studies conducted so far lacked in systematic and statistically robust analysis of distinct plastic

types. Numerous former studies focussed on the comparison of randomly collected diverse

marine plastics of unknown exposure time and origin (Amaral-Zettler et al., 2015; De Tender

et al., 2015; Oberbeckmann et al., 2014; Zettler et al., 2013). Random sampling of plastic-

attached communities impede a proper evaluation of substrate specificity due to unknown

exposure and biofilm realities such as environmental conditions (e.g. temperature, light,

salinity, and shear stress), physico-chemical properties of the substrate (e.g. hydrophobicity,

roughness, surface conditioning, nutrient enrichment) (Dang and Lovell, 2016) and the

differences in biofilm age.

The Plastisphere and the environment

Within a long term exposure experiment (15 month) the biofilm communities studied in

Chapter I were exposed to natural variation of several environmental factors in the North Sea

such as temperature or nutrient variation (Fig 1). The experiment was carried out using a natural

seawater flow-through system at the very well documented Long Term Ecological Research

(LTER) site “Helgoland Roads”. Hence, the microbial seawater community from Helgoland

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Roads, which is representative for the community passing through the flow-through system, is

proven to be recurrent (Chafee et al., 2017; Lucas et al., 2015; Teeling et al., 2012).

Fig 1 Environmental parameters (monthly means) recorded from 01. August 2013 – 30. November 2014 at

Helgoland Roads. T: temperature, S: salinity, Chl a: chlorophyll a.

The Plastisphere can be primarly considered as a general marine biofilm. Since it is well

established that the composition of biofilm communities is strongly driven by environmental

factors (Salta et al., 2013), the survival and successful growth of potentially plastic-specific

microorganism is likely also favoured by specific environmental conditions. For instance,

Amaral-Zettler et al. (2015) found that Plastisphere communities of the Atlantic and Pacific

Ocean clustered to a great extent by geographic location. This finding is in accordance with the

studies of Oberbeckmann et al. (2014); (2016), they showed that Plastisphere communities in

marine habitats are primarily driven by spatial and seasonal effects.

Abiotic conditions can also influence the abundance of individual species within biofilms.

Chapter III highlights differences in the geographic distribution of potentially pathogenic Vibrio

spp. on randomly collected, floating microplastics. Vibrio parahaemolyticus was, with one

exception, exclusively detected on microplastics in coastal and estuarine regions of the North

Sea. Vibrio parahaemolyticus are known as prevailing inhabitants of North Sea waters (Böer et

al., 2013; Oberbeckmann et al., 2011b). Oberbeckmann et al. (2011b) detected V.

parahaemolyticus during summer months and reported that their abundance was influenced by

a combination of specific environmental conditions while no single environmental parameter

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could explain the overall community structure of V. parahaemolyticus populations in the

German Bight (Oberbeckmann et al., 2011b). The authors also reported that free-living and

plankton-attached Vibrio spp. abundances were mainly driven by the same environmental

parameters (Oberbeckmann et al., 2011b). This suggests that the abundance of potentially

pathogenic V. parahaemolyticus detected both on North Sea microplastics and in seawater

samples of one station may have been similarly influenced by environmental conditions.

The conditions present in the natural seawater flow-through system used in Chapter I and II,

with, less shear forces and no light, may have influenced Plastisphere composition. De Tender

and colleagues (2017) carried out a one year exposure experiment of PE in two different

environments, harbour and offshore, in the North Sea. Interestingly, they detected a shift

towards more secondary colonizers of PE biofilms at later stages only in the harbour

environment, which is less exposed to shear and current forces. Furthermore, they observed that

plastic samples taken offshore, either with known history or randomly sampled, were most

similar to early phase biofilms observed on plastics incubated in the harbour (De Tender et al.,

2017). Four genera detected by De Tender et al. (2017) in the harbour were also abundant in

the mature biofilm communities studied in Chapter I. This suggests that the Plastisphere, which

developed during the experiment, may not represent a community from another season or from

open waters.

The survival and successful growth of potentially plastic-specific microorganisms is likely

driven by environmental conditions. Optimally, to delineate the effects of season, habitat

variation, and substrate specificity on community composition, Plastisphere communities

should be monitored at close time intervals over more than one seasonal cycle, and at different

locations.

The Plastisphere: Does age make a difference?

Within this PhD project, long-term experiments were conducted, in which nine different plastic

substrates were incubated under the same conditions over a period of 15 months (Chapter I)

and 21 months (Chapter II). To the best of of my knowledge, there exists only one other long-

term study which monitored the development of the Plastisphere. However, this one-year long

experiment only used one type of plastic as substrate (PE) of two different colour and surface

properties (dolly rope and sheet) (De Tender et al., 2017). Therefore, Chapter I and II of this

PhD thesis present unique data from experiments rarely performed in this subject area.

Moreover, the majority of studies conducted so far, investigated the Plastisphere on floating

particles of unknown age, or, alternatively, over a short incubation time (days to weeks). None

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of these studies detected significant differences between distinct plastic types or between

plastics and other inert substrates (Kettner et al., 2017; Oberbeckmann et al., 2018;

Oberbeckmann et al., 2016). The discrepancy between these observations and the differences

between 15 months old Plastisphere communities, detected in Chapter I, is surprising since it

has been demonstrated that bacterial communities present on dissimilar surfaces evolve to a

similar community structure over time (De Tender et al., 2017; Jones et al., 2007; Salta et al.,

2013). Moreover, the influence of the substrate type should decrease over time since the

accumulation of biota in a dense mature biofilm covers up the substrate surface (Jones et al.,

2007). Sensing of a non-soluble surface followed by successful colonization are the first two

steps for marine bacteria in biofilm formation (Dang and Lovell, 2016; Sivan, 2011). One

possible explanation for the similarities of “young” Plastisphere communities might be that

plastics, as any other surface in the marine environment, become conditioned or coated by

organic polymers, which generates a chemical modification (Bhosle et al., 2005) potentially

masking the physico-chemical surface properties of diverse plastic types. This effect has been

previously suggested as an explanation for the fact that young marine biofilms are

indistinguishable within the first four days of development on stainless steel and polycarbonate

surfaces (Jones et al., 2007). However, Dang and Lovell (2016) stated that surface properties,

and resulting chemodynamics like surface conditioning or nutrient enrichment, may play a role

in forming distinct biofilm communities. Bravo et al. (2011) observed fewer taxa on plastic jar

surfaces than on Styrofoam pieces in early stage biofilm formation, and hypothesized that

substrate surface rugosity may facilitate the initial colonization of marine plastics. Also, De

Tender et al. (2017) observed the development of different microbial communities on two types

of PE (plastic sheets and dolly ropes), with slightly higher bacterial diversity on dolly ropes

within the first few weeks of exposure to Belgian North Sea waters.

As already mentioned, the Plastisphere can be primarly considered as a general marine biofilm.

With increasing age, a natural biofilm becomes increasingly complex in terms of taxonomic

diversity and architecture. The Plastisphere is, metaphorically speaking, a multicultural city of

marine microbes (marine biofilm) including ethnical majority and minority groups (abundant

and rare taxa), all with very different abilities, built on an artificial substrate (plastics). Biofilm

development on artificial substrates follows a general pattern (Artham et al., 2009; Bravo et al.,

2011; Lobelle and Cunliffe, 2011), starting with the adsorption of dissolved organic molecules,

followed by the attachment of bacterial cells, and by the attachment of unicellular eukaryotes,

concluded by the attachment of larvae and spores (Dobretsov, 2010). Even though a growing

number of studies focus on the comparison of Plastisphere communities present on distinct

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plastic types, only two studies describe their complete prokaryotic and eukaryotic community

composition (Bryant et al., 2016; Oberbeckmann et al., 2016). De Tender et al. (2017)

investigated additionally to the bacterial also fungal Plastisphere communities of PE in parallel,

using 16S rDNA and ITS2 metabarcoding. These three studies observed a high variability in

eukaryotic or fungal community composition, which is consistent with the general

heterogeneity of eukaryotic communities presented in Chapter I. However, the interplay of

diverse groups of organisms within the Plastisphere is highly complex and far from being

understood. Cell to cell interactions, such as competition and cooperation, are likely to have an

effect on the biofilm community structure (Flemming et al., 2016; McLean et al., 2005).

Chesson and Kuang (2008) suggested that competition dynamics at lower trophic levels

(bacteria and microflagellates) may have consequences for protists’ dynamics. The bacterial

layer might attract different eukaryotic predators feeding on specific bacterial groups, which

may, in turn, control through top-down forces active bacterial populations in a mature biofilm

(Andersson et al., 2010).

In Chapter I, I investigated complete Plastisphere communities at one time point. Future studies

should monitor the Plastisphere development at close time intervals during the initial phase to

understand the influence of different plastic types and other substrates on initial colonization,

and subsequently take monthly samples over an annual cycle to identify the impact of the

Plastisphere age on substrate specificity. This will be necessary to indentify the timing at which

specific species or assemblages appear or disappear during biofilm development and aging.

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The Plastisphere and its substrate specificity

Within this PhD project, Plastisphere communities that colonized nine distinct plastic foils, as

well as glass as neutral control, were analysed to assess substrate specificity. The term substrate

refers here, to a surface on which an organism grows or is attached and which might serve as a

carbon source. The knowledge gained in Chapter I and II can be divided in the following major

outcomes regarding the substrate specificity of the Plastisphere. 1. Prokaryotic Plastisphere

communities were different from glass communities, and significant differences were detected

between various plastic types (Chapter I). 2. A general marine prokaryotic biofilm community

serves as shared core on all plastics and plastic “specific” microbes/assemblages rather account

to the rare biosphere (Chapter I & II). 3. The term Plastisphere is valid for prokaryotic but may

not be valid for eukaryotic biofilm communities since the communities appear generally

heterogenic (Chapter I).

1. So far, little is known on the specificity of marine biofilms, since only a few studies in marine

environments investigated the consequences of exposure to diverse plastic substrate properties

on the taxonomic composition of the Plastisphere community by comparing distinct plastic

types to other substrates incubated under similar conditions. Ogonowski et al. (2018) incubated

cellulose, glass, PE, PP and PS, using natural sediments as source community for the different

substrate types, and found significant differences between plastic and non‐plastic colonizing

microbial communities. However, the specificity of these communities on their respective

chemically distinct plastic types remains unclear. Comparing the PET and glass associated

microbiomes, Oberbeckmann et al. (2016) did not detect significant differences in the

prokaryotic community composition. Contrariwise, in Chapter I of this thesis, the biofilms

associated to PET were significantly different from those associated to glass. A possible

explanation for the contradictory results of these two studies might be the differences in biofilm

age and/or environmental conditions applied. Recently, Oberbeckmann et al. (2018)

investigated the Plastisphere communities by comparing HDPE and PS with wood, and found

the plastic-associated communities to be different from those associated to wood. Moreover,

studies comparing HDPE and PS communities found no differences (Oberbeckmann et al.,

2018), which is consistent with the findings of this study (Chapter I, Fig 1).

2. Significant differences in the composition of biofilm communities associated to diverse

plastics and glass were found and described in Chapter I. It is important to note that these were

generally low, indicating that the shared core of the various biofilms is a rather substrate

unspecific one. Differences in sequencing depth could explain why no other study could detect

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significant differences in the composition of biofilms covering diverse plastic types. This

highlights the difficulties of comparing the outcomes from different culture-independent

sequencing-based methodological approaches. The strongest contribution to the total

dissimilarity between the diverse substrates was often given by less abundant OTUs (Chapter

I). However, these observations indicate that the rather rare species within the Plastisphere,

which interact with the general biofilm community, are the species with high substrate

specificity. In general, microbial communities consist of a few abundant taxa while a large

proportion of rare taxa makes up the so called “rare biosphere” (Pedrós-Alió, 2006). To account

for the rare species within the Plastisphere, OTUs with a mean relative abundance of <0.1% in

the 15 months old biofilms associated to diverse plastics were additionally investigated in the

context of Chapter I (Fig 2). Principle coordinates and PERMANOVA analyses of the rare

Plastisphere give a similar impression to the abundant Plastisphere communities (>0.1%;

Chapter I), in the light of glass communities being distinct from all Plastisphere communities

(Fig 2, PERMANOVA = p (perm) < 0.05). Following to glass, PLA was shown to harbour the

most distinct rare Plastisphere community, as the PLA community was significantly different

from three other plastic types (Fig 2, PERMANOVA = p (perm) < 0.05).

Fig 2 Rare biospheres. OTUs with a mean relative abundance of <0.1% (n=50) were analysed. (a) Principle

Coordinate Ordination relating variation in rare prokaryotic community composition between different synthetic

polymers and glass biofilm. PCOs representing similarity of biofilm communities based on relative abundances of

OTUs across samples. (b) PERMANOVA & PERMDISP pair-wise tests of rare prokaryotic biofilm

communities on different plastics and glass based on Hellinger distance of operational taxonomic units (OTUs).

Significant results (p (perm) < 0.05) are highlighted in blue (PERMANOVA) and yellow (PERMDISP), green

indicates significant results in both tests.

Based on the knowledge gained in Chapter I, and considering that the predation and competition

pressure in mature biofilms can be particularly high (e.g. for space or nutrients) (Andersson et

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al., 2010), in Chapter II tightly surface-attached were uncovered in order to investigate potential

“rare plastic specific” genera. Combining the datasets of Chapter I and Chapter II revealed that

70% of the uncovered potential plastic “specific” OTUs were assigned to the rare biosphere

(<0.1%) of the Plastisphere communities investigated in Chapter I. It remains unclear whether

the rare biosphere is representing an active part of the microbial community, and if so, which

role it plays in community dynamics and ecosystem functioning (Wilhelm et al., 2014). On the

one hand, rare phylotypes were previously reported to tend to stay rare (Galand et al., 2009;

Kirchman et al., 2010). On the other hand, Besemer et al. (2012) demonstrated that at least a

certain portion of rare OTUs is active, indicating that those have the potential to increase in

abundance, under favourable environmental conditions (Andersson et al., 2010). Wilhelm et al.

(2014) found a large proportion of rare taxa with higher relative abundances in rRNA compared

with rDNA, suggesting that the rare biosphere contributes disproportionately to microbial

community dynamics. Having the potential to increase in their abundance, our findings clearly

support the idea that potential plastic “specific” species are, at least partly, controlled by

competitive interactions in mature dense biofilms (Chapter II).

3. The only study addressing the eukaryotic community composition of the Plastisphere in an

exposure experiment, found no significant differences between glass and PET biofilms

(Oberbeckmann et al., 2016). While significant differences detected between PET, as well as

other plastic types, and glass, statistical tests of dispersion revealed that these differences were

most likely the result of within-system heterogeneities (PERMDISP, Chapter I). The eukaryotic

communities of all tested substrates appeared generally heterogeneous (within group

dispersion) which is in accordance with the findings of Oberbeckmann et al. (2016). As part of

the eukaryotic community, fungi are often “the forgotten ones” in microbial ecology studies.

To account for the fungal Plastisphere, the fungal community composition of the biofilms

associated to diverse plastics was also investigated in the context of Chapter I (Fig 3). Principle

coordinates and PERMANOVA analyses indicate that the fungal Plastisphere communities are

overall highly heterogeneous (Fig 3). Despite this heterogeneity, PP and glass fungal

communities were found to be significantly different (Fig 1, p (perm) < 0.05).

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Fig 3 Fungal communities. OTUs with a mean relative abundance of at least 0.1% in one substrate type (n = 5)

were analysed. (a) Principle Coordinate Ordination relating variation in fungal community composition

between different synthetic polymers and glass biofilm. PCOs representing similarity of biofilm communities

based on relative abundances of OTUs across samples. (b) PERMANOVA & PERMDISP pair-wise tests of

fungal biofilm communities on different plastics and glass based on Hellinger distance of operational taxonomic

units (OTUs). Significant results (p (perm) < 0.05) are highlighted in blue (PERMANOVA) and yellow

(PERMDISP).

Recently, Plastisphere communities were shown to differ from wood associated communities,

but, no significant differences were detected comparing fungal Plastisphere HDPE and PS

communities (Kettner et al., 2017), which is consistent with my findings (Fig 3). These results

indicate that fungi in the Plastisphere are generally more heterogeneous (PERMANOVA = p

(perm) > 0.05), and that preferences for a particular plastic type may not be detected because

of their “random” growth. However, since fungi are of particular interest in their function as

potential plastic degraders in the environment (Grossart and Rojas-Jimenez, 2016; Krueger et

al., 2015), the role as part of the Plastiphere and their impact on plastic as a substrate in marine

environments needs further investigations.

The scientific community of the Plastisphere intensively discussed the possibility of plastic

“specific” organisms/assemblages to be potentially involved in biodegradation (Amaral-Zettler

et al., 2015; Bryant et al., 2016; De Tender et al., 2017; De Tender et al., 2015; Oberbeckmann

et al., 2018; Oberbeckmann et al., 2014; Oberbeckmann et al., 2016; Zettler et al., 2013).

Concluding my findings, there are strong indications that plastic specific

organisms/assemblages exist in the marine environment, but that their development is

controlled or even suppressed by natural conditions and interactions/competition with other

organisms, which impede the establishment of a “truly” plastic specific community.

Considering the enormous reservoir of genetic diversity of the “rare Plastisphere” with the

general potential of several microbes to degrade complex organic compounds, I believe that

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their importance, e.g. their potential degrading ability, within the Plastisphere is

underestimated. Therefore, future studies of the Plastisphere should not only focus on

taxonomic composition and most abundant species but also assess the rRNA expression as

indicator for the active Plastisphere community. Future studies should also screen for specific

enzymes of given microbial strains which may enable them to use plastics as their main carbon

source (Pathak and Navneet, 2017; Yoshida et al., 2016).

The Plastisphere as potential vector and accumulation site for pathogens

Most plastic types are poorly degradable and are, as any other surface, rapidly colonized by

microorganisms. When entering the oceans, plastics could be consequently transported over

long distances in marine environments, as compared to naturally occurring polymers, and

therefore function as a vector for the dispersal of harmful or even human pathogenic species.

Within this PhD project, specific attention has been paid to identify potentially human

pathogenic Vibrio spp. on floating microplastics in the North and Baltic Sea (Chapter III). The

first indication of the presence of Vibrio spp. on marine plastics was published by Zettler et al.

(2013), who reported high abundances of this genus with up to 24 % of the whole Plastisphere

community. Later on, De Tender et al. (2015) detected members of the Vibrionaceae family on

marine plastics from the Belgian North Sea. However, due to short read lengths, a conclusive

identification on the species level was not provided so far (De Tender et al., 2015; Zettler et al.,

2013). The outcome of Chapter III highlight for the first time the presence of cultivable Vibrio

spp. on marine microplastic particles, including potentially pathogenic V. parahaemolyticus

strains. As they are persistent materials, plastics may not only serve as a vector for the dispersal

but also as an accumulation site of pathogenic species. Jambeck et al. (2015) estimated that 4.8

to 12.7 million MT of mismanaged plastic waste entered the oceans in 2010. Considering the

yearly growing amount of mismanaged plastic litter entering and accumulating in the oceans

and the, mainly due to fragmentation resulting various size fractions, the accumulation and

transport of pathogens and alien species may have consequences for various ecosystems, for

different trophic levels of the food web, as well as for human and animal health. For instance,

Schmidt et al. (2014) demonstrated with the use of oligotyping that Vibrio communities present

on plastic substrates include several species potentially pathogenic for fish, corals, and bivalves.

Recently, Viršek et al. (2017) identified the fish pathogen Aeromonas salmonicida on

microplastics of the North Adriatic Sea, and suggested that microplastics serve as a vector for

this harmful invasive species. Beside bacteria, plastic may also serve as vector for harmful

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eukaryotes as the drift of potentially harmful algae species, barnacles and bryozoans on plastic

litter has been already reported (Barnes, 2002) (Masó et al., 2003).

Research has just started to unravel ecological implications of pathogens and alien species of

the Plastisphere. The fact that V. parahaemolyticus was identified, exclusively on randomly

collected PE, PP and PS particles, highlights the urgent need to further address the 1.

biogeography, 2. persistence, 3. substrate specificity, and 4. co-occurrence of specific

genotypes on microplastic and surface water of these hitchhikers.

Final conclusion

The main purpose of this thesis was the detailed description of the Plastisphere associated to

various plastic types. The combination of the high sample replication with usage of culture-

independent high-resolution techniques, like 16S rRNA tag sequencing and visual tools (SEM)

for the description of the prokaryotic and eukaryotic Plastisphere, allowed for sensitive and

statistically robust observations of Plastisphere communities, and to analyse substrate

dependent specificities. The combination of selective enrichment and isolation, MALDI-TOF

MS, and PCRs of regulatory and virulence-related genes in the culture-dependent approach

enabled a conclusive identification on the species level of potential Plastisphere pathogens. At

the onset of this PhD project, 2014, Osborn and Stojkovic (2014) reviewed the knowledge

regarding “Marine Microbes in the Plastic Age” and formulated key questions that need to be

answered in order to understand the diversity and ecology of the Plastisphere. The outcome of

this thesis provides answers to two of these questions;

“Do plastic surfaces select specifically for particular microbial species and/or alternatively,

are plastic surfaces just primarily a convenient substrate for colonisation?”

Originally, the term “Plastisphere” was reffered to a diverse microbial community, detected on

diverse plastic samples and which was found to be distinct from the surrounding seawater

communities. Considering these two habitats, plastic surfaces select for particular microbial

species, since the Plastisphere can be primarly considered as a general marine biofilm.

Compairing the communities associated to diverse substrates, unambiguously, plastic surfaces

are primarily a convenient substrate for colonisation since the microbial community of plastics

and glass in the marine environment is a more general than a specific one. However, the

outcome of this thesis indicate also that plastic surfaces select specifically for particular

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microbial species but that these specificities for a distinct plastic type are related to the rather

rare biosphere and might be controlled by top-down forces, competition pressure, and

environmental conditions.

“Are plastic surfaces a potential site for accumulation of pathogenic microorganisms?”

Plastic surfaces serve as a potential site for accumulation of pathogenic microorganisms and

plastic might therefore serve as potential vector for their distribution. Within this PhD project,

the presence of cultivable Vibrio spp. was conclusively confirmed on 13 % of collected marine

microplastic particles, including potentially pathogenic V. parahaemolyticus strains detected on

12 microplastic particles.

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95

FUTURE PERSPECTIVES

Overall, this thesis provides a complete overview of the Plastisphere eukaryotic and prokaryotic

community composition, their specificities to diverse plastic types and glass, and the

Plastispheres role as reservoir for potential pathogenic bacteria in marine environments.

Chapters I to III comprise comprehensive, statistically robust, and descriptive approaches which

provide an in-depth picture and solid base for future research on the Plastisphere.

Chapter I and II focused on the substrate specificity of Plastisphere communities associated

with distinct plastic types and glass. The knowledge gained during these studies indicates that

plastic “specific” microorganisms/assemblages account to the rather rare biosphere, likely

because of slow growth of respective organisms, or the biomass of these organisms are

controlled by environmental and biotic (e.g., competition, grazing) pressures. Subsequently, in

the marine environment their biomass might be too low to have a potential role in the biological

degradation of plastics over ecological relevant time scales. Due to their longevity, plastic items

entering in marine environments, accumulate and become fragmented into various sizes over

time. Consequently, (micro)-plastics are detected worldwide in various marine environments

(Cole et al., 2011; Eriksen et al., 2014) representing a major threat for marine life (Galgani,

2015; Gregory, 2009) and might have severe implications for human health and the

environment. Whether it is motivated by applied or fundamental research, one of the main goals

to study the Plastisphere nowadays is to identify plastic “specific” microorganisms/assemblages

which can degrade this highly complex substrate that pollutes the marine environment.

To prove biodegradation of plastic, one can use (1) a substrate based approach, in which the

plastic/product itself is analysed and/or (2) a biological approach, in which the organism or

assemblage is assessed.

(1) Commonly, in the substrate based approach, biological degradation of plastics is assessed

by growing organisms on medium enriched with a synthetic polymer as sole carbon source;

followed by gravimetrically determining the resulting mass loss and size reduction of the

polymer. The analysis of degradation products, e.g. the amount of produced metabolites, such

as CO2, can be assessed through biodegradation assays (Pathak and Navneet, 2017).

Additionally, changes of different functional groups of the respective polymer can be measured

to prove microbial degradation by Fourier Transform Infrared Spectroscopy (Harshvardhan and

Jha, 2013; Nowak et al., 2012). While the focus of this PhD was predominantly on the

specificities of Plastisphere communities, Scanning Electron Microscopy was used to observe

physical changes of the surface morphology of different plastic types, like embrittlement and

FUTURE PERSPECTIVES

96

micro-cracks (Arutchelvi et al., 2008), which could possibly result from degradation or

biofouling processes. SEM investigations of various plastic types were performed on pristine

plastics surfaces and after removal of the 21 months old biofilm in the context of Chapter II

(Fig 4). While, overall, most plastic surfaces appeared visually smooth and unaltered, signs of

alteration of different degrees on the surface of all visualized plastic types were observed. The

diverse surfaces seem to be partly deformed, in places embrittled and cracks or holes developed

over time (Fig 4). The largest changes in surface morphology were, by far, observed for PLA

(Fig 4). The pristine PLA foil has a flat, smooth surface with minimal imperfections. The PLA

foils after incubation showing countless pits of porous structure distributed all over the surface

(Fig 4). On the one hand, PLA is known to be

biodegradable when composted, it seems likely

that these erosions are caused by microbes of the

Plastisphere attacking first vulnerabilities in the

polymer structure, like additives or thinner

surface structures. On the other hand, the

degradation mechanism of PLA starts with

chemical hydrolysis in the presence of water at

elevated temperatures, followed by biological

degradation (Shah et al., 2008), hence biotic

degradation seem rather unlikely. Interestingly,

the rare and abundant Plastisphere community of

PLA was the most distinct compared to the other

tested plastic substrates (Chapter I). However,

sole visual inspection does not suffice to

conclude whether the porous structure of the aged

PLA is the result of biodegradation or whether

this structure was already present but hidden

under a thin polymeric layer and unravelled by

erosion of the surface over time.

Generally, the longevity of plastics in the marine

environment is a matter for debate, and estimates

range from hundreds to thousands of years

depending on the chemical and physical

properties of the synthetic polymer (Barnes et al.,

Fig 4 Aged plastic surfaces. Scanning Electron

microscopic images of selected pristine plastic

surfaces, and after removal of 21 months old

biofilms. Scale bar = 1 µm.

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97

2009). To gain knowledge on the influence of the Plastisphere on the longevity and alteration

of distinct plastic types in the marine environment, long-term incubation experiments of distinct

plastic types under comparable conditions, like the one conducted during this PhD project, are

needed. Regular assessment of the plastic substrates by the combination of visual (e.g. SEM)

and spectroscopic techniques (e.g. ATR FT-IR) will provide invaluable information on

structural changes of plastics over time.

(2) The above-mentioned visual techniques only provide indications on the

alteration/degradation of the plastics investigated, but hard evidence for biological degradation

is missing. In Chapter II of this thesis, tightly attached potentially plastic “specific” microbes

were uncovered. As successful colonisation of a plastic surface is no proof for biological

degradation, the degradation ability of an organism or assemblage needs to be additionally

addressed.

Biological degradation can be determined, for example, by assessing specific enzymatic activity

of a given microbial strain (Pathak and Navneet, 2017; Yoshida et al., 2016). Therefore, as a

first step, microbes need to be isolated and identified, which I did in the context of Chapter II.

I enriched and isolated bacteria and fungi from distinct plastic types and glass. Bacteria were

isolated from HDPE, PS, PET, SAN, PESTUR and glass, fungi from HDPE, PS, PESTUR and

glass. Bacterial isolates were de-replicated by MALDI-TOF MS, which allowed selection of

representative isolates per substrate prior to Sanger Sequencing (see detailed information on

enrichment, isolation, de-replication, DNA extraction and sequencing in the supplementary

information). The resulting 47 bacterial isolates were taxonomically classified to the genera

Thalassospira, Marinobacter, Pseudoalteromonas, Alteromonas, Muricaudap, Sporosarcina,

Jeotgalibacillus, Micrococcus, Sulfitobacter, Celeribacter and Bacillus (Fig S1). Strains of the

genus Bacillus and Micrococcus were previously reported to be associated with polymer

degradation (Pathak and Navneet, 2017). Pseudoalteromonas spp., which are known as

hydrocarbon degraders, are regularly detected as part of the Plastisphere (Oberbeckmann et al.,

2016; Zettler et al., 2013). Since fungi are of particular interest in their role as potential plastic

degraders in the environment (Grossart and Rojas-Jimenez, 2016; Krueger et al., 2015), 12

fungal strains were isolated, sequenced, and taxonomically assigned to the classes of

Tremellomycetes, Cystobasidiomycetes, Microbotryomycetes, Leotiomycetes,

Sordariomycetes, Eurotiomycetes and Exobasidiomycetes (Table S1).

However, at this stage it is impossible to state if the isolated strains actively degraded one of

the given plastic types, which needs to be addressed in future studies. Moreover, active

organisms that could not be isolated might play a specific role in interspecies interactions

FUTURE PERSPECTIVES

98

(cooperation) in plastic-degrading microbial assemblages. Most marine microorganisms are

viable but non-culturable (Eilers et al., 2000) which makes it particularly difficult, or even

impossible to isolate specific plastic degrading microbes/assemblages. Stable Isotope Probing

(SIP), the analysis of incorporated 13C in the DNA of the belonging metabolizer (Bernard et al.,

2007) can provide hard evidence for plastic degraders in a microbial community. The

substantial disadvantage of this technique is that 13C labeled plastics are either unavailable or

expensive. Nevertheless, this approach has great potential due to the advantage to asses a

microbial community, instead of a pure culture, and might be used in future research to address

biodegradation by Plastisphere communities.

SUMMARY

99

SUMMARY

Plastic litter is entering and accumulating in our oceans and can be found in the marine

environment all over the globe. When entering marine waters, plastics as any other surface, is

rapidly colonized by a plethora of organisms, which form dense biofilms on the plastic surface,

the so-called “Plastisphere”. Despite growing concerns about the ecological impact of plastics

on the marine environment during the last decade, the number of studies addressing

Plastisphere-related questions remains limited.

This thesis aimed to tackle this knowledge gap by comprehensively describing and analysing

specificities of Plastisphere communities attached to chemically distinct plastic types.

The specificity of mature Plastisphere communities was investigated on nine chemically

different plastic types, and compared to the inert control substrate glass. The Plastisphere

communities attached to diverse plastic types were found to be distinct from glass-associated

communities. A more general marine biofilm core community serves as shared core among all

tested plastic types and glass, rather than specific Plastisphere communities. The general

heterogeneity of eukaryotic communities was much higher, indicating that the term Plastisphere

is valid for mature prokaryotic biofilm communities, but may not be for eukaryotic ones.

This work also showed that the prokaryotic shared core of the various mature Plastisphere

communities are rather substrate unspecific, pointing towards the importance of rather rare

species in plastic associated marine biofilms. A high-pressure water Jet treatment technique

was developed to remove the cohesive layer of mature biofilms, while the adhesive layer

remains on the plastics surface . It was shown that tightly attached microorganisms might

account rather to the rare biosphere in mature Plastisphere communities, which suggests the

presence of plastic “specific” microorganisms/assemblages.

Due to their longevity, plastics could be transported over long distances in marine

environments, and therefore may function as a vector for the dispersal of pathogenic species.

To test this, plastic particles were collected in the North Sea and the Baltic Sea and screened

for the presence of pathogens. Potentially pathogenic Vibrio parahaemolyticus were discovered

on a number of microplastic particles, e.g. polyethylene, polypropylene and polystyrene.

Mostly, this species co-occurred also in surrounding seawater, suggesting that seawater serves

as a possible source for Vibrio colonization on microplastics. The confirmed occurrence of

potentially pathogenic bacteria on marine microplastics highlights the urgent need for detailed

biogeographical analyses of marine microplastics.

SUMMARY

100

The results from this thesis substantially increase our understanding of the diversity and

specificity of Plastisphere communities. This thesis comprises a detailed and descriptive

approach, which provides a fundamental knowledge basis for future research on Plastisphere

questions related to e.g. potential biodegradation of marine plastics and the vector function for

alien and potentially pathogenic species.

ZUSAMMENFASSUNG

101

ZUSAMMENFASSUNG

Plastikmüll gelangt über diverse Eintragungswege in unsere Meere und wird weltweit in allen

marinen Gewässern gefunden. Wie alle anderen Oberflächen, wird auch Plastik im Meerwasser

schnell von einer Vielzahl von Organismen besiedelt, die auf der Plastikoberfläche dichte

Biofilme bilden, die sogenannte „Plastisphere". Trotz der im letzten Jahrzehnt wachsenden

Besorgnis über die ökologischen Auswirkungen der Plastikvermüllung in den Meeren, ist die

Zahl der Studien, die sich mit speziellen Fragen wie die der „Plastisphere“ beschäftigen,

begrenzt. Daher sind deren ökologische Relevanz und die resultierenden Konsequenzen dieser

„Plastisphere“ noch weitgehend unverstanden. Um einen Teil dieser Wissenslücke zu schließen

liefert die vorliegende Arbeit eine umfassende Beschreibung und Analyse dieser „Plastisphere“

Gemeinschaften, besonders im Hinblick auf deren Spezifität auf verschiedenen chemisch

unterschiedlichen Kunststoffen.

Die Spezifität ausgereifter „Plastisphere“ Gemeinschaften wurde am Beispiel von neun

chemisch unterschiedlichen Kunststoffen untersucht und mit den Gemeinschaften auf dem

inerten Kontrollsubstrat Glas verglichen. Die „Plastisphere“ Gemeinschaften, assoziiert mit

diversen Kunststoffen, unterschieden sich von Glas Gemeinschaften. „Plastisphere“

Gemeinschaften erscheinen jedoch eher als generelle marine Biofilm Gemeinschaften mit einer

gemeinsamen Kerngemeinschaft aller getesteten Kunststoffe aber auch von Glas.

Eukaryotische Gemeinschaften waren generell viel heterogener, sowohl im Vergleich diverser

Substrate zueninander als auch innerhalb der jeweiligen Substrat Replikate. Dies deutet darauf

hin, dass der Begriff „Plastisphere“ für ausgereifte prokaryotische Biofilme zutreffend ist, aber

nicht für eukaryotische Biofilm Gemeinschaften.

Da die ausgereiften prokaryotischen Kerngemeinschaften der Plastisphere eher unspezifisch

sind, fokussiert diese Arbeit weitergehend auf eher seltener Arten in den „Plastisphere“

Gemeinschaften. Es wurde eine Hochdruck-Wasser-Jet Behandlungstechnik entwickelt, um die

kohäsive Schicht ausgereifter Biofilme zu entfernen, während die adhäsive Schicht auf der

Kunststoffoberfläche verbleibt. Stark assoziierte Mikroorganismen zählten zu der eher seltenen

Biosphäre in den ausgereiften Plastisphere Gemeinschaften, was einen Hinweis darauf liefert

das "spezifische" Mikroorganismen oder Consortia auf unterschiedlichen Plastik Substraten

nicht abundant aber dennoch vorhanden sind.

Plastik kann aufgrund der langen Lebensdauer in marinen Umgebungen über weite

Entfernungen transportiert werden und kann daher als Vektor für die Verbreitung pathogener

Arten fungieren. Um dies zu testen, wurden Mikroplastikpartikel in der Nord- und Ostsee

ZUSAMMENFASSUNG

102

gesammelt und auf potentiell pathogene Bakterien untersucht. Die potentiell pathogene Art

Vibrio parahaemolyticus wurde auf einer Reihe von Mikroplastikpartikeln entdeckt, unter

anderem auf Polyethylen, Polypropylen und Polystyrol. Meistens trat diese Arten auch im

umgebenden Seewasser auf, was darauf schließen lässt, dass Seewasser allgemein als mögliche

Quelle für die Besiedlung durch die Gattung Vibrio auf Mikroplastik dient. Der Nachweis

potenziell pathogener Bakterien auf marinem Mikroplastik unterstreicht den dringenden Bedarf

an detaillierten biogeographischen Analysen mariner Mikroplastikpartikel.

Die Ergebnisse dieser Arbeit verbessern sowohl unser Verständnis über die Vielfalt

eukaryotischer und prokaryotischer „Plastisphere“ Gemeinschaften, als auch deren Spezifität

zu verschiedenen Kunstoffen und anderen inerten Materialien erheblich. Desweiteren, umfasst

diese Arbeit einen detaillierten und deskriptiven Ansatz, der eine grundlegende Wissensbasis

für zukünftige Studien zur „Plastisphere“ bietet. Themen wie z.B. den potentiellen biologischen

Abbau von marinem Plastik, oder die Rolle als Vektor für nichtheimische und potenziell

pathogene Arten, könnten dabei im Focus stehen.

Supplement

The Supplement contains four subsections, one for each of the Chapters I to III. One subsection

contains information of the methods used and documentation of the preliminary results of the

section “Future Perspectives”.

Supplementary material for Chapter I

Mature biofilm communities on synthetic polymers in seawater -

Specific or general?

Four figures illustrating the experimental design, environmental conditions, abundance profiles

of eukaryotic kingdoms and phyla, and the most abundant, characteristic and discriminative

prokaryotic and eukaryotic OTUs. Further nine tables giving detailed information about

synthetic polymers, PERMANOVA and PERMDISP tests, SIMPER analysis, taxonomic path

of OTUs with a mean relative abundance of at least 0.1% in at least one sample including tested

similarities within prokaryotic and eukaryotic communities, and prokaryotic classes detected in

biofilms compared to Helgoland Roads communities.

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108

Fig S1 Flow-through incubation system for foil-samples of different synthetic polymers mounted in

conventional slide-frames. (a) Mounting frames with different polymer foils in the seawater system, (b)

appearance of the polymer foils at the start in August 2013, (c) appearance of the polymer foils in September 2014.

(d) Environmental parameters (monthly means) recorded from 01. August 2013 – 30. November 2014 at

Helgoland Roads. T: temperature, S: salinity, Chl a: chlorophyll a.

SUPPLEMENT CHAPTER I

109

Fig S2 Eukaryotic biofilm community composition on different synthetic polymers and glass. Abundance

profiles of eukaryotic (a) kingdoms and (b) phyla on different synthetic polymers and glass. OTUs with a mean

relative abundance of at least 0.1% in one substrate type (n = 5) were analysed. A * indicates the term

“unclassified”, `indicate the term “Incertae sedis”.

SUPPLEMENT CHAPTER I

110

Fig S3 Most abundant and discriminative prokaryotic OTUs of the nine different synthetic polymers and

glass (n=5). OTUs with a mean relative abundance of at least 0.1% (n=5) in at least one substrate type were

analysed. Displayed are OTUs with a mean relative abundance of at least 1% or jointly contributing, with a

minimum of 1%, to the total dissimilarity between different statistically significant (PERMANOVA p<0.05) glass

and synthetic polymer groups. Groups showing both, PERMANOVA and PERMDISP significant p values were

rejected. The amount of contribution is indicated by the colour of cells, darker colours represent higher

contributions. Bold lines indicate OTUs contributing to the same phylum. A * indicates the term “unclassified”.

SUPPLEMENT CHAPTER I

111

Fig S4 Most discriminative eukaryotic OTUs of the nine different synthetic polymers and glass (n=5). The

analysis is based on presence / absence matrix of eukaryotic OTUs. Displayed are OTUs jointly contributing, with

a minimum of 0.5%, to the total dissimilarity between different statistically significant (PERMANOVA p<0.05)

glass and synthetic polymer groups. Groups showing both, PERMANOVA and PERMDISP significant p values

were rejected. The amount of contribution is indicated by the colour of cells, darker colours represent higher

contributions. Bold lines indicate OTUs contributing to the same phylum. A * indicates the term “unclassified”. #

indicates the term “Superkingdom”. ` indicates the term “Incertae sedis”.

SUPPLEMENT CHAPTER I

112

Table S1 Sample information about synthetic polymers used within this study.

Polymer Abbreviation Monomer Manufacturer

Low density polyethylene LDPE (C2H4)n ORBITA-FILM GmbH

High density polyethylene HDPE (C2H4)n ORBITA-FILM GmbH

Polypropylene PP (C3H6)n ORBITA-FILM GmbH

Polystyrene PS (C8H8)n Ergo.fol norflex GmbH

Styrene acrylonitrile SAN (C8H8)n-(C3H3N)m Ergo.fol norflex GmbH

Polyurethane prepolymer PESTUR (C4H4O5)n Bayer

Polylactic acid PLA (C3H4O2)n Folienwerk Wolfen GmbH

Polyethylene terephthalate PET (C10H8O4)n Mitsubishi Polyester Film

Polyvynil chloride PVC (C2H2Cl)n Leitz

Table S2 PERMANOVA main tests of prokaryotic and eukaryotic biofilm community on different synthetic

polymers and glass based on Hellinger distance (Prokaryotes, Fungi) and Jaccard (Eukaryotes) of operational

taxonomic units (OTUs). P-values were obtained using type III sums and 9999 permutations under the full model.

d.f.: degrees of freedom, SS: sums of squares; MS: mean squares, perms: number of unique permutations per

comparison. 1Significant results (p (perm) < 0.05) are highlighted in bold.

Prokaryotes

Source of variation d.f. SS MS Pseudo-F p (perm)1 perms

Substrate 9 0.20563 0.0228 3.8052 0.0001 9801

Res 40 0.24017 0.0060

Total 49 0.4458

Eukaryotes

Source of variation d.f. SS MS Pseudo-F p (perm)1 perms

Substrate 9 23052 2561.3 1.2264 0.0001 9495

Res 40 83541 2088.5

Total 49 1.0659E+05

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113

Table S3 PERMANOVA pair-wise tests of prokaryotic and eukaryotic biofilm communities on different

synthetic polymers and glass based on Hellinger distance and Jaccard index (Eukaryotes) of operational

taxonomic units (OTUs). 1Significant results (p (perm) < 0.05) are highlighted in bold.

Prokaryotes Eukaryotes

Comparison t (perm) p (perm)1 t (perm) p (perm)1

Glass vs.

HDPE 3.011 0.008 1.2784 0.0073

LDPE 2.942 0.007 1.238 0.0076

PESTUR 3.02 0.008 1.2408 0.0087

PET 3.333 0.008 1.544 0.0067

PLA 3.284 0.009 1.4681 0.008

PP 2.909 0.008 1.2722 0.0074

PS 3.26 0.008 1.5016 0.0081

PVC 3.01 0.008 1.0849 0.0544

SAN 3.018 0.007 1.1334 0.0583

HDPE vs.

LDPE 0.969 0.48 0.97751 0.6152

PESTUR 1.428 0.017 1.063 0.1058

PET 1.315 0.064 0.99083 0.5195

PLA 1.676 0.007 1.0155 0.3937

PP 1.006 0.458 0.99627 0.5221

PS 1.144 0.16 0.95682 0.7587

PVC 1.346 0.031 1.0655 0.178

SAN 1.077 0.361 0.89472 0.9674

LDPE vs.

PESTUR 1.292 0.097 0.98827 0.5906

PET 1.406 0.051 1.1238 0.0435

PLA 1.782 0.008 1.1419 0.0541

PP 1.104 0.232 0.89898 0.9405

PS 1.174 0.163 0.96542 0.7153

PVC 1.293 0.08 1.1142 0.0369

SAN 0.998 0.383 0.9773 0.6774

PESTUR vs.

PET 1.816 0.007 1.2486 0.0087

PLA 2.158 0.009 1.2087 0.0084

PP 1.374 0.063 1.0048 0.4488

PS 1.716 0.007 1.1234 0.0396

PVC 1.015 0.411 0.90223 0.991

SAN 1.417 0.032 0.88111 0.9828

PET vs.

PLA 1.368 0.054 1.0721 0.1494

PP 1.212 0.15 1.1666 0.018

PS 1.045 0.294 1.0608 0.1756

PVC 1.563 0.007 1.2945 0.0074

SAN 1.498 0.015 1.1092 0.117

PLA vs.

PP 1.758 0.009 1.2538 0.0155

PS 1.406 0.024 1.1106 0.0578

PVC 1.898 0.007 1.1577 0.0221

SAN 1.523 0.008 1.0488 0.224

PP vs.

PS 1.021 0.37 0.98839 0.5434

PVC 1.282 0.062 1.015 0.3793

SAN 1.135 0.184 0.98037 0.5587

PS vs. PVC 1.482 0.016 1.1766 0.0304

SAN 1.19 0.1 0.96935 0.6431

PVC vs. SAN 1.199 0.065 0.89216 0.9431

SUPPLEMENT CHAPTER I

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Table S4 PERMDISP pair-wise tests of prokaryotic and eukaryotic biofilm communities on different synthetic

polymers and glass based on Hellinger distance and Jaccard index (Eukaryotes) of operational taxonomic units

(OTUs). 1Significant results (p (perm) < 0.05) are highlighted in bold.

Prokaryotes Eukaryotes

Comparison t (perm) p (perm)1 t (perm) p (perm)1

Glass vs.

HDPE 2.982 0.009 6.1072 0.0083

LDPE 2.319 0.072 5.93 0.009

PESTUR 2.964 0.023 1.6132 0.1244

PET 2.05 0.082 8.8325 0.0082

PLA 2.204 0.074 2.2482 0.0069

PP 1.696 0.114 6.0339 0.0079

PS 3.333 0.009 4.6126 0.0075

PVC 2.541 0.039 2.4745 0.01

SAN 2.688 0.025 4.9305 0.0094

HDPE vs.

LDPE 0.367 0.701 0.026393 0.9771

PESTUR 0.724 0.526 3.7934 0.0087

PET 1.248 0.173 0.0027481 1

PLA 0.279 0.803 2.4303 0.0497

PP 2.101 0.007 2.5103 0.0074

PS 0.587 0.541 2.074 0.0661

PVC 0.251 0.848 3.1883 0.008

SAN 0.109 0.931 2.7485 0.0074

LDPE vs.

PESTUR 0.902 0.401 3.7306 0.0085

PET 0.611 0.5 0.029177 0.9775

PLA 0.041 0.938 2.4111 0.0621

PP 1.174 0.278 2.4541 0.007

PS 0.803 0.504 2.0464 0.0556

PVC 0.122 0.946 3.1387 0.0075

SAN 0.397 0.701 2.6899 0.0097

PESTUR vs.

PET 1.548 0.195 4.7257 0.0084

PLA 0.803 0.481 0.84592 0.45

PP 2.08 0.127 2.3404 0.0703

PS 0.331 0.805 2.1231 0.0875

PVC 0.839 0.478 0.6704 0.571

SAN 0.547 0.647 1.8703 0.102

PET vs.

PLA 0.595 0.587 2.8281 0.0462

PP 0.654 0.458 3.7175 0.008

PS 1.767 0.073 2.7087 0.0207

PVC 0.807 0.425 3.993 0.0102

SAN 1.075 0.365 3.8906 0.007

PLA vs.

PP 1.09 0.352 0.83729 0.4529

PS 0.669 0.556 0.85598 0.4479

PVC 0.068 0.982 0.28652 0.7728

SAN 0.324 0.818 0.52505 0.632

PP vs.

PS 2.655 0.024 0.16079 0.8325

PVC 1.445 0.175 1.5234 0.2436

SAN 1.675 0.175 0.54294 0.4593

PS vs. PVC 0.729 0.56 1.4193 0.2688

SAN 0.338 0.799 0.56216 0.4997

PVC vs. SAN 0.3 0.792 1.0833 0.3514

SUPPLEMENT CHAPTER I

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Table S5 SIMPER analysis of prokaryotic communities jointly contributing to the total similarity within and

dissimilarity between different groups of synthetic polymers. Av.Si%: average percentage similarity within the

different groups, Av.δi%: average dissimilarity between the different groups.

Av.Si% Av.δi%

LDPE 95.74 LDPE 4.13

PP 95.51 PP 4.25 4.37

PS 95.97 PS 4.19 4.26 4.35

PET 95.52 PET 4.71 4.83 4.86 4.33

PLA 96.01 PLA 4.84 4.96 5.29 4.36 4.69

SAN 95.92 SAN 4.16 4.15 4.46 4.23 4.92 4.62

PESTUR 96.16 PESTUR 4.40 4.29 4.57 4.75 5.20 5.49 4.42

PVC 96.14 PVC 4.34 4.34 4.54 4.50 4.98 5.26 4.26 3.90

Glass 95.25 Glass 7.64 7.71 7.79 8.11 8.93 8.60 7.72 7.67 7.77

HDPE 95.86 HDPE LDPE PP PS PET PLA SAN PESTUR PVC

Table S6 SIMPER analysis of eukaryotic communities jointly contributing to the total similarity within and

dissimilarity between different groups of synthetic polymers. Av.Si%: average percentage similarity within the

different groups, Av.δi%: average dissimilarity between the different groups.

Av.Si% Av.δi%

LDPE 45.78 LDPE 53.80

PP 52.26 PP 50.87 49.50

PS 51.90 PS 50.45 50.63 47.74

PET 45.97 PET 53.92 56.34 53.66 52.03

PLA 54.45 PLA 50.07 52.17 50.56 48.44 50.91

SAN 53.09 SAN 48.99 50.22 47.04 47.07 52.23 46.91

PESTUR 57.23 PESTUR 49.39 48.33 45.31 47.17 52.43 47.10 43.39

PVC 55.45 PVC 50.33 51.17 46.34 48.93 54.28 47.27 44.36 42.49

Glass 60.66 Glass 51.11 50.47 47.41 51.61 56.22 49.42 44.88 44.18 43.01

HDPE 45.88 HDPE LDPE PP PS PET PLA SAN PESTUR PVC

.

SUPPLEMENT CHAPTER I

116

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hag

ia

Ekh

idna

174

0.6

0

0.5

7

0.5

4

0.6

4

0.5

0

0.4

2

0.5

8

0.5

2

0.5

7

0.5

7

Fla

mm

eovi

rgace

ae_

uncu

ltu

red*

1

85

0.6

5

0.8

3

0.8

6

0.8

0

0.8

3

0.8

1

0.8

3

0.8

4

0.7

9

0.8

1

Rho

do

ther

ma

cea

e_un

cult

ure

d*

1

90

0.8

7

0.9

6

0.9

4

0.9

5

1.0

3

0.9

4

0.9

9

0.9

9

0.9

7

0.9

2

Fla

voba

cter

iia

Ow

enw

eeks

ia

200

0.5

9

0.5

3

0.5

1

0.5

2

0.5

1

0.4

8

0.5

4

0.5

2

0.4

9

0.4

8

Aqu

ibact

er

206

0.6

7

0.5

8

0.5

4

0.5

3

0.5

7

0.6

2

0.5

8

0.6

0

0.5

2

0.5

9

Ba

cter

oid

etes

G

ilvi

ba

cter

2

31

0.6

6

0.9

4

1.0

4

0.9

5

1.0

0

0.9

2

1.1

0

1.0

1

0.9

8

0.9

7

Lep

tob

act

eriu

m

240

0.3

9

0.5

6

0.3

9

0.5

8

0.5

6

0.6

4

0.5

3

0.5

2

0.5

4

0.5

8

Ma

rixa

nth

om

ona

s 2

46

0.3

9

0.4

6

0.4

3

0.4

8

0.4

8

0.4

5

0.6

2

0.5

4

0.4

5

0.4

3

Ulv

iba

cter

2

76

0.5

0

0.7

0

0.6

9

0.7

5

0.7

6

0.6

9

0.7

5

0.7

6

0.6

7

0.6

7

Sph

ing

oba

cter

iia

C

hit

inop

hag

ace

ae_

uncu

ltu

red*

2

94

1.0

7

0.9

7

0.9

7

0.8

9

0.9

3

0.8

7

0.9

9

0.9

5

0.9

0

0.9

1

Ta

ble

S7

Ch

ara

cter

isti

c p

rok

ary

oti

c O

TU

s o

f th

e n

ine

dif

feren

t sy

nth

etic

po

lym

ers

an

d g

lass

(n

=5

). O

TU

s w

ith

a m

ean

rel

ativ

e ab

und

ance

of

at lea

st 0

.1%

(n=

5)

in a

t le

ast

on

e su

bst

rate

ty

pe

wer

e an

aly

sed

. H

igh

lig

hte

d i

n g

reen

are

OT

Us

join

tly

con

trib

uti

ng

, w

ith

a m

inim

um

of

1%

to

th

e to

tal

sim

ilar

ity o

f g

lass

an

d

syn

thet

ic p

oly

mer

gro

up

s. B

old

lin

es i

nd

icat

e O

TU

s co

ntr

ibu

tin

g t

o t

he

sam

e p

hy

lum

. A

* i

nd

icat

es t

he

term

“un

clas

sifi

ed”.

A

v.S

i.%

SUPPLEMENT CHAPTER I

Table S7: continued

117

Ba

cter

oid

etes

Lew

inel

la

309

1.0

5

0.9

6

0.9

9

0.8

8

0.9

5

0.9

2

0.9

7

0.9

7

0.8

9

0.9

4

Pha

eoda

ctyl

iba

cter

3

10

0.6

3

0.6

1

0.6

6

0.6

0

0.5

8

0.6

7

0.6

3

0.5

8

0.5

8

0.6

6

Po

rtib

act

er

311

1.1

3

0.9

8

0.9

6

0.9

6

0.9

1

0.9

4

1.0

0

0.9

9

0.9

1

0.9

8

Sap

rosp

ira

ceae_

un

cult

ure

d*

314

1.1

3

1.0

0

1.0

1

1.0

0

1.0

0

1.0

2

1.0

7

1.0

2

1.0

1

1.0

7

Chla

myd

iae

Chla

myd

iae

Chla

myd

iace

ae_

uncu

ltu

red*

3

28

0.6

2

0.4

6

0.5

1

0.4

4

0.4

6

0.4

6

0.4

7

0.4

7

0.5

2

0.4

8

Cand

idatu

s F

rits

chea

3

34

0.4

1

0.4

1

0.4

2

0.3

5

0.4

0

0.3

9

0.4

2

0.4

4

0.4

3

0.4

4

Chlo

rofl

exi

Ard

enti

cate

nia

A

rden

tica

ten

ale

s*

355

0.6

5

0.6

2

0.6

1

0.8

3

0.7

1

0.7

5

0.6

7

0.7

4

0.8

2

0.6

9

Ard

enti

cate

nia

_un

cult

ure

d*

3

56

1.2

2

1.1

6

1.1

9

1.1

9

1.1

9

1.1

5

1.2

0

1.1

7

1.1

7

1.1

3

Cald

ilin

eae

Cald

ilin

eace

ae_

uncu

ltu

red*

3

59

1.5

5

1.5

7

1.5

0

1.5

1

1.5

7

1.6

3

1.5

2

1.5

9

1.5

1

1.5

0

Cya

nob

act

eria

Cya

nob

act

eria

_C

hlo

ropla

st*

C

yanob

act

eria

_C

hlo

ropla

st*

3

78

0.5

4

0.5

2

0.5

0

0.4

7

0.4

7

0.5

1

0.5

1

0.4

6

0.4

8

0.5

4

Mel

ain

abact

eria

O

bsc

uri

ba

cter

ale

s*

389

0.4

6

0.5

7

0.5

6

0.5

4

0.5

3

0.5

8

0.5

6

0.5

9

0.5

6

0.5

8

Va

mpir

ovi

bri

ona

les*

3

90

0.4

4

0.5

3

0.5

2

0.4

9

0.5

2

0.5

4

0.4

6

0.5

3

0.5

0

0.5

2

Def

erri

ba

cter

es

Def

erri

ba

cter

es I

nce

rta

e S

edis

*

Cald

ith

rix

391

0.7

5

0.7

1

0.7

0

0.7

0

0.7

0

0.6

3

0.6

9

0.7

0

0.6

9

0.6

8

Dei

no

cocc

us-

Th

erm

us

Dei

no

cocc

i T

ruep

era

393

0.7

9

0.7

2

0.6

9

0.6

7

0.7

3

0.7

8

0.7

3

0.7

2

0.6

7

0.7

5

Gem

ma

tim

on

ad

etes

G

emm

ati

mon

ad

etes

G

emm

ati

mon

ad

etes

_B

D2

-11

ter

rest

ria

l g

rou

p*

5

40

1.0

4

1.1

8

1.1

6

1.1

3

1.2

5

1.2

0

1.1

8

1.2

0

1.1

8

1.1

3

Gem

ma

tim

on

ad

etes

_P

AU

C4

3f

ma

rine

ben

thic

gro

up

*

544

0.3

8

0.5

6

0.5

9

0.5

0

0.6

2

0.5

9

0.5

5

0.6

0

0.5

5

0.5

7

La

tesc

iba

cter

ia

La

tesc

iba

cter

ia*

L

ate

scib

act

eria

*

551

0.9

0

0.6

6

0.6

8

0.6

5

0.6

5

0.6

4

0.6

3

0.6

6

0.6

3

0.6

5

Len

tisp

ha

erae

Len

tisp

ha

erae_

LD

1-P

B3

*

Len

tisp

ha

erae_

LD

1-P

B3

*

560

0.4

7

0.5

2

0.5

1

0.5

2

0.5

7

0.5

8

0.5

0

0.5

1

0.5

1

0.5

6

Oli

go

sph

aer

ia

Oli

go

sph

aer

ia*

5

65

1.1

2

1.0

2

1.0

4

1.0

9

0.9

0

0.7

9

1.0

2

0.9

3

1.0

4

1.0

4

Nit

rosp

irae

Nit

rosp

ira

Nit

rosp

ira

576

1.7

1

1.7

8

1.7

9

1.9

0

1.8

0

1.6

5

1.7

7

1.7

1

1.8

3

1.7

5

Om

nit

rop

hic

a

Om

nit

rop

hic

a_

NP

L-U

PA

2*

Om

nit

rop

hic

a_

NP

L-U

PA

2*

579

0.6

4

0.6

1

0.6

1

0.6

4

0.6

1

0.5

7

0.6

2

0.6

0

0.6

5

0.6

0

Pa

rcub

act

eria

P

arc

ub

act

eria

*

Pa

rcub

act

eria

*

582

0.7

8

0.5

7

0.6

0

0.5

5

0.6

4

0.6

4

0.5

9

0.6

1

0.5

5

0.5

7

Pla

nct

om

ycet

es

Pla

nct

om

ycet

es_02

8H

05

-P-B

N-P

5*

Pla

nct

om

ycet

es_02

8H

05

-P-B

N-P

5*

583

0.6

2

0.5

6

0.5

4

0.5

6

0.5

1

0.5

2

0.5

9

0.5

0

0.5

4

0.5

5

Pla

nct

om

ycet

es_B

D7

-11*

Pla

nct

om

ycet

es_B

D7

-11*

584

0.6

5

0.5

6

0.5

5

0.5

9

0.4

8

0.4

9

0.5

7

0.5

5

0.6

1

0.6

0

Pla

nct

om

ycet

es_

OM

190

*

Pla

nct

om

ycet

es_

OM

190

*

586

1.4

5

1.4

3

1.4

1

1.4

3

1.4

0

1.2

9

1.4

6

1.3

8

1.4

3

1.4

1

Ph

ycis

ph

aer

ae

Ph

ycis

ph

aer

ace

ae_

SM

1A

02*

6

06

0.5

4

0.6

6

0.7

0

0.6

9

0.7

5

0.6

5

0.6

7

0.7

1

0.7

0

0.7

2

Ph

ycis

ph

aer

ace

ae_

Ura

nia

-1B

-19 m

ari

ne

sedim

ent

gro

up*

6

07

0.7

5

0.7

9

0.8

2

0.8

2

0.8

7

0.8

1

0.8

4

0.8

5

0.8

5

0.8

3

Ph

ycis

ph

aer

ace

ae_

uncu

ltu

red*

6

09

1.1

6

0.9

7

0.9

2

1.0

3

0.9

2

0.8

5

0.9

7

0.9

0

1.0

5

0.9

8

Ph

ycis

ph

aer

ae_

S-7

0*

612

0.3

6

0.4

8

0.4

7

0.4

9

0.4

4

0.5

4

0.4

7

0.5

2

0.5

0

0.5

3

SUPPLEMENT CHAPTER I

Table S7: continued

118

P

lan

cto

myc

etes

_P

la3 l

inea

ge*

P

lan

cto

myc

etes

_P

la3 l

inea

ge*

6

16

0.8

4

0.7

5

0.7

6

0.7

7

0.7

2

0.6

8

0.7

5

0.7

5

0.7

4

0.7

6

P

lan

cto

myc

etes

_P

la4 l

inea

ge*

P

lan

cto

myc

etes

_P

la4 l

inea

ge*

6

17

0.5

7

0.5

4

0.5

7

0.5

7

0.4

8

0.4

2

0.5

7

0.5

0

0.5

2

0.5

2

Pla

nct

om

ycet

es

Pla

nct

om

ycet

aci

a

Pla

nct

om

ycet

ace

ae*

6

19

0.2

3

0.4

3

0.4

9

0.5

0

0.4

2

0.4

6

0.5

4

0.4

9

0.4

8

0.5

4

Bla

sto

pir

ellu

la

620

1.2

0

1.1

6

1.1

6

1.2

0

1.1

7

1.1

6

1.1

6

1.1

3

1.1

3

1.1

3

Byt

hopir

ellu

la

621

0.7

7

1.0

1

1.0

7

1.0

1

1.1

0

1.0

5

1.0

6

1.0

3

1.0

1

1.0

8

Pla

nct

om

ycet

ace

ae_

Pir

4 l

inea

ge*

6

25

0.6

0

0.7

7

0.8

3

0.7

6

0.8

1

0.8

1

0.7

8

0.7

3

0.7

4

0.8

1

Pir

ellu

la

626

0.4

8

0.5

1

0.5

1

0.5

4

0.5

2

0.5

0

0.5

0

0.5

3

0.5

4

0.5

6

Pla

nct

om

yces

6

27

1.1

7

1.0

8

1.0

9

1.0

6

1.0

7

1.0

7

1.0

8

1.0

7

1.0

5

1.0

8

Rho

dop

irel

lula

6

28

1.0

8

1.0

9

1.1

3

1.0

7

1.0

7

1.0

4

1.0

8

1.0

7

1.0

5

1.1

1

Pla

nct

om

ycet

ace

ae_

un

cult

ure

d*

6

32

0.9

0

0.9

2

0.9

6

0.9

3

0.8

7

0.8

5

0.9

7

0.8

9

0.8

9

0.9

4

P

lan

cto

myc

etes

_va

din

HA

49

*

Pla

nct

om

ycet

es_va

din

HA

49

*

635

0.4

8

0.5

4

0.5

4

0.5

4

0.5

1

0.4

8

0.5

2

0.5

1

0.5

0

0.5

7

P

rote

oba

cter

ia_

AE

GE

AN

-24

5*

Pro

teo

ba

cter

ia_

AE

GE

AN

-24

5*

637

0.8

3

0.7

2

0.7

2

0.7

0

0.6

9

0.7

2

0.7

6

0.7

3

0.7

3

0.6

9

P

rote

oba

cter

ia_

AR

KIC

E-9

0*

Pro

teo

ba

cter

ia_

AR

KIC

E-9

0*

639

0.7

0

0.6

4

0.5

7

0.6

3

0.5

5

0.5

2

0.5

8

0.5

5

0.4

8

0.5

4

A

lpha

pro

teo

ba

cter

ia

Alp

ha

pro

teo

ba

cter

ia_

DB

1-1

4*

661

0.8

6

0.8

8

0.9

3

0.9

6

1.0

0

1.0

5

0.8

9

0.9

2

1.0

1

0.9

6

Alp

ha

pro

teo

ba

cter

ia_

OC

S11

6 c

lad

e*

666

1.2

5

1.3

2

1.2

8

1.2

5

1.4

2

1.3

5

1.2

8

1.3

5

1.2

6

1.3

0

Pa

rvula

rcula

6

68

0.8

1

0.9

0

0.8

9

0.9

1

0.9

5

0.9

4

0.9

2

0.9

2

0.8

9

0.8

9

Fil

om

icro

biu

m

691

1.0

3

1.1

2

1.0

9

1.0

7

1.1

8

1.2

0

1.1

3

1.1

4

1.0

8

1.0

9

Ped

om

icro

biu

m

694

0.4

9

0.7

7

0.7

6

0.7

3

0.8

2

0.8

1

0.7

8

0.8

0

0.7

3

0.7

8

Hyp

ho

mic

robia

cea

e_u

ncu

ltu

red_u

nid

6

98

0.7

0

0.7

6

0.7

0

0.6

6

0.7

2

0.7

9

0.7

3

0.7

5

0.6

7

0.7

1

Pse

uda

hre

nsi

a

715

1.0

2

0.9

9

0.9

6

0.9

5

1.0

1

1.0

2

0.9

9

1.0

3

0.9

1

1.0

0

And

erse

nie

lla

730

0.5

9

0.6

3

0.6

0

0.5

6

0.6

6

0.6

4

0.6

2

0.6

3

0.6

2

0.6

3

Rho

dob

iace

ae_

un

cult

ure

d*

7

38

0.6

5

0.7

3

0.7

1

0.6

5

0.7

2

0.7

8

0.6

9

0.7

6

0.7

1

0.7

4

Pro

teo

ba

cter

ia

R

ho

dob

act

erace

ae_

uncu

ltu

red*

8

31

1.2

1

1.0

8

1.1

4

1.1

6

1.1

4

1.1

1

1.1

1

1.1

2

1.1

6

1.1

4

Rho

do

spir

illa

les_

AT

-s3

-44

*

833

0.7

2

0.6

9

0.7

4

0.8

7

0.7

5

0.7

8

0.6

6

0.7

2

0.8

6

0.7

1

Def

luvi

icocc

us

851

0.7

8

0.8

3

0.8

1

0.8

7

0.7

8

0.8

0

0.7

8

0.8

1

0.8

2

0.8

4

Pel

agib

ius

864

0.5

4

0.5

0

0.5

2

0.6

5

0.5

4

0.5

8

0.5

2

0.5

6

0.6

0

0.5

9

Rho

do

spir

illa

cea

e_uncu

ltu

red*

8

72

1.1

4

1.2

1

1.2

6

1.2

9

1.2

3

1.1

9

1.2

2

1.2

5

1.2

6

1.2

2

Ric

kett

siale

s_SM

2D

12

*

907

0.8

5

0.8

1

0.8

3

0.8

5

0.8

3

0.8

7

0.8

0

0.8

3

0.9

0

0.8

6

Sph

ing

orh

ab

du

s 9

34

0.5

2

0.4

9

0.5

4

0.4

7

0.4

9

0.5

2

0.4

7

0.5

1

0.4

0

0.5

0

B

eta

pro

teo

ba

cter

ia

Lim

no

ba

cter

9

48

0.5

1

0.5

7

0.5

7

0.5

6

0.6

4

0.5

8

0.6

0

0.6

1

0.5

5

0.5

5

Hyd

rogen

op

hil

ace

ae_

un

cult

ure

d*

9

94

0.5

6

0.5

2

0.5

7

0.6

1

0.4

2

0.4

2

0.5

2

0.4

2

0.5

3

0.4

9

SUPPLEMENT CHAPTER I

Table S7: continued

119

Nit

roso

mona

s 1

005

1.0

5

1.2

8

1.3

1

1.2

9

1.3

2

1.2

5

1.2

9

1.2

4

1.2

7

1.3

0

Del

tap

rote

obact

eria

Ba

cter

iovo

raca

cea

e*

1021

0.5

3

0.5

3

0.5

2

0.5

2

0.4

0

0.5

2

0.5

2

0.5

3

0.5

3

0.5

5

H

alo

ba

cter

iovo

rax

1024

0.6

1

0.4

2

0.4

6

0.5

1

0.4

3

0.4

8

0.4

6

0.4

8

0.5

1

0.5

2

P

ered

iba

cter

1

025

0.5

7

0.5

2

0.5

6

0.6

5

0.4

9

0.6

0

0.5

5

0.5

9

0.6

7

0.6

4

B

del

lovi

bri

o

1027

0.6

9

0.7

0

0.6

9

0.7

2

0.6

7

0.6

8

0.7

1

0.6

7

0.7

2

0.6

9

B

del

lovi

bri

ona

cea

e_O

M2

7 c

lad

e*

1028

0.7

9

0.8

3

0.8

1

0.7

8

0.8

1

0.8

5

0.8

3

0.8

5

0.8

3

0.8

1

C

and

idatu

s E

nto

theo

nel

la

1058

1.5

7

1.2

2

1.3

1

1.3

0

1.1

7

1.1

8

1.2

8

1.3

1

1.2

9

1.2

3

N

itro

spin

a

1059

0.9

6

0.8

2

0.7

6

0.8

3

0.6

4

0.6

3

0.7

1

0.6

7

0.8

2

0.7

7

D

esu

lfu

rell

ace

ae_

un

cult

ure

d*

1

067

0.4

8

0.4

2

0.4

8

0.5

3

0.3

8

0.4

7

0.2

7

0.4

3

0.4

4

0.4

0

D

esu

lfu

rom

onad

ale

s_G

R-W

P33

-58*

1076

0.6

3

0.6

2

0.6

1

0.6

4

0.6

0

0.6

2

0.6

2

0.6

3

0.6

6

0.6

6

D

elta

pro

teo

bact

eria

_G

R-W

P3

3-3

0*

1084

0.4

9

0.5

3

0.5

5

0.5

2

0.5

5

0.5

9

0.5

0

0.5

4

0.5

3

0.5

6

H

ali

an

giu

m

1093

0.7

8

0.7

9

0.7

7

0.7

2

0.7

7

0.8

3

0.7

6

0.8

1

0.7

8

0.8

0

N

ann

ocy

stis

1

098

0.6

0

0.6

6

0.5

8

0.6

3

0.6

3

0.6

5

0.6

8

0.6

3

0.5

7

0.6

3

M

yxo

cocc

ale

s_P

3O

B-4

2*

1102

0.7

6

0.7

6

0.7

7

0.7

5

0.7

3

0.7

1

0.7

6

0.7

5

0.7

7

0.7

4

S

and

ara

cin

us

1112

0.7

9

0.7

0

0.6

9

0.6

7

0.7

5

0.7

3

0.7

2

0.7

4

0.7

1

0.6

8

S

and

ara

cin

ace

ae_

uncu

ltu

red*

1

113

0.6

3

0.7

2

0.7

0

0.6

6

0.7

6

0.7

5

0.7

0

0.7

4

0.6

9

0.7

2

O

ligofl

exa

les*

1

120

0.6

1

0.5

4

0.5

6

0.5

6

0.5

7

0.6

0

0.5

9

0.5

7

0.5

9

0.6

1

O

ligofl

exa

cea

e*

1121

0.5

9

0.5

5

0.5

5

0.5

9

0.5

3

0.6

0

0.5

4

0.6

1

0.6

3

0.5

9

D

elta

pro

teo

bact

eria

_Sh

765

B-T

zT-2

9*

1123

1.4

3

1.6

1

1.6

2

1.6

2

1.7

1

1.6

4

1.6

5

1.6

2

1.6

3

1.5

9

Pro

teo

ba

cter

ia

Ep

silo

np

rote

oba

cter

ia

Sulf

uro

vum

1

139

0.5

8

0.6

1

0.5

4

0.5

3

0.5

9

0.6

7

0.5

9

0.6

1

0.5

4

0.6

1

Ga

mm

ap

rote

obact

eria

Shew

an

ella

1

173

0.4

1

0.3

2

0.2

7

0.2

3

0.2

9

0.2

9

0.2

6

0.2

8

0.2

8

0.3

0

A

renic

ella

cea

e*

1174

0.5

9

0.3

5

0.3

2

0.3

3

0.3

4

0.4

4

0.3

6

0.2

9

0.3

1

0.3

8

G

am

map

rote

obact

eria

_B

D3

-1*

1178

0.4

6

0.5

7

0.5

6

0.5

8

0.5

6

0.6

3

0.4

8

0.5

5

0.5

5

0.5

4

G

am

map

rote

obact

eria

_B

D7

-8 m

ari

ne

gro

up*

1

179

0.9

0

0.7

2

0.7

1

0.7

6

0.6

4

0.6

5

0.7

4

0.6

9

0.7

4

0.7

1

C

ellv

ibri

ona

ceae_

un

cult

ure

d*

1

200

0.7

6

0.6

8

0.6

6

0.6

6

0.6

6

0.6

5

0.6

8

0.6

7

0.7

2

0.6

8

H

ali

ea

1204

0.5

8

0.5

7

0.5

2

0.5

0

0.5

5

0.5

6

0.5

6

0.5

2

0.5

2

0.5

4

H

ali

eace

ae_

OM

60

(NO

R5

) cl

ade*

1

207

0.5

7

0.5

5

0.5

7

0.4

9

0.5

6

0.5

8

0.5

8

0.5

5

0.5

3

0.5

6

P

ort

ico

ccus

1211

0.8

1

0.6

0

0.5

9

0.6

2

0.6

3

0.5

9

0.6

5

0.6

4

0.6

1

0.5

8

S

pon

gii

ba

cter

ace

ae_

BD

1-7

cla

de*

1

214

0.8

5

0.5

9

0.6

1

0.6

3

0.5

7

0.5

9

0.6

3

0.5

9

0.6

8

0.6

4

N

itro

soco

ccus

1222

1.1

5

1.0

4

1.0

6

1.0

7

1.0

3

1.0

2

1.0

4

1.0

2

1.0

5

1.0

3

G

ranu

losi

cocc

us

1243

1.0

9

1.3

5

1.3

4

1.3

4

1.4

1

1.3

7

1.3

4

1.3

5

1.3

6

1.3

1

G

am

map

rote

obact

eria

_E

01

-9C

-26

ma

rin

e g

rou

p*

1

246

0.4

7

0.4

9

0.4

8

0.4

5

0.4

3

0.5

2

0.4

5

0.4

9

0.4

6

0.5

1

SUPPLEMENT CHAPTER I

Table S7: continued

120

Ga

mm

ap

rote

obact

eria

_H

OC

36*

1

267

0.6

1

0.6

0

0.6

3

0.6

2

0.6

2

0.6

5

0.6

4

0.6

5

0.6

1

0.6

2

G

am

map

rote

obact

eria

_K

I89

A c

lad

e*

1269

1.1

5

1.2

3

1.2

0

1.2

5

1.2

3

1.1

6

1.2

4

1.2

0

1.2

2

1.1

4

P

seudo

spir

illu

m

1327

0.7

0

0.7

8

0.7

8

0.7

4

0.6

9

0.6

3

0.7

8

0.6

7

0.7

9

0.6

9

M

ari

nic

ella

1

341

0.8

4

0.8

8

0.8

8

0.8

6

0.9

1

0.9

5

0.8

9

0.9

0

0.8

8

0.8

9

X

an

tho

mon

ada

les_

JTB

255

ma

rin

e ben

thic

gro

up

*

1390

1.5

1

1.5

6

1.5

2

1.6

0

1.6

4

1.5

7

1.5

4

1.5

6

1.6

0

1.5

1

Pro

teo

ba

cter

ia

Xan

tho

mon

ada

les_

un

cult

ure

d*

1

407

0.9

0

0.9

9

0.9

8

1.0

3

1.0

1

1.0

1

0.9

6

0.9

6

1.0

5

0.9

9

P

rote

oba

cter

ia_

JTB

23*

P

rote

oba

cter

ia_

JTB

23*

1

408

0.6

5

0.5

7

0.5

7

0.5

6

0.5

2

0.4

9

0.5

9

0.5

5

0.5

9

0.5

5

P

rote

oba

cter

ia_S

C3

-20*

Pro

teo

ba

cter

ia_S

C3

-20*

1413

0.5

4

0.7

3

0.6

9

0.7

1

0.6

9

0.6

5

0.7

2

0.7

1

0.7

4

0.7

2

P

rote

oba

cter

ia_S

PO

TS

OC

T00

m8

3*

P

rote

oba

cter

ia_S

PO

TS

OC

T00

m8

3*

1

414

1.0

4

1.0

3

1.0

6

1.0

5

1.0

5

1.0

2

1.0

4

1.0

3

1.0

4

1.0

0

P

rote

oba

cter

ia_

TA

18

*

Pro

teo

ba

cter

ia_

TA

18

*

1416

0.7

4

0.7

5

0.7

9

0.7

4

0.8

1

0.9

1

0.7

6

0.7

9

0.7

7

0.8

1

Sacc

hari

bact

eria

S

acc

hari

bact

eria

*

Sacc

hari

bact

eria

*

1420

0.8

4

0.7

3

0.7

3

0.6

2

0.7

4

0.7

9

0.6

9

0.7

4

0.6

4

0.6

9

Ver

ruco

mic

rob

ia

Op

ituta

e C

era

sico

ccu

s 1

454

0.6

0

0.5

6

0.5

0

0.5

5

0.5

0

0.4

7

0.4

9

0.4

8

0.5

4

0.4

9

Ver

ruco

mic

rob

iae

Ver

ruco

mic

rob

iale

s_D

EV

007

*

1470

0.7

8

0.6

9

0.7

1

0.6

6

0.6

9

0.6

9

0.6

9

0.7

0

0.6

7

0.6

6

SUPPLEMENT CHAPTER I

121

Kin

gd

om

Ph

ylu

mC

lass

Ge

nu

sO

TU

Glass

HDPE

LDPE

PESTUR

PET

PLA

PP

PS

PVC

SAN

Euk

ary

ota

*E

ukar

yo

ta*

Euk

ary

ota

*E

ukar

yo

ta*

10

00

00

00

00

0

Am

oeb

ozo

aD

isco

sea

Fla

bell

inia

Ver

mis

tell

a2

0.4

80

00

.37

00

00

0.1

0.3

3

Van

nel

la3

0.2

40

00

00

00

0.6

80

.12

Tub

ulin

eaE

uam

oeb

ida

Har

tman

nel

la4

00

00

00

00

00

Lep

tom

yx

ida

Lep

tom

yx

ida*

50

.08

00

00

00

00

0

Lep

tom

yx

a6

0.2

30

.50

.15

00

.69

0.3

90

.11

0.9

60

.33

0.4

2

Rh

izam

oeb

a8

0.0

80

.16

00

00

00

00

Ch

loro

pla

stid

a*C

hlo

rop

last

ida*

Ch

loro

pla

stid

a*9

00

00

00

00

00

Ph

ragm

op

last

op

hy

ta*

Co

leo

chae

te1

20

00

0.1

10

00

00

0

Ph

ragm

op

last

op

hy

taE

mbr

yo

ph

yta

Em

bry

op

hy

ta_

Am

b-1

8S-

11

21

*1

30

.48

0.1

80

.48

0.3

50

.18

0.1

30

00

.10

.11

Mag

no

lio

ph

yta

*1

40

00

00

00

00

0

Ho

rdeu

m1

50

00

00

00

00

0

Py

rus

16

00

00

00

00

00

Ch

loro

pla

stid

a_C

lade

VII

*C

hlo

rop

last

ida_

Subc

lade

A*

20

00

00

00

00

00

Mam

iell

op

hy

ceae

Mam

iell

op

hy

ceae

*2

10

00

00

0.1

30

00

0

Mam

iell

op

hy

ceae

_D

SGM

-81

*2

20

00

0.3

40

.25

0.1

30

00

0

Ch

loro

pla

stid

aC

rust

om

asti

x2

30

00

00

.22

0.9

10

00

0.1

1

Do

lich

om

asti

x2

40

00

.15

0.1

20

00

0.1

70

.33

0

Mam

iell

a2

50

.25

0.5

40

00

0.1

40

.14

00

0

Mic

rom

on

as2

60

.08

00

00

0.9

10

.14

00

.31

0

Ch

loro

pla

stid

a*O

stre

oco

ccus

27

00

00

00

00

00

Nep

hro

selm

ido

ph

yce

aeN

eph

rose

lmis

28

00

00

00

0.4

00

.31

0

Pra

sin

op

hy

tae*

29

0.4

90

.97

0.1

70

.35

1.3

90

.79

0.3

80

.43

1.0

80

.1

Pra

sin

ode

rma

30

00

00

.11

00

.13

0.1

10

.19

0.3

10

.16

Pra

sin

op

hy

tae

Cy

mbo

mo

nas

31

00

00

00

00

00

Po

lybl

eph

arid

es3

20

00

00

00

00

0

Ta

ble

S8

Ch

ara

cter

isti

c eu

ka

ryo

tic

OT

Us

of

the

nin

e d

iffe

ren

t sy

nth

etic

po

lym

ers

an

d g

lass

(n

=5

). A

nal

ysi

s is

bas

ed o

n p

rese

nce

/ a

bse

nce

mat

rix

of

det

ecte

d O

TU

s. H

igh

lig

hte

d i

n g

reen

are

OT

Us

join

tly

co

ntr

ibu

tin

g,

wit

h a

min

imu

m o

f 1%

to

th

e to

tal

sim

ilar

ity

of

gla

ss a

nd

sy

nth

etic

po

lym

er g

roup

s. B

old

lin

es in

dic

ate

OT

Us

con

trib

uti

ng

to th

e sa

me

kin

gdo

m. A

* in

dic

ates

th

e te

rm “

un

clas

sifi

ed”,

# in

dic

ates

th

e te

rm “

Su

per

kin

gd

om

”, `

in

dic

ates

th

e te

rm “

Ince

rta

e

Sed

is”.

A

v.S

i.%

SUPPLEMENT CHAPTER I

Table S8: continued

122

Pte

rosp

erm

a3

30

00

00

0.1

40

0.1

40

.63

0

Py

ram

imo

nas

34

00

00

00

00

00

Tre

boux

iop

hy

ceae

Tre

boux

iop

hy

ceae

*3

50

.81

0.2

10

0.1

10

0.7

90

.14

0.4

40

.31

0.7

2

Pro

toth

eca

37

00

00

00

00

00

Ch

loro

pla

stid

aC

hlo

rop

last

ida*

Dic

tyo

chlo

rop

sis

39

00

00

00

00

00

Ell

ipto

chlo

ris

40

00

00

00

00

00

Ko

liel

la4

10

00

00

00

00

0

Ulv

op

hy

ceae

Ulv

op

hy

ceae

*4

30

.08

0.6

0.1

70

.11

00

.46

0.4

60

0.3

0

Bli

din

gia

45

00

00

00

00

00

Pse

uden

docl

on

ium

46

00

00

.11

00

00

00

Ulv

a4

70

.48

1.7

40

.17

1.2

0.6

90

.39

0.8

50

.14

0.6

30

.82

Ulv

ella

48

00

00

00

00

00

Ch

loro

pla

stid

a*C

hlo

rop

last

ida*

49

00

00

00

00

00

Rh

odo

ph

yce

aeF

lori

deo

ph

yci

dae

Co

rall

ino

ph

yci

dae

Co

rall

ino

ph

yci

dae*

50

00

00

00

00

.17

00

Ph

ym

ato

lith

on

51

00

00

00

00

00

Nem

alio

ph

yci

dae

Aud

oui

nel

la5

20

00

00

00

00

0

Co

laco

nem

a5

30

00

00

00

00

0

Rh

ody

men

iop

hy

cida

eC

ho

ndr

us5

40

00

00

00

0.1

70

0

Del

esse

ria

55

00

00

00

00

00

Har

vey

ella

56

00

00

00

00

00

Hy

po

glo

ssum

57

00

00

00

00

00

Mas

toca

rpus

58

00

00

00

00

00

Pey

sso

nn

elia

59

00

00

00

00

00

Cen

tro

hel

ida

Cen

tro

hel

ida*

Cen

tro

hel

ida*

Cen

tro

hel

ida*

60

0.4

80

.54

00

00

0.8

30

.14

0.3

10

.34

Aca

nth

ocy

stid

aeA

can

tho

cyst

idae

*C

ho

ano

cyst

is6

10

.81

0.4

61

.01

1.2

0.2

30

.13

1.4

40

.14

1.0

81

.29

Pte

rocy

stis

62

0.2

60

00

00

0.1

60

00

Rai

ner

iop

hry

s6

30

.08

00

.49

00

00

00

0

Cen

tro

hel

ida*

Cen

tro

hel

ida*

Cen

tro

hel

ida_

H1

5-6

*6

40

00

00

00

00

0

Cen

tro

hel

ida*

Cen

tro

hel

ida_

H2

6-1

*6

50

00

00

00

00

0

Het

ero

ph

ryid

aeH

eter

op

hry

idae

*O

xn

erel

la6

70

00

00

00

00

0

Cen

tro

hel

ida*

Cen

tro

hel

ida*

Cen

tro

hel

ida_

M1

-18

D0

8*

68

00

00

00

00

00

SUPPLEMENT CHAPTER I

Table S8: continued

123

Cry

pto

ph

yce

aeC

ryp

top

hy

ceae

*C

ryp

top

hy

ceae

*G

on

iom

on

as7

00

00

00

00

00

0

Kat

hab

lep

har

idae

Kat

hab

lep

har

idae

*H

aten

a7

10

.48

0.1

40

.48

0.3

50

01

.44

0.9

40

.35

0.3

3

Kat

able

ph

aris

72

00

00

00

00

00

Leu

cocr

yp

tos

73

0.0

80

00

00

0.1

40

0.1

10

Kat

hab

lep

har

idae

*7

40

.08

0.1

40

0.1

10

00

0.1

40

.13

0

Dis

coba

Jak

obi

daJa

ko

bida

*Ja

ko

bida

*7

50

.25

00

00

00

00

0

Jak

obi

da_

RM

1-S

GM

49

*7

60

00

00

00

00

0

Jak

obi

da*

77

00

0.1

70

00

00

.14

00

Euk

ary

ota

`*A

ncy

rom

on

adid

aA

ncy

rom

on

adid

a*A

ncy

rom

on

as7

90

.81

00

0.3

50

00

00

.10

.82

Ap

uso

mo

nad

idae

An

cyro

mo

nad

ida*

80

0.8

11

.74

1.8

81

.22

.31

0.8

91

.44

1.6

61

.08

1.2

9

Am

asti

gom

on

as8

10

.81

0.1

71

.88

1.2

0.1

81

.47

1.4

40

.95

1.0

81

.29

Ap

uso

mo

nas

82

00

0.2

30

00

0.4

50

0.1

0.1

1

Ap

uso

mo

nad

idae

*F

abo

mo

nas

83

0.0

80

0.2

00

00

00

0

Bre

via

tea

Bre

via

tea

Subu

lato

mo

nas

84

00

.19

00

00

0.1

30

.14

00

Euk

ary

ota

`*E

ukar

yo

ta`*

Man

tam

on

as8

50

00

00

00

00

0

Rig

ifil

ida

Rig

ifil

ida*

Mic

ron

ucle

aria

86

00

00

.12

00

0.1

30

00

Euk

ary

ota

`*E

ukar

yo

ta`*

Tel

on

ema

87

0.5

00

.17

0.3

50

.59

00

0.1

70

.10

.4

Op

isth

ok

on

ta*

Fre

shw

ater

Op

isth

ok

on

ta*

Fre

shw

ater

Op

isth

ok

on

ta*

Fre

shw

ater

Op

isth

ok

on

ta*

88

0.4

90

.14

0.1

70

.12

00

0.1

30

0.3

10

.72

Ho

lozo

a*A

can

tho

ecid

aA

can

tho

ecid

a*8

90

.48

0.1

50

.16

0.1

20

00

.92

0.1

40

.31

0.3

4

Aca

nth

oec

idae

*9

00

.81

0.1

60

.17

0.1

20

00

.45

0.1

40

0.1

Aca

nth

oec

a9

10

.81

0.1

60

.53

0.3

60

00

.40

.14

0.1

0.3

3

Cal

liac

anth

a9

20

.81

00

.17

1.2

00

0.1

30

.13

0.3

10

.34

Aca

nth

oec

idae

_F

V2

3-1

A4

*9

30

.81

00

.17

00

00

.13

0.1

40

0.1

Ho

lozo

a#Sa

vil

lea

94

0.8

10

0.2

00

00

.16

0.1

40

0.3

4

Dia

ph

ano

eca

95

0.4

80

00

00

.13

0.4

30

00

Mar

ine

Ch

oan

ofl

agel

late

s 1

96

0.8

11

.74

1.0

11

.21

.32

0.7

90

.83

1.6

61

.08

1.2

9

Step

han

oec

a9

70

00

00

00

00

0

Ho

lozo

a_C

lade

L*

Ho

lozo

a_C

lade

L*

98

0.8

10

.46

1.8

81

.20

0.3

81

.44

0.9

51

.08

1.2

9

Cra

sped

ida

Mo

no

siga

99

0.0

80

00

.11

00

00

0.1

0

Fre

shw

ater

Ch

oan

ofl

agel

late

s 1

10

00

00

00

00

00

0

Ho

lozo

a*L

agen

oec

a1

01

0.8

10

.97

1.8

81

.21

.39

0.1

31

.44

0.9

51

.08

0.7

4

SUPPLEMENT CHAPTER I

Table S8: continued

124

Cra

sped

ida_

OL

I11

04

1*

10

20

00

00

00

00

0

Cra

sped

ida

Salp

ingo

ecid

ae*

10

30

.48

0.1

70

00

0.1

30

00

.31

0.3

9

Ch

oan

oec

a1

04

0.0

80

00

00

00

00

Fre

shw

ater

Ch

oan

ofl

agel

late

s 2

10

50

0.1

70

0.7

00

.47

00

.49

0.1

0.7

4

Salp

ingo

eca

10

60

00

00

.23

0.1

30

0.1

90

0.3

4

Co

rall

och

ytr

eaC

ora

llo

chy

trid

aC

ora

llo

chy

triu

m1

07

00

00

00

00

00

Ho

lozo

a#H

olo

zoa*

Fil

aste

rea

Mar

ine

Gro

up*

10

80

.48

0.1

50

.17

0.1

30

.76

0.1

30

.83

0.9

50

.66

0.7

4

Min

iste

ria

10

90

.81

0.5

0.5

90

.70

.23

00

.17

0.1

90

.35

0.7

4

Ich

thy

osp

ore

aD

erm

ocy

stid

aR

hin

osp

ori

deac

ae*

11

00

.08

00

00

00

00

0

Ich

thy

op

ho

nae

Abe

ofo

rmid

ae*

11

10

00

00

00

00

0

Abe

ofo

rma

11

20

00

00

00

00

0

Mar

ine

Ich

thy

osp

ore

ans

11

13

00

00

00

00

00

Pir

um1

14

0.2

40

.60

00

.18

0.7

90

0.5

90

.11

0.3

4

Ich

thy

op

ho

nus

11

50

00

00

00

00

0

Cre

oli

max

11

70

00

00

00

00

0

Pse

udo

per

kin

sus

11

80

00

00

00

00

0

An

nel

ida

An

nel

ida`

*F

auv

elo

psi

s1

19

00

00

00

00

00

Po

lych

aeta

Eun

icid

a*1

20

00

00

00

00

00

Ph

yll

odo

cida

*1

22

00

00

00

00

00

Sabe

llid

a*1

23

00

00

00

00

00

Cap

itel

lida

*1

24

0.4

90

.17

00

00

00

00

Co

ssur

idae

*1

25

00

00

00

00

00

Met

azo

a

(An

imal

ia)

Op

hel

iida

e*1

26

0.8

10

0.2

31

.21

.31

.47

1.4

41

.66

1.0

81

.29

Spio

nid

a*1

27

00

00

00

00

00

Ter

ebel

lida

*1

28

0.5

00

0.1

10

00

0.1

90

0

Ara

chn

ida

Ara

chn

ida*

13

00

00

00

00

00

.10

Art

hro

po

daO

pil

ion

es*

13

10

00

00

00

00

0

Max

illo

po

daM

axil

lop

oda

*1

32

00

00

00

00

00

Cal

ano

ida*

13

30

00

00

00

00

0

Har

pac

tico

ida*

13

40

.81

1.7

41

.12

1.2

1.4

80

.93

0.9

20

.94

1.0

81

.29

Art

hro

po

daM

axil

lop

oda

Mis

op

hri

oid

a*1

35

00

00

00

00

00

SUPPLEMENT CHAPTER I

Table S8: continued

125

Sip

ho

no

sto

mat

oid

a*1

36

00

00

.12

00

00

00

Sess

ilia

*1

37

00

00

00

00

00

Bry

ozo

aG

ym

no

laem

ata

Cte

no

sto

mat

ida*

13

90

00

00

00

00

0

Tun

icat

aA

scid

iace

aE

nte

rogo

na*

14

00

.81

0.5

0.1

70

.73

1.3

0.7

90

.92

0.9

61

.08

0.7

4

Sto

lido

bran

chia

*1

41

0.2

50

.14

1.0

10

0.2

60

.14

0.1

70

.44

00

.42

Ver

tebr

ata

Act

ino

pte

rygi

iA

ctin

op

tery

gii*

14

20

0.1

40

00

00

00

0

Mam

mal

iaM

amm

alia

*1

43

0.8

11

.74

1.8

81

.22

.31

1.4

71

.44

1.6

61

.08

1.2

9

Ech

ino

derm

ata

Ast

ero

idea

Ast

ero

idea

*1

44

0.2

50

00

00

00

1.0

80

Cri

no

idea

Cri

no

idea

*1

45

00

00

00

00

00

Op

hiu

roid

eaO

ph

iuro

idea

*1

47

00

00

00

00

00

En

top

roct

aSo

lita

ria

Lo

xo

som

atid

ae*

14

80

.25

0.2

10

.54

1.2

1.4

80

.89

0.1

30

.48

0.1

0.7

4

Met

azo

a

(An

imal

ia)

Gas

tro

tric

ha

Gas

tro

tric

ha*

Ch

aeto

no

tida

*1

49

0.2

51

.74

1.8

80

.71

.32

0.9

1.4

41

.66

0.3

51

.29

Mo

llus

caG

astr

op

oda

Cae

no

gast

rop

oda

*1

51

00

00

00

00

00

Ner

itim

orp

ha*

15

20

00

00

00

00

0

Nem

ato

daC

hro

mad

ore

aA

raeo

laim

ida*

15

30

00

00

00

00

0

Ch

rom

ado

rida

*1

54

0.5

1.7

40

.54

0.1

20

.21

.47

0.4

60

.95

0.6

30

.86

Mo

nh

yst

erid

a*1

55

0.2

50

.97

0.6

10

.12

0.6

90

.38

0.1

11

.66

0.6

30

.33

En

op

lea

En

op

lida

*1

56

0.2

40

.17

0.2

0.3

60

.20

.49

0.1

10

.13

0.1

0.7

2

An

op

laP

aleo

nem

erte

a*1

58

00

00

00

00

00

Pla

tyh

elm

inth

esR

hab

dito

ph

ora

Rh

abdo

coel

a*1

59

00

00

00

00

00

Ro

tife

raM

on

ogo

no

nta

Flo

scul

aria

cea*

16

00

00

00

00

00

0

Plo

imid

a*1

61

00

0.5

90

.11

0.2

50

0.1

40

.95

00

.12

Pri

apul

ida

Pri

apul

ida*

Tub

iluc

hid

ae*

16

20

00

00

00

00

0

Met

azo

aun

id*

Met

azo

aun

id_

Cla

ssM

etaz

oa*

16

30

0.5

00

00

.79

00

0.1

0

Cn

idar

iaA

nth

ozo

aA

ctin

iari

a*1

64

0.2

40

00

00

00

0.1

10

An

tip

ath

aria

*1

65

00

00

00

00

0.1

30

Co

rall

imo

rph

aria

*1

66

00

00

00

00

00

Zo

anth

aria

*1

67

0.0

80

00

00

00

00

.12

Hy

dro

zoa

An

tho

ath

ecat

a*1

69

00

00

00

0.1

10

.45

00

Lep

toth

ecat

a*1

70

00

00

00

00

00

Hy

dro

zoa*

17

10

00

00

00

00

0

SUPPLEMENT CHAPTER I

Table S8: continued

126

Scy

ph

ozo

aR

hiz

ost

om

eae*

17

40

.24

00

.20

.11

00

0.1

10

0.1

10

Scy

ph

ozo

aSe

mae

ost

om

eae*

17

50

.24

0.1

70

.17

0.3

80

00

.37

0.4

80

.11

0.3

4

Stau

rom

edus

ae*

17

60

.81

1.0

41

.88

1.2

1.4

61

.47

1.4

41

.66

1.0

81

.29

Cte

no

ph

ora

Cy

clo

coel

aB

ero

ida*

17

70

00

00

00

00

0

Lo

bata

*1

78

00

00

00

00

00

Po

rife

raC

alca

rea

Cal

care

a*1

80

00

.19

1.2

0.3

50

00

.38

00

.33

0.3

3

Bae

rida

*1

81

0.8

11

.01

1.2

1.2

0.2

11

.47

1.4

40

.96

1.0

81

.29

Leu

coso

len

ida*

18

20

.08

0.1

41

.20

.35

0.2

10

.38

0.4

0.1

40

.11

0.1

Met

azo

a

(An

imal

ia)

Lit

ho

nid

a*1

83

0.0

80

00

.35

00

0.1

10

0.1

0.1

Cla

thri

nid

a*1

84

0.2

40

0.2

0.3

50

.21

0.1

70

.38

00

.11

0.1

Mur

ray

on

ida*

18

50

00

00

00

00

0

Dem

osp

on

giae

Den

dro

cera

tida

*1

86

0.4

81

.01

0.6

71

.20

1.4

70

.85

0.9

41

.08

0.7

4

Had

rom

erid

a*1

87

0.8

11

.74

1.1

1.2

2.3

11

.47

1.4

41

.66

1.0

81

.29

Hal

ich

on

drid

a*1

88

00

00

.13

00

0.1

30

00

Hap

losc

leri

da*

18

90

0.1

40

.19

0.1

10

0.1

70

.14

0.1

70

0

Po

ecil

osc

leri

da*

19

00

.51

.08

1.1

0.7

41

.46

0.9

10

.38

0.9

40

.30

.39

Spir

op

ho

rida

*1

91

00

00

00

00

00

.11

Ver

on

gida

*1

92

00

00

00

00

0.1

0

Nuc

letm

yce

a#N

ucle

tmy

cea*

Nuc

letm

yce

a*N

ucle

tmy

cea*

19

30

.81

0.1

60

.16

1.2

0.7

0.1

30

.37

0.9

40

.33

0.8

2

Dis

cicr

isto

idea

Fo

nti

culi

daF

on

ticu

lida

*F

on

ticu

lida

*1

94

0.4

91

.74

1.8

81

.20

.75

1.4

71

.44

1.6

61

.08

0.7

4

Dis

cicr

isto

idea

`*D

isci

cris

toid

ea`*

Dis

cicr

isto

idea

`*1

95

0.0

80

00

00

0.4

20

0.1

0

Ch

ytr

idio

my

cota

Ch

ytr

idio

my

cete

s`C

hy

trid

iom

yce

tes`

*1

96

00

00

00

00

00

.33

Ch

ytr

idia

les*

19

70

.50

.46

0.1

50

0.5

90

.40

.13

00

.31

0.3

3

Rh

izo

ph

ydi

um1

98

00

00

00

00

00

.1

Neo

kar

lin

gia

19

90

.25

00

00

.18

0.1

40

00

0

Rh

izo

ph

lyct

idal

es*

20

00

00

00

00

00

0

Rh

izo

ph

ydi

ales

*2

01

00

00

00

00

00

Rh

izo

ph

ydi

ales

*2

02

00

00

00

00

00

Spiz

ello

my

ceta

les*

20

30

.25

00

.17

00

00

00

0

Po

wel

lom

yce

s2

04

00

00

00

00

00

Tri

par

tica

lcar

20

60

00

00

0.1

30

00

0

SUPPLEMENT CHAPTER I

Table S8: continued

127

Ch

ytr

idio

my

cete

s`*

20

70

.50

.46

0.1

70

0.6

40

.91

0.3

70

.14

0.1

30

.72

Mo

no

blep

har

ido

my

cete

s`M

on

obl

eph

arid

ales

*2

08

0.5

00

00

0.1

30

00

0

Hy

alo

rap

hid

ium

20

90

00

00

00

00

0

Cry

pto

my

cota

Cry

pto

my

cota

`*R

oze

lla

21

00

.24

0.4

60

00

.18

0.4

00

00

.16

Cry

pto

my

cota

*C

ryp

tom

yco

ta*

21

10

.25

00

0.1

20

0.1

30

0.5

30

.10

.11

Asc

om

yco

taD

oth

ideo

my

cete

sG

uign

ardi

a2

12

00

00

00

00

00

Cap

no

dial

es*

21

30

00

00

00

00

0

Co

cco

din

ium

21

40

00

00

00

00

0

Cla

dosp

ori

um2

15

00

00

00

00

00

My

cosp

hae

rell

a2

16

00

00

00

00

00

Ple

osp

ora

les*

22

00

00

00

00

00

0

Ph

aeo

sph

aeri

a2

21

00

00

00

00

00

Fun

giE

uro

tio

my

cete

sC

hae

toth

yri

ales

*2

22

00

00

00

00

00

Eur

oti

ales

*2

23

00

.18

00

00

00

00

Asp

ergi

llus

22

40

00

00

00

00

0

Pen

icil

lium

22

50

00

00

00

00

0

Th

ysa

no

ph

ora

22

60

00

00

00

00

.10

Ver

ruca

ria

22

70

00

00

00

00

0

Lec

ano

rom

yce

tes

Lec

ano

rale

s*2

28

00

00

00

00

00

Leo

tio

my

cete

sB

otr

yo

tin

ia2

29

00

00

00

00

00

Sord

ario

my

cete

sH

yp

ocr

eale

s*2

32

0.2

40

.46

00

00

00

00

Em

eric

ello

psi

s2

34

00

.17

00

00

00

00

Geo

smit

hia

23

50

00

00

00

00

0

Sacc

har

om

yce

tes

Sacc

har

om

yce

tale

s*2

38

0.0

90

00

.34

0.1

80

.14

00

.54

00

.16

Can

dida

23

90

00

00

00

00

0

Bas

idio

my

cota

Aga

rico

my

cete

sA

gari

com

yce

tes*

24

10

.81

1.7

40

.49

0.7

1.4

81

.47

1.4

40

.13

1.0

80

.39

Leu

coag

aric

us2

42

00

00

00

00

00

Bo

lbit

ius

24

30

00

00

00

00

0

Ch

amae

ota

25

00

00

00

00

00

0

Agr

ocy

be2

51

00

00

00

.13

00

00

Psi

locy

be2

53

00

00

00

00

00

SUPPLEMENT CHAPTER I

Table S8: continued

128

Ch

rom

ose

ra2

54

00

00

00

00

00

Fun

giA

can

tho

lich

en2

56

00

00

00

00

00

Dip

lom

ito

po

rus

25

80

00

00

00

00

0

Tra

met

es2

59

00

00

00

00

00

Pen

iop

ho

ra2

60

00

00

00

00

00

Tre

mel

lom

yce

tes

Cry

pto

cocc

us2

62

00

00

00

00

00

Dio

szeg

ia2

63

00

00

00

00

00

Mic

robo

try

om

yce

tes

Spo

ridi

obo

lale

s*2

64

00

00

00

00

00

Rh

odo

toru

la2

66

00

00

00

00

00

Ex

oba

sidi

om

yce

tes

Mal

asse

zia

26

80

00

00

00

00

0

Qua

mba

lari

a2

69

00

00

00

00

00

Neo

call

imas

tigo

my

cota

Neo

call

imas

tigo

my

cete

sN

eoca

llim

asti

gace

ae*

27

00

00

00

00

00

0

Zy

gom

yco

taZ

ygo

my

cota

`*M

ort

iere

llal

es*

27

20

.50

.20

.16

0.1

20

00

00

.31

0

Fun

giun

id*

Fun

giun

id*

Fun

gi*

27

40

00

00

00

00

0.1

1

Nuc

letm

yce

a#N

ucle

tmy

cea_

LK

M1

5*

Nuc

letm

yce

a_L

KM

15

*N

ucle

tmy

cea_

LK

M1

5*

27

50

00

00

00

00

0

Euk

ary

ota

*P

ico

zoa

Pic

ozo

a*P

ico

zoa*

27

60

00

00

00

00

0

Pic

om

on

adid

aP

ico

mo

nas

27

70

00

00

00

00

0

Euk

ary

ota

*E

ukar

yo

ta*

Euk

ary

ota

*2

78

00

00

00

00

00

Alv

eola

ta*

Alv

eola

ta*

Alv

eola

ta*

27

90

0.1

40

00

00

00

0

Ap

ico

mp

lex

aC

on

oid

asid

aG

ous

sia

28

00

00

0.1

20

00

00

0

Mar

goli

siel

la2

81

0.2

40

.19

0.4

91

.20

.64

0.3

81

.44

0.1

30

.31

1.2

9

Rh

yti

docy

stis

28

20

00

00

00

00

0

Alv

eola

taSe

len

idiu

m2

83

00

.46

0.5

30

.13

00

0.1

40

.96

0.1

0.1

2

Lan

kes

teri

a2

84

00

.16

0.1

70

00

0.1

30

.17

00

Eug

rega

rin

ori

da*

28

50

00

0.1

20

00

00

0

No

vel

Ap

ico

mp

lex

a C

lass

1N

ov

el A

pic

om

ple

xa

Cla

ss 1

*2

86

0.0

80

.97

1.1

00

.18

0.4

90

.46

0.9

50

0.1

2

No

vel

Ap

ico

mp

lex

a C

lass

2N

ov

el A

pic

om

ple

xa

Cla

ss 2

*2

87

00

.17

00

00

00

00

Alv

eola

ta*

Alv

eola

ta*

Alv

eola

ta_

BO

LA

91

4*

28

80

00

00

00

00

0

Cil

iop

ho

raIn

tram

acro

nuc

leat

aSo

roge

na

29

10

00

00

00

00

0

Oli

goh

ym

eno

ph

ore

a*2

92

00

00

00

00

00

Cin

eto

chil

um2

93

00

00

00

00

0.1

0

SUPPLEMENT CHAPTER I

Table S8: continued

129

Hy

alo

ph

ysa

29

40

00

00

.69

0.1

30

0.1

40

0.1

6

Vam

py

rop

hry

a2

95

00

00

00

00

00

.16

An

op

lop

hry

a2

97

00

00

00

00

00

Par

acla

usil

oco

la3

00

00

00

00

00

00

Oli

goh

ym

eno

ph

ore

a*3

01

0.8

10

.49

0.5

91

.21

.39

0.3

80

.80

0.6

30

.1

Ap

oca

rch

esiu

m3

02

0.8

10

0.1

90

.72

00

0.4

30

0.1

0.3

3

Ast

ylo

zoo

n3

03

00

00

00

00

00

Car

ches

ium

30

40

.09

00

0.1

20

00

00

0

Ep

icar

ches

ium

30

50

.49

00

.19

0.7

40

.18

0.1

30

.42

0.1

40

.31

0.3

3

Op

hry

dium

30

70

00

00

00

00

0

Alv

eola

taO

pis

tho

nec

ta3

08

00

00

00

00

00

Pse

udo

vo

rtic

ella

30

90

.81

1.7

41

.88

1.2

2.3

10

.91

1.4

40

.95

1.0

81

.29

Tel

otr

och

idiu

m3

10

0.8

10

.49

0.5

40

.74

2.3

10

.13

0.1

30

.14

0.6

30

.33

Vag

inic

ola

31

10

.08

00

00

00

00

0

Vo

rtic

ella

31

20

.81

0.1

80

.19

1.2

00

.13

0.8

50

.43

0.6

30

.38

Zo

oth

amn

ium

31

30

.09

00

0.1

30

00

00

0

Oli

goh

ym

eno

ph

ore

a*3

14

00

00

00

00

00

Car

dio

sto

mat

ella

31

50

00

00

00

.13

0.1

40

.10

Cy

clid

ium

31

60

00

00

00

00

0

Dex

iotr

ich

a3

17

0.0

80

00

00

00

00

Dex

itri

chid

es3

18

00

00

00

00

00

Gla

uco

nem

a3

19

0.2

40

.57

00

.35

00

.16

0.1

60

.17

0.3

0.1

2

Hip

po

com

os

32

00

00

00

00

00

0

Ho

mal

oga

stra

32

10

.08

00

00

00

00

.10

Mes

ano

ph

rys

32

20

00

00

00

00

0

Mia

mie

nsi

s3

23

0.0

80

00

00

00

0.1

0.3

3

Par

ano

ph

rys

32

40

1.0

10

.19

0.1

10

.69

00

0.1

90

.33

0.4

Ph

ilas

ter

32

50

00

00

00

00

0

Pla

gio

py

liel

la3

26

00

.17

00

00

00

00

Pse

udo

coh

nil

embu

s3

27

00

00

00

00

00

Uro

nem

a3

28

00

00

00

00

00

SUPPLEMENT CHAPTER I

Table S8: continued

130

Wil

bert

ia3

29

00

00

00

00

00

Oli

goh

ym

eno

ph

ore

a*3

30

01

.04

00

00

0.1

60

00

Oli

goh

ym

eno

ph

ore

a*3

31

00

00

00

00

0.1

10

Ap

ort

ho

tro

chil

ia3

32

00

00

00

00

00

Dy

ster

ia3

33

0.8

11

.74

0.4

80

.70

.63

0.9

31

.44

1.6

60

.68

0.7

4

Har

tman

nul

a3

34

00

00

00

00

00

Het

ero

har

tman

nul

a3

35

00

.14

00

00

00

00

Pit

hit

es3

36

00

00

00

00

00

Alv

eola

taT

rich

op

odi

ella

33

70

00

00

00

.11

00

0

Tro

chil

ia3

38

00

.21

00

00

00

00

Tro

chil

ioid

es3

39

0.2

50

.51

.01

1.2

00

.13

0.1

41

.66

0.3

10

.74

Aci

net

a3

40

0.8

11

.74

1.8

81

.22

.31

1.4

71

.44

1.6

61

.08

1.2

9

Ep

alx

ella

34

10

00

.17

00

00

00

0

Lit

ost

om

atea

*3

43

00

00

00

00

00

Ep

iph

yll

um3

46

0.0

80

00

00

0.1

40

0.1

0

Hem

iop

hry

s3

47

00

00

.12

00

0.1

30

0.1

10

Lit

on

otu

s3

48

00

00

00

00

0.1

0

Lo

xo

ph

yll

um3

49

0.0

90

00

00

00

00

Spir

otr

ich

ea*

35

00

.48

0.1

41

.10

.70

.59

0.1

30

.92

1.6

60

.11

.29

Ch

ore

otr

ich

ia*

35

10

00

.19

00

00

00

0

Asp

idis

ca3

52

0.0

80

.14

0.2

30

.35

0.2

30

0.3

80

.17

0.6

60

.1

Dis

coce

ph

alus

35

30

00

00

00

00

0

Eup

lote

s3

54

00

00

00

00

00

Hy

po

tric

hia

*3

55

0.4

80

.49

00

.73

1.3

90

.41

00

0.6

30

.38

An

teh

olo

stic

ha

35

60

.48

00

00

00

00

0.1

1

Go

no

sto

mum

35

80

.23

0.1

80

00

.22

00

.13

00

.10

.11

Hal

teri

a3

59

0.2

30

.18

00

00

00

00

Ho

lost

ich

a3

60

00

.17

0.1

50

.35

00

0.4

50

00

.12

Ort

ham

ph

isie

lla

36

20

.23

00

00

00

.13

00

.10

.11

Ox

ytr

ich

a3

63

00

00

0.2

20

00

00

Par

abis

tich

ella

36

40

0.1

80

00

00

00

0

SUPPLEMENT CHAPTER I

Table S8: continued

131

Par

aste

rkie

lla

36

50

00

00

00

00

0

Ple

uro

tric

ha

36

60

00

00

00

00

0

Po

ntu

rost

yla

36

70

00

00

00

00

0

Psa

mm

om

itra

36

80

00

00

00

00

0

Spir

otr

ach

elo

sty

la3

70

00

00

00

00

00

Alv

eola

taSt

ylo

ny

chia

37

10

00

00

00

00

0

Th

igm

ok

ero

no

psi

s3

72

0.0

80

00

00

00

00

Uro

lep

tus

37

40

00

00

00

00

0

Hy

po

tric

hia

*3

75

0.2

30

00

.11

00

00

0.1

0

Lic

no

ph

ora

37

60

00

00

00

00

0

Oli

gotr

ich

ia*

37

70

00

00

00

00

0

Pse

udo

ton

ton

ia3

78

0.0

80

00

00

00

00

Stro

mbi

dium

37

90

.25

00

.15

00

00

00

.10

Var

istr

om

bidi

um3

80

00

00

00

00

00

Oli

gotr

ich

ia*

38

10

00

00

00

00

.10

Ox

ytr

ich

idae

*3

82

0.4

80

0.4

80

.34

0.1

80

0.4

40

0.1

0

Pro

tocr

uzia

38

30

00

00

00

00

0

Po

stci

lio

desm

ato

ph

ora

Euf

oll

icul

ina

38

40

00

00

00

.92

00

0

Per

itro

mus

38

50

0.1

80

00

00

.13

00

.11

0.1

Din

ofl

agel

lata

*D

ino

flag

ella

ta*

38

60

00

00

00

00

0

Din

op

hy

ceae

Din

op

hy

ceae

*3

87

0.2

41

.01

0.4

91

.20

.21

.47

0.1

10

1.0

81

.29

Din

op

hy

ceae

_B

AQ

K0

3*

38

80

.24

0.9

70

.15

0.7

0.6

31

.47

0.1

10

.45

0.6

30

.72

Din

ofl

agel

lata

Din

op

hy

ceae

_C

CM

P1

87

8*

38

90

00

00

00

00

.31

0

Din

op

hy

ceae

_D

24

4*

39

00

00

00

00

00

0

Din

op

hy

sis

39

20

00

00

00

00

0

Sin

op

hy

sis

39

30

.25

0.9

71

.20

.72

0.2

20

.41

0.4

0.4

40

.66

1.2

9

Am

ph

idin

ium

39

40

00

00

00

.14

00

0

Gy

mn

odi

niu

m c

lade

*3

95

00

.17

00

00

00

00

Ch

ytr

iodi

niu

m3

96

0.0

80

.46

00

00

00

00

Ery

thro

psi

din

ium

39

70

.09

00

00

00

00

.31

0

Gy

mn

odi

niu

m c

lade

_F

V1

8-2

D9

*3

98

0.0

80

00

00

00

0.3

10

SUPPLEMENT CHAPTER I

Table S8: continued

132

Gy

mn

odi

niu

m4

00

0.8

10

.14

00

.12

00

.39

0.1

30

.14

0.6

30

.34

Lep

ido

din

ium

40

10

.09

00

0.1

20

00

00

0.1

3

Nem

ato

din

ium

40

20

00

00

00

00

0

Par

agy

mn

odi

niu

m4

03

00

00

00

.14

00

0.3

10

Spin

ifer

odi

niu

m4

04

00

00

.11

00

00

0.3

30

Gy

rodi

niu

m4

05

0.8

10

.46

0.1

50

.70

.22

0.4

10

.40

.48

1.0

80

.74

Ap

ico

po

rus

40

60

00

00

00

00

0

Co

chlo

din

ium

40

70

00

00

00

00

0

Alv

eola

taD

ino

flag

ella

taK

aren

ia4

08

00

00

00

00

00

Kar

lodi

niu

m4

09

0.4

80

.97

00

.36

0.6

30

.79

00

.13

0.3

0.3

3

Gy

mn

odi

nip

hy

cida

e_SC

M2

7C

9*

41

10

.08

0.1

50

00

00

.11

00

.11

0.1

2

Sues

siac

eae*

41

20

00

00

00

00

0

Sues

siac

eae_

3b-

C9

*4

13

00

00

00

00

00

Bie

chel

eria

41

40

00

0.1

10

0.1

30

00

0

Pel

ago

din

ium

41

50

.08

0.1

90

0.1

10

0.1

40

.11

0.1

40

.66

0.3

3

Po

lare

lla

41

60

00

00

00

00

0

Pro

todi

niu

m4

17

00

00

00

00

0.1

0

Sym

bio

din

ium

41

80

00

00

00

00

0

Aza

din

ium

41

90

.81

0.9

70

.49

0.7

1.3

91

.47

0.3

70

.95

0.6

30

.72

Din

op

hy

ceae

_N

IF-4

G9

*4

20

00

00

00

00

00

Py

rodi

niu

m4

21

00

00

00

00

00

Am

ph

idin

iop

sis

42

30

00

00

00

00

0

Cry

pth

eco

din

ium

42

40

0.2

10

00

.30

00

00

Gle

no

din

ium

42

50

00

00

00

00

0

Het

ero

cap

sa4

26

0.8

11

.74

1.8

81

.22

.31

1.4

70

.80

.95

1.0

81

.29

Isla

ndi

niu

m4

27

00

.15

00

00

0.1

40

00

Per

idin

iop

sis

42

80

00

00

00

00

0

Per

idin

ium

42

90

.81

0.9

70

.49

0.3

60

.75

1.4

70

.83

0.9

60

.31

0.7

4

Pro

top

erid

iniu

m4

30

0.0

80

00

0.7

60

.14

00

00

Po

dola

mp

adac

eae*

43

10

0.1

40

00

.30

00

00

.1

Les

sard

ia4

32

00

00

00

00

00

SUPPLEMENT CHAPTER I

Table S8: continued

133

Th

ora

cosp

hae

race

ae*

43

40

0.1

60

0.1

20

00

00

.35

0.1

2

Cry

pto

per

idin

iop

sis

43

50

.08

00

00

00

0.1

30

0

Din

ofl

agel

lata

Oo

din

ium

43

60

.23

00

.16

0.3

50

.68

0.1

30

.13

0.1

30

.31

0.1

1

Pfi

este

ria

43

70

.08

0.1

60

00

.22

00

0.5

40

0

Scri

pp

siel

la4

38

0.0

80

00

.12

00

00

0.3

0

Th

ora

cosp

hae

ra4

39

00

00

00

00

00

Th

ora

cosp

hae

race

ae*

44

00

.08

00

00

00

00

0

Alv

eola

taE

xuv

iael

la4

41

00

00

00

00

0.1

0

Pro

roce

ntr

um4

42

0.0

90

.15

00

.11

0.2

31

.47

00

.43

0.6

60

.12

Din

op

hy

ceae

_SC

M3

7C

58

*4

44

00

00

00

00

00

Din

op

hy

ceae

_SL

16

3A

10

*4

46

0.0

80

00

0.2

20

0.1

30

0.1

0

Din

op

hy

ceae

_cL

A1

1G

01

*4

47

0.0

80

00

.12

00

00

00

Din

op

hy

ceae

*4

48

0.2

50

.16

0.1

50

00

0.1

10

.13

00

.1

Din

ofl

agel

lata

`*B

last

odi

niu

m4

49

0.0

80

.14

00

0.6

30

.13

00

00

Pau

lsen

ella

45

00

00

0.1

20

00

00

0

Din

ofl

agel

lata

*D

ino

flag

ella

ta_

S9*

45

10

.08

00

00

00

00

.11

0

Din

ofl

agel

lata

_SC

M2

8C

5*

45

30

00

00

00

00

0

Alv

eola

ta*

Alv

eola

ta*

Alv

eola

ta_

H6

7*

45

40

.08

00

0.1

20

00

00

.10

Alv

eola

ta_

NIF

-4C

10

*4

56

0.8

11

.74

1.0

11

.20

.59

1.4

70

.80

.44

0.6

30

.74

Alv

eola

ta_

OL

I11

25

5*

45

70

.23

0.1

40

.15

0.1

10

00

.14

00

.30

Pro

talv

eola

taC

hro

mer

ida

Ch

rom

era

45

80

.09

00

00

00

00

0

Co

lpo

dell

ida

Co

lpo

dell

ida*

45

90

00

00

00

0.4

80

0

Co

lpo

dell

a4

60

00

00

.12

00

00

00

Pro

talv

eola

ta*

Ox

yrr

his

46

10

00

00

00

00

0

Per

kin

sida

eP

erk

insi

dae_

A3

1*

46

20

.08

00

00

00

.18

00

0.1

1

Par

vil

ucif

era

46

30

00

00

00

00

0

Syn

din

iale

sSy

ndi

nia

les*

46

40

.48

0.1

60

.15

1.2

1.3

91

.47

0.3

70

.45

0.6

30

.72

Am

oeb

op

hry

a4

65

00

00

00

00

0.1

0

Alv

eola

taD

ubo

scqu

ella

46

60

00

00

00

.14

00

0

Syn

din

iale

s_G

roup

I*

46

80

.49

0.4

60

.48

1.2

2.3

11

.47

0.8

51

.66

1.0

80

.74

Syn

din

iale

s_G

roup

II*

46

90

00

00

0.1

30

0.1

40

0.1

SUPPLEMENT CHAPTER I

Table S8: continued

134

Alv

eola

ta*

Alv

eola

ta*

Alv

eola

ta_

SCM

37

C5

2*

47

00

00

00

00

00

0

Alv

eola

ta_

SGU

H9

42

*4

71

00

00

00

00

00

Cer

cozo

aC

erco

zoa*

Cer

cozo

a*4

72

0.4

80

00

00

0.1

30

0.3

30

.12

Cer

cozo

a_7

-2.3

*4

73

0.8

10

00

.12

00

00

0.3

10

.12

Cer

cozo

a_7

-5.4

*4

74

0.0

90

0.1

70

00

.13

00

00

Cer

cozo

a_B

asal

Gro

up T

*4

75

00

0.2

40

.13

00

0.1

60

00

Cer

cozo

a_C

CW

10

*4

76

0.8

11

.04

1.0

11

.21

.30

.93

1.4

40

.17

1.0

80

.72

Cer

com

on

adid

aeC

erco

mo

nas

47

80

.24

00

00

00

00

0

Ch

lora

rach

nio

ph

yta

Gy

mn

och

lora

47

90

.25

00

00

00

00

0

Ch

lora

rach

nio

ph

yta

*4

80

0.8

11

.74

1.8

81

.21

.32

1.4

71

.44

1.6

61

.08

1.2

9

Gli

sso

mo

nad

ida

Gli

sso

mo

nad

ida*

48

10

.81

00

.16

0.1

10

00

.83

00

.31

0

Bo

dom

orp

ha

48

20

00

00

00

00

0

Het

ero

mit

a4

83

00

00

00

00

00

Rh

izar

iaG

ran

ofi

lose

aM

assi

ster

ia4

84

0.8

10

.19

0.1

70

.72

0.6

90

0.9

20

.49

0.6

30

.33

Imbr

icat

eaM

arim

on

adid

a*4

85

00

00

00

00

00

Aur

anti

cord

is4

86

0.0

80

0.1

70

00

.14

0.5

10

0.1

0

Mar

imo

nad

ida_

NA

MA

KO

-15

*4

87

0.0

90

.16

00

.12

0.3

0.4

00

0.1

10

.11

Pse

udo

pir

son

ia4

88

0.5

0.1

71

.01

1.2

1.3

91

.47

1.4

40

.45

0.6

30

.72

Imbr

icat

ea_

No

vel

Cla

de 3

*4

89

00

00

00

00

00

Nud

ifil

a4

90

00

00

00

00

00

Eug

lyp

hid

a*4

91

00

00

00

00

00

.16

Eug

lyp

hid

a_1

3-1

.8*

49

20

00

00

00

00

0

Eug

lyp

hid

a*4

93

0.0

80

00

00

0.4

50

.13

0.1

0

Cy

ph

ode

ria

49

40

00

00

00

00

0

Eug

lyp

ha

49

50

00

00

00

00

0

Pau

lin

ella

49

60

.48

01

.12

0.7

00

.90

.92

0.1

40

.30

Th

aum

ato

mo

nad

ida_

D2

P0

4A

09

*4

97

0.0

80

00

.12

00

00

0.1

0

Esq

uam

ula

49

80

00

00

00

00

0

Gy

rom

itus

49

90

.51

.01

0.4

80

.72

0.6

30

1.4

40

.94

1.0

81

.29

Th

aum

ato

mo

nad

ida*

50

00

00

0.3

50

00

.14

00

0.3

3

Th

aum

ato

mo

nad

ida_

D6

*5

01

00

0.1

60

00

0.1

30

0.3

10

SUPPLEMENT CHAPTER I

Table S8: continued

135

Th

aum

ato

mas

tix

50

20

.08

00

.20

.12

00

00

.14

0.6

30

.1

Spo

ngo

mo

nas

50

40

00

00

00

00

0

Imbr

icat

ea_

p1

5D

09

*5

05

0.8

10

0.1

50

00

00

0.3

30

.74

Cer

cozo

a`*

Gy

mn

op

hry

s5

06

0.2

40

00

.70

00

00

0

Met

rom

on

adea

Met

op

ion

50

70

.25

00

.17

0.1

20

.18

0.1

40

.37

0.4

90

.31

0.1

3

Mic

rom

eto

pio

n5

08

00

00

00

00

00

Cer

cozo

a*C

erco

zoa_

N-P

or*

50

90

.49

0.2

0.2

0.1

10

00

.17

00

.33

0.1

5

Cer

cozo

a_N

ov

el C

lade

12

*5

10

00

00

00

00

00

Rh

izar

iaC

erco

zoa_

No

vel

Cla

de 2

*5

11

0.4

80

.16

00

.70

0.4

70

.38

0.1

71

.08

0.7

4

Cer

cozo

a_N

ov

el C

lade

Gra

n-1

*5

12

0.4

80

.19

00

.11

00

.38

00

.20

.35

0.4

Cer

cozo

a_N

ov

el C

lade

Gra

n-3

*5

13

0.8

10

.50

.19

0.7

30

0.4

11

.44

0.1

41

.08

0.3

3

Cer

cozo

a_N

ov

el C

lade

Gra

n-4

*5

14

0.8

10

.46

1.1

20

.35

0.2

30

0.9

20

.45

0.3

30

.1

Cer

cozo

a_N

ov

el C

lade

Gra

n-5

*5

15

00

00

00

00

00

Cer

cozo

a_N

ov

el C

lade

Gra

n-6

*5

16

0.2

30

0.1

60

00

00

00

Ph

yto

my

xea

Ph

yto

my

xea

*5

17

0.8

11

.04

00

.34

00

.91

0.1

40

.48

0.6

30

.72

Cer

cozo

a_R

M2

-SG

M5

8*

Cer

cozo

a_R

M2

-SG

M5

8*

51

80

00

00

00

00

0

Th

eco

filo

sea

Th

eco

filo

sea*

51

90

00

00

00

00

0

Th

eco

filo

sea_

BO

LA

32

2*

52

00

.49

00

.48

0.7

0.1

80

.40

.40

0.3

10

.1

Cry

oth

eco

mo

nas

52

10

00

00

.76

0.1

70

0.1

40

.31

0

Pro

tasp

is5

22

0.8

11

.74

1.8

81

.22

.31

1.4

71

.44

1.6

61

.08

1.2

9

Rh

ogo

sto

ma

52

30

00

00

00

00

0

Th

eco

filo

sea_

DSG

M-5

0*

52

40

00

00

00

00

0

Ebr

ia*

52

50

.49

01

.10

.12

00

.79

0.9

20

.19

0.6

30

.1

Th

eco

filo

sea_

NIF

-3A

7*

52

60

.08

00

.20

.13

00

00

00

Th

eco

filo

sea_

NW

61

7*

52

70

.81

0.5

21

.88

0.7

01

.47

0.3

81

.66

0.6

30

.74

Th

eco

filo

sea_

WH

OI-

LI1

-14

*5

28

00

00

0.1

80

.13

00

0.1

0.1

3

Rh

izar

iaT

hec

ofi

lose

a*5

29

0.5

0.4

61

.01

0.7

0.6

80

.13

0.3

70

.44

1.0

80

.74

Vam

py

rell

idae

Vam

py

rell

idae

*5

30

0.8

11

.74

1.8

81

.21

.39

1.4

71

.44

1.6

61

.08

1.2

9

Cer

cozo

a*C

erco

zoa*

53

10

.81

0.2

0.5

91

.20

01

.44

0.1

41

.08

1.2

9

Ret

aria

Ret

aria

_R

AD

A*

Ret

aria

_R

AD

A*

53

20

.08

0.1

70

00

00

00

0.1

1

SUPPLEMENT CHAPTER I

Table S8: continued

136

Bic

oso

ecid

aB

ico

soec

ida*

Bic

oso

eca

53

40

.81

1.7

41

.88

1.2

2.3

10

.89

1.4

41

.66

1.0

81

.29

Pse

udo

bodo

53

60

.09

00

00

00

00

0

Ric

tus

53

70

.24

0.1

90

.15

0.3

60

00

.14

00

.66

0.3

3

Bic

oso

ecid

a*5

38

0.2

60

.16

00

.12

00

00

0.1

10

.33

Hy

ph

och

ytr

iom

yce

tes

Hy

ph

och

ytr

iom

yce

tes*

Hy

ph

och

ytr

iale

s*5

39

0.2

41

.08

0.6

10

.73

0.6

80

.79

0.8

31

.66

1.0

81

.29

Hy

ph

och

ytr

ium

54

00

00

00

0.1

30

00

0

Rh

izid

iom

yce

s5

41

0.0

80

00

.11

0.2

0.4

00

0.3

10

.11

Stra

men

op

iles

`*St

ram

eno

pil

es`*

Dev

elo

pay

ella

54

20

.49

00

.59

0.3

50

00

.16

00

.11

0.1

2

Pir

son

ia5

43

0.8

10

.97

0.4

81

.20

.18

1.4

70

.85

1.6

61

.08

1.2

9

Lab

yri

nth

ulo

my

cete

sL

aby

rin

thul

om

yce

tes*

Lab

yri

nth

ulo

my

cete

s_A

B3

F1

4R

J3E

10

*5

44

00

00

00

00

00

Lab

yri

nth

ulo

my

cete

s_A

I5F

15

RM

1E

10

*5

45

00

00

00

00

00

Lab

yri

nth

ulo

my

cete

s_D

2P

04

F0

1*

54

60

.08

00

.17

00

00

00

.10

Stra

men

op

iles

Lab

yri

nth

ulo

my

cete

s_D

52

*5

47

00

00

00

00

00

Lab

yri

nth

ulo

my

cete

s_H

E0

01

00

5.1

12

*5

48

0.0

80

0.1

70

00

00

00

Lab

yri

nth

ulac

eae

Lab

yri

nth

ula

54

90

00

00

00

00

.10

Lab

yri

nth

ulo

my

cete

s*L

aby

rin

thul

om

yce

tes_

PW

19

*5

50

0.4

90

00

00

00

0.1

0

Lab

yri

nth

ulo

my

cete

s_T

AG

IRI-

15

*5

51

00

00

00

00

00

Lab

yri

nth

ulo

my

cete

s_T

AG

IRI-

17

*5

52

0.8

11

.74

1.0

11

.22

.31

1.4

71

.44

1.6

61

.08

1.2

9

Th

raus

toch

ytr

iace

ae*

55

30

00

00

00

00

0

Ap

lan

och

ytr

ium

55

40

.81

1.7

41

.01

1.2

2.3

11

.47

1.4

41

.66

0.6

31

.29

Th

raus

toch

ytr

iace

ae_

BS1

*5

55

0.2

40

00

.12

00

00

00

Th

raus

toch

ytr

iace

ae_

E1

70

*5

56

0.0

80

00

00

00

00

Lab

yri

nth

ulo

ides

55

70

00

00

00

.14

00

0

Sch

izo

chy

triu

m5

58

00

00

00

00

00

Th

raus

toch

ytr

ium

55

90

.24

00

00

0.1

40

.14

00

.31

0

Stra

men

op

iles

*St

ram

eno

pil

es*

Stra

men

op

iles

_M

AST

-12

*5

60

0.0

80

.54

0.4

90

.35

00

.14

0.1

40

.20

.10

.1

Stra

men

op

iles

_M

AST

-12

A*

56

10

.08

0.5

00

.70

0.3

90

00

0.1

1

Stra

men

op

iles

_M

AST

-12

E*

56

30

00

00

00

00

0

Stra

men

op

iles

_M

AST

-1C

*5

64

0.4

90

.47

0.4

90

.70

.70

.91

1.4

40

.94

1.0

80

Stra

men

op

iles

_M

AST

-3E

*5

65

0.0

80

00

00

00

00

Stra

men

op

iles

_M

AST

-3F

*5

66

00

00

00

00

00

Stra

men

op

iles

_M

AST

-3J*

56

70

.81

0.1

61

.21

.20

.59

0.9

10

.85

0.4

31

.08

0.3

3

SUPPLEMENT CHAPTER I

Table S8: continued

137

Stra

men

op

iles

_M

AST

-4D

*5

69

00

00

00

00

00

Stra

men

op

iles

*5

70

0.5

0.4

60

.15

1.2

0.2

0.1

30

.38

0.1

30

.30

.42

Stra

men

op

iles

_M

AST

-7B

*5

71

0.0

80

00

00

00

00

Stra

men

op

iles

*5

72

0.2

50

.21

0.2

00

0.1

30

.13

0.9

40

0.4

2

Stra

men

op

iles

_M

AST

-9D

*5

73

00

00

00

00

.14

00

Och

rop

hy

taO

chro

ph

yta

*O

chro

ph

yta

_A

NT

37

-16

*5

74

0.8

11

.01

0.5

90

.72

0.2

10

.38

1.4

40

.94

0.6

80

.82

Stra

men

op

iles

Bo

lido

mo

nas

57

50

00

.17

0.1

20

0.1

40

0.1

30

.11

0.3

8

Ch

ryso

ph

yce

aeC

hry

sop

hy

ceae

*5

76

00

00

00

00

00

Ch

ryso

ph

yce

ae_

Am

b-1

8S-

46

2*

57

70

.81

00

.15

00

0.1

30

00

0.1

Ch

ryso

ph

yce

ae_

CC

I40

*5

78

0.2

50

00

.12

0.1

80

00

0.1

10

Ch

rom

ulin

ales

*5

79

00

00

00

00

00

Ch

rom

ulin

ales

_A

MT

15

-27

-30

*5

80

00

00

00

00

00

Uro

glen

a5

83

00

00

00

00

00

Ch

ryso

ph

yce

ae_

E2

22

*5

85

0.8

11

.74

1.8

81

.21

.32

1.4

71

.44

1.6

61

.08

1.2

9

Ch

ryso

ph

yce

ae_

LG

01

-04

*5

86

00

00

00

00

00

Och

rop

hy

taC

hry

sop

hy

ceae

_L

G0

1-0

9*

58

70

.08

00

00

00

00

0

Ch

ryso

ph

yce

ae_

LG

07

-07

*5

88

0.0

80

00

00

00

00

Och

rom

on

adal

es*

59

00

00

00

00

00

0

Och

rom

on

as5

93

00

00

00

00

00

Par

aph

yso

mo

nas

59

40

.81

1.0

41

.12

1.2

2.3

11

.47

0.8

0.9

61

.08

1.2

9

Ch

ryso

ph

yce

ae_

P3

4.4

5*

59

50

00

0.1

20

00

00

0

Ch

ryso

ph

yce

ae_

P3

4.4

8*

59

60

00

00

00

00

0

Mal

lom

on

as5

97

00

00

00

00

00

.12

Tes

sell

aria

59

80

00

00

00

00

0

Ch

ryso

ph

yce

ae*

59

90

.08

00

00

00

00

0

Dia

tom

ea_

3b-

B4

*6

00

00

00

00

00

00

Bac

illa

rio

ph

yce

ae*

60

10

.08

00

00

00

00

0

Ach

nan

thes

60

20

00

00

00

00

0

Am

ph

ora

60

30

.24

00

0.1

20

00

00

0

Ast

erio

nel

lop

sis

60

40

00

00

00

00

0

Bac

illa

ria

60

50

00

00

00

00

0

SUPPLEMENT CHAPTER I

Table S8: continued

138

C

occ

on

eis

60

60

00

00

00

00

0

Cra

ticu

la6

07

00

00

00

00

00

Cy

lin

dro

thec

a6

08

00

00

00

00

00

Dia

tom

a6

09

00

00

00

00

00

Eun

oti

a6

10

00

00

00

00

00

Fis

tuli

fera

61

10

00

00

00

00

0

Nav

icul

a6

13

0.8

10

.16

00

.34

00

00

0.1

10

.11

Nit

zsch

ia6

14

0.0

80

0.4

90

00

00

00

Ph

aeo

dact

ylu

m6

15

00

00

00

00

00

Stra

men

op

iles

Och

rop

hy

taD

iato

mea

Ple

uro

sigm

a6

16

00

00

00

00

00

Pse

udo

-nit

zsch

ia6

17

0.2

50

00

.35

00

.41

00

0.3

50

.11

Rh

aph

on

eis

61

80

.50

.49

1.1

0.3

60

0.9

00

.14

0.1

10

.1

Sell

aph

ora

61

90

00

00

00

00

0

Stau

ron

eis

62

00

00

00

00

00

0

Bac

illa

rio

ph

yce

ae_

Zeu

k1

0*

62

10

00

00

00

00

0

Med

iop

hy

ceae

*6

22

0.8

10

.16

0.5

30

.12

0.2

11

.47

0.4

30

.49

1.0

80

.34

Bid

dulp

hia

62

30

.09

00

00

00

00

0

Bid

dulp

hio

psi

s6

24

00

00

00

0.1

30

00

Cam

py

losi

ra6

25

00

00

00

00

00

Ch

aeto

cero

s6

26

0.8

11

.01

00

.12

0.6

90

.41

1.4

40

.45

0.6

60

.34

Cy

clo

tell

a6

27

00

.15

00

00

00

00

Cy

mat

osi

ra6

28

00

00

00

00

00

Dis

cost

ella

62

90

.23

00

00

00

.13

00

0

Eun

oto

gram

ma

63

00

.25

00

00

00

00

0

Ex

tubo

cell

ulus

63

10

00

00

00

00

0

Ley

anel

la6

32

00

00

00

00

.14

0.1

0.1

Med

iop

hy

ceae

_M

E-E

uk-D

BT

11

6*

63

30

.49

0.1

60

.53

00

0.1

30

00

0

Min

idis

cus

63

40

.81

0.1

51

.88

1.2

0.5

91

.47

0.1

31

.66

0.6

30

.74

Pap

ilio

cell

ulus

63

50

.08

00

00

.21

00

00

0

Pie

rrec

om

per

ia6

36

0.0

80

00

0.2

00

.14

00

0.1

1

Pla

gio

gram

ma

63

70

00

00

00

00

0

SUPPLEMENT CHAPTER I

Table S8: continued

139

Pla

gio

gram

mo

psi

s6

38

0.8

10

.97

0.5

90

.70

.26

1.4

70

.46

1.6

61

.08

0.7

4

Pla

nk

ton

iell

a6

39

0.2

50

00

00

.14

00

00

Ro

undi

a6

40

0.2

60

00

0.2

10

00

00

.1

Dia

tom

eaSk

elet

on

ema

64

10

.08

00

.16

00

00

00

0

Step

han

odi

scus

64

20

.08

00

00

00

00

0

Th

alas

sio

sira

64

40

.81

1.7

41

.88

1.2

1.3

1.4

71

.44

1.6

60

.63

1.2

9

Fra

gila

rial

es*

64

60

00

00

00

00

0

Stra

men

op

iles

Och

rop

hy

taL

icm

op

ho

ra6

50

0.0

80

00

00

00

00

Nan

ofr

ustu

lum

65

10

.48

0.1

60

00

00

00

0.1

Op

eph

ora

65

20

.08

00

00

00

00

0

Stau

rosi

ra6

53

00

00

00

0.1

40

.14

00

En

dict

ya

65

40

00

00

00

00

0

Hy

alo

disc

us6

55

00

00

00

00

00

Step

han

op

yx

is6

56

0.0

80

.17

00

00

00

00

Dia

tom

ea_

ME

-Euk

-FW

10

*6

57

0.8

10

0.5

30

.12

0.3

0.4

00

0.1

10

.11

Dic

tyo

cho

ph

yce

aeP

seud

op

edin

ella

65

80

00

00

0.1

40

00

0

Rh

izo

chro

mul

ina

65

90

00

00

00

00

0

Och

rop

hy

ta`*

Pic

op

hag

us6

61

0.8

11

.01

0.2

0.1

20

.59

0.4

80

.13

00

.68

0.8

2

Des

mar

esti

a6

62

0.4

91

.74

0.4

80

.35

2.3

11

.47

0.8

0.9

50

.63

0.7

4

Ect

oca

rpal

es*

66

40

0.2

0.1

60

.11

1.3

20

.39

0.1

30

0.1

0.4

Ast

ero

clad

on

66

50

00

00

0.1

40

00

0

Hal

oth

rix

66

60

.08

0.5

00

0.1

80

0.3

70

.54

00

.12

Ph

aeo

ph

yce

aeIs

hig

e6

67

00

00

00

00

00

Py

laie

lla

66

80

00

00

00

00

0

Scy

tosi

ph

on

66

90

00

00

00

.13

00

0

Ak

kes

iph

ycu

s6

70

00

00

00

00

00

Po

stel

sia

67

10

00

00

00

00

0

Och

rop

hy

taP

hae

op

hy

ceae

Sacc

har

ina

67

20

.81

1.7

41

.01

1.2

2.3

11

.47

1.4

41

.66

1.0

81

.29

Spo

roch

on

us6

73

00

00

00

00

00

Rap

hid

op

hy

ceae

Go

ny

ost

om

um6

74

00

00

00

00

00

Har

amo

nas

67

50

.08

00

00

00

00

0

SUPPLEMENT CHAPTER I

Table S8: continued

140

Ch

atto

nel

lale

s_M

OC

H-3

*6

76

00

00

00

00

00

Per

on

osp

oro

my

cete

sP

ero

no

spo

rom

yce

tes*

Per

on

osp

oro

my

cete

s*6

78

0.2

40

.19

0.1

71

.20

0.7

90

.11

0.1

70

.68

0.4

2

Stra

men

op

iles

Ap

han

om

yce

s6

79

00

00

00

00

0.1

10

Hal

iph

tho

ros

68

10

.49

00

0.1

10

.25

0.1

60

.42

0.1

70

.31

0.3

9

Hal

ocr

usti

cida

68

20

00

.48

0.1

10

.25

0.1

30

0.4

40

.31

0

Hal

op

hy

top

hth

ora

68

30

.08

00

00

0.1

30

00

.10

.15

Hap

togl

oss

a6

84

00

00

00

00

00

Lag

enid

ium

68

50

.08

01

.21

.20

0.3

90

0.5

31

.08

0.3

9

Lep

tole

gnia

68

60

00

00

00

00

0

Olp

idio

psi

s6

87

0.4

80

.54

0.4

80

.34

1.4

60

.90

.81

.66

0.6

61

.29

Ph

yti

um6

88

00

00

00

00

0.1

0

Ph

yto

ph

tho

ra6

89

0.2

30

.54

0.1

60

.71

.46

0.9

0.8

1.6

61

.08

1.2

9

Py

thiu

m6

90

0.4

80

1.1

20

.11

0.2

0.9

0.1

40

1.0

81

.29

Per

on

osp

oro

my

cete

s*6

92

0.8

10

.16

1.8

80

.70

.20

.79

0.3

71

.66

1.0

81

.29

SUPPLEMENT CHAPTER I

141

Table S9 Prokaryotic taxonomic classes detected in pooled biofilm samples (n=50, incubated from August

2013 – November 2014) associated to nine different synthetic polymers and glass, and seawater samples (n=42,

collected weekly from March 2012 – February 2013) of Helgoland Roads. Classes are represented by the number

of OTUs present in the given environment; in bold: classes exclusive present in one habitat. A * indicates the

term `unclassified class`.

Class Biofilm Seawater

Acidimicrobiia 4 3

Acidobacteria 5

Actinobacteria 1

Alphaproteobacteria 16 28

Ardenticatenia 2

Bacteria_BD1-5* 1

Betaproteobacteria 3 3

Caldilineae 1

Caldilineae 2

Chloroflexi* 1

Cyanobacteria 1

Cyanobacteria_Chloroplast* 1 1

Cytophagia 3 2

Deferribacteres 1

Deferribacteres Incertae Sedis* 1

Deinococci 1

Deltaproteobacteria 18 4

Epsilonproteobacteria 1 1

Flavobacteria 6 19

Gammaproteobacteria 18 37

Gemmatimonadetes 2 1

Holophagae 3

Latescibacteria* 1

Lentisphaerae_LD1-PB3* 1

Melainabacteria 2

Nitrospira 1

Oligosphaeria 1

Omnitrophica_NPL-UPA2* 1

Opitutae 1 2

Parcubacteria* 1

Phycisphaerae 4

Planctomycetacia 8 2

Planctomycetes_028H05-P-BN-P5* 1

Planctomycetes_BD7-11* 1

Planctomycetes_OM190* 1

Planctomycetes_Pla3 lineage* 1

Planctomycetes_Pla4 lineage* 1

Planctomycetes_vadinHA49* 1

Proteobacteria_AEGEAN-245* 1 1

Proteobacteria_ARKICE-90* 1

Proteobacteria_JTB23* 1

Proteobacteria_SC3-20* 1

Proteobacteria_SPOTSOCT00m83* 1 1

Proteobacteria_TA18* 1

Saccharibacteria* 1

Sphingobacteriia 5 2

Thaumarchaeota_Marine Group I* 2 1

Thermoplasmata 1

Verrucomicrobia_OPB35 soil group 1

Verrucomicrobiae 1 2

Supplement material for Chapter II

The Plastisphere –

Uncovering tightly attached plastic “specific” microorganisms

Detailed information on the development of the high-pressure treatment technique and the

staining procedure in order to visualize high-pressure treated biofilms is given in the supporting

file. Furthermore, three figures illustrating the cell numbers counted per mm2 evaluated at

different time and pressures, the abundance profiles of short- and long-term incubated

communities on the family level, and most abundant and discriminative OTUs. Further ten

tables giving detailed information about plastic types, GLM model results, PERMANOVA and

PERMDISP tests and Univariate Diversity indices.

SUPPLEMENT CHAPTER II

144

Development of the new high-pressure treatment technique

To develop the method and to evaluate whether there is a significance of time or pressure a pre-

test with Polypropylene (PP) in triplicate has been performed (Fig S1). The high-pressure

device (LicoJet) needs to be affiliated to a compressed air supply to create an air flow with high

velocity through the device towards the restricted opening of the nozzle. The liquid inside the

device nozzle gets pressed out by the pressurized air with the adjusted pressure. The LicoJet

was held in a mounting structure to ensure time of spraying and nozzle distance to be controlled.

Sterile seawater (0.2 µm filtered and autoclaved) was shot vertical, with a working distance of

1 cm on the biofilm associated to the different substrates in a time series of 2, 3 and 4 minutes,

and a change in pressure at 2, 3 and 4 bar. The exposed spots were stained with SYBR Gold to

determine the total cell count. Evaluation of the cell counts of remaining strongly attached cells

on the substrate showed that neither the impacted pressure of the water current nor the duration

of the pressure had any significant influence on the amount of cells (Fig S1, Table S2).

Visualization of high-pressure treated biofilms

To distinguish cells with membrane integrity from the ones with a damaged cell membrane

after high-pressure treatment double staining with propidium iodide (PI) and SybrGreen was

performed. In this study a mix of both stains was prepared according to the concentrations

investigated by Falcioni et al (2008). In total, 20 µl of the double stain were added on each

high-pressure treated spot and stained for 30 minutes at room temperature in the dark. After the

staining process the polymeric foils were washed in deionized water to remove the unbound

staining solution and dried with Whatman paper. To prevent the fluorescent from rapid

photobleaching, the sample got fixed with a 0.1% (v/v) p-phenylenediamine anti-fade mounting

medium. SybrGreen stained cells were detected with the optical microscope Axioplan2,

imaging (Zeiss; Oberkochen, Germany) using a bandpass excitation filter with the wavelengths

between 450 to 490 nm and a longpass emission filter of 515 nm into the IR spectra (filter set

09; Zeiss; Oberkochen, Germany). To evaluate how many cells of the total amount have a

damaged cell membrane a bandpass excitation filter that passes light at a wavelength of 534 to

558 nm and is therefore ideal to excite PI has been used (filter set 20; Zeiss; Oberkochen,

Germany). The emission filter passes the fluorescence from 575 to 640 nm and therefore

transmits the emission of PI (617 nm) but excludes emission of SYBR Green (512 nm).

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Table S1 Sample information about synthetic polymers used within this study.

Polymer Abbreviation Monomer Manufacturer

Low density polyethylene LDPE (C2H4)n ORBITA-FILM GmbH

High density polyethylene HDPE (C2H4)n ORBITA-FILM GmbH

Polypropylene PP (C3H6)n ORBITA-FILM GmbH

Polystyrene PS (C8H8)n Ergo.fol norflex GmbH

Styrene acrylonitrile SAN (C8H8)n-(C3H3N)m Ergo.fol norflex GmbH

Polyurethane prepolymer PESTUR (C4H4O5)n Bayer

Polylactic acid PLA (C3H4O2)n Folienwerk Wolfen GmbH

Polyethylene terephthalate PET (C10H8O4)n Mitsubishi Polyester Film

Polyvynil chloride PVC (C2H2Cl)n Leitz

Table S2 GLM model results of cell counts against exposure time and pressure. Both variables and their

interaction resulted not significant (p-value > 0.05). Est. Average represents the estimated average, Std. Error

represents the standard error.

Est. Average Std. Error p-value

Pressure 0.0007133 0.0006328 0.271

Time 0.0008184 0.0006655 0.231

Time * Pressure -0.0002071 0.0002241 0.365

Table S3 GLM model results of cell count distinguished in membrane damaged and intact cells after a high

pressure water treatment at 4 bars for 2 minutes and staining with PI and SYBR Green. Both variables and their

interaction resulted significant (p-value < 0.05). Est. Average represents the estimated average of the mean cell

counts, Std. Error represents the standard error of the mean cell counts.

Membrane damaged Membrane intact

Samples Est. Average Std. Error p-value Est. Average Std. Error p-value

HDPE 693.2000 5.0794 < 0.05 626.1000 4.8221 < 0.05

LDPE 1307.2000 6.9672 < 0.05 805.1000 5.4664 < 0.05

PESTUR 491.2000 4.2801 < 0.05 116.1000 2.0889 < 0.05

PET 18.2000 0.8948 < 0.05 36.1000 1.1834 < 0.05

PLA 109.2000 2.0432 < 0.05 141.1000 2.2998 < 0.05

PP 2127.2000 8.8832 < 0.05 2755.1000 10.1047 < 0.05

PS 1006.2000 6.1150 < 0.05 707.1000 5.1237 < 0.05

PVC 439.2000 4.0489 < 0.05 149.1000 2.3634 < 0.05

SAN 288.2000 3.2864 < 0.05 197.1000 2,7135 < 0.05

Glass 1.8000 0.2449 < 0.05 0.9000 0.1732 < 0.05

Table S4 PERMANOVA main tests of biofilm community on different re-colonized synthetic polymers and

glass based on Hellinger distance of operational taxonomic units (OTUs). P-values were obtained using type III

sums and 9999 permutations under the full model. d.f.: degrees of freedom, SS: sums of squares; MS: mean

squares, perms: number of unique permutations per comparison. Significant results (p (perm) < 0.05) are

highlighted in bold.

Source of variation d.f. SS MS Pseudo-F p (perm)1 perms

Substrate 9 16.945 1.8827 56.281 0.0001 9847

Res 40 1.3381 0.0335

Total 49 18.283

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Table S5 PERMANOVA and PERMDISP pair-wise tests biofilm communities on different re-colonized

synthetic polymers and glass based on Hellinger distance of operational taxonomic units (OTUs). Significant

results (p (perm) < 0.05) are highlighted in bold.

PERMANOVA PERMDISP

Comparison t (perm) p (perm)1 t (perm) p (perm)1

Glass vs.

HDPE 7.798 0.007 1.987 0.138

LDPE 8.351 0.007 1.969 0.140

PESTUR 4.917 0.007 1.512 0.270

PET 6.402 0.007 1.709 0.204

PLA 4.889 0.009 0.654 0.608

PP 7.703 0.008 0.892 0.577

PS 7.234 0.007 1.409 0.336

PVC 6.527 0.007 1.539 0.265

SAN 5.774 0.009 0.978 0.528

HDPE vs.

LDPE 6.739 0.009 0.131 0.912

PESTUR 8.299 0.008 0.339 0.812

PET 7.550 0.007 0.015 0.969

PLA 8.311 0.008 1.352 0.345

PP 7.710 0.008 1.986 0.161

PS 7.400 0.008 0.628 0.615

PVC 8.205 0.007 0.950 0.484

SAN 7.739 0.006 1.375 0.306

LDPE vs.

PESTUR 8.816 0.007 0.250 0.827

PET 7.664 0.009 0.085 0.930

PLA 9.296 0.007 1.312 0.395

PP 6.750 0.007 2.054 0.142

PS 8.277 0.009 0.547 0.695

PVC 9.321 0.007 0.893 0.463

SAN 8.167 0.007 1.345 0.310

PESTUR vs.

PET 6.510 0.008 0.271 0.825

PLA 5.616 0.010 0.902 0.505

PP 7.637 0.007 1.138 0.360

PS 8.081 0.009 0.216 0.856

PVC 6.785 0.009 0.356 0.818

SAN 6.187 0.010 0.800 0.516

PET vs.

PLA 7.220 0.008 1.132 0.406

PP 8.099 0.009 1.408 0.293

PS 8.459 0.008 0.495 0.696

PVC 8.080 0.008 0.673 0.602

SAN 6.914 0.008 1.065 0.418

PLA vs.

PP 8.754 0.006 0.073 0.983

PS 7.128 0.007 0.756 0.562

PVC 6.590 0.008 0.803 0.658

SAN 5.080 0.008 0.253 0.832

PP vs.

PS 9.128 0.010 0.997 0.440

PVC 8.756 0.010 1.380 0.199

SAN 8.362 0.009 0.290 0.849

PS vs. PVC 7.067 0.008 0.109 0.928

SAN 7.783 0.008 0.628 0.597

PVC vs. SAN 7.828 0.007 0.701 0.571

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Table S6 Univariate Diversity indices of biofilm communities on different re-colonized synthetic polymers and

glass based on read counts of operational taxonomic units (OTUs). S: Total species, N: Total individuals, d:

Species richness (Margalef), J': Pielou`s evenness, H'(log2): Shannon.

Sample S N d J' H'(log2)

Glass_source 73 20955 7.236 0.3211 1.988

Glass_1 86 17459 8.702 0.4509 2.897

Glass_2 70 12466 7.316 0.4806 2.946

Glass_3 47 7621 5.146 0.4414 2.452

Glass_4 91 25996 8.853 0.4651 3.027

Glass_5 90 20852 8.949 0.4435 2.879

HDPE_source 250 22046 24.9 0.5646 4.497

HDPE_1 96 11970 10.12 0.6059 3.99

HDPE_2 67 14732 6.877 0.5891 3.573

HDPE_3 88 8869 9.571 0.5348 3.454

HDPE_4 103 20693 10.26 0.5247 3.508

HDPE_5 128 21242 12.75 0.5463 3.824

LDPE_source 157 17389 15.98 0.5413 3.948

LDPE_1 96 24979 9.382 0.4477 2.948

LDPE_2 98 27832 9.478 0.46 3.043

LDPE_3 98 20035 9.793 0.4376 2.895

LDPE_4 92 21794 9.11 0.4337 2.83

LDPE_5 104 22508 10.28 0.4833 3.238

PESTUR_source 163 34157 15.52 0.379 2.785

PESTUR_1 84 20485 8.361 0.5603 3.581

PESTUR_2 79 12688 8.255 0.5542 3.493

PESTUR_3 74 14687 7.608 0.5536 3.437

PESTUR_4 83 17803 8.378 0.5871 3.743

PESTUR_5 77 25107 7.502 0.5535 3.469

PET_source 173 20954 17.29 0.5822 4.328

PET_1 113 24762 11.07 0.4958 3.382

PET_2 83 14111 8.582 0.5451 3.475

PET_3 104 23971 10.21 0.5579 3.738

PET_4 93 25589 9.064 0.5439 3.556

PET_5 94 23672 9.233 0.5301 3.475

PLA_source 187 28842 18.11 0.4914 3.709

PLA_1 103 17109 10.46 0.4159 2.781

PLA_2 94 29710 9.03 0.3386 2.22

PLA_3 72 22882 7.073 0.3602 2.223

PLA_4 78 31240 7.44 0.4121 2.59

PLA_5 89 28330 8.584 0.3245 2.101

PP_source 84 12495 8.799 0.5073 3.243

PP_1 61 16602 6.175 0.4798 2.846

PP_2 60 15213 6.127 0.5376 3.175

PP_3 53 15154 5.402 0.5017 2.874

PP_4 74 16802 7.503 0.4911 3.05

PP_5 48 24150 4.657 0.5134 2.867

PS_source 190 24781 18.68 0.6684 5.06

PS_1 122 22754 12.06 0.5377 3.726

PS_2 118 15715 12.11 0.5568 3.832

PS_3 103 23498 10.13 0.5518 3.69

PS_4 95 27453 9.197 0.5682 3.733

PS_5 126 40903 11.77 0.5437 3.793

PVC_source 168 38893 15.8 0.5776 4.27

PVC_1 65 15088 6.652 0.4716 2.84

PVC_2 86 17938 8.678 0.5083 3.266

PVC_3 104 31841 9.934 0.5593 3.748

PVC_4 62 18382 6.212 0.6244 3.718

PVC_5 79 16165 8.049 0.5112 3.222

SAN_source 186 34111 17.72 0.5278 3.979

SAN_1 91 22864 8.967 0.3977 2.588

SAN_2 75 8973 8.13 0.4687 2.919

SAN_3 92 26426 8.937 0.4497 2.934

SAN_4 93 38640 8.71 0.3896 2.547

SAN_5 83 30449 7.943 0.3482 2.22

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Table S7 SIMPER analysis of re-colonized communities jointly contributing to the total similarity within and

dissimilarity between different groups of synthetic polymers and glass. Av.Si%: average percentage similarity

within the different groups, Av.δi%: average dissimilarity between the different groups.

Av.Si% Av.δi%

LDPE 88.16 LDPE 40.29

PP 84.55 PP 56.24 44.87

PS 87.55 PS 42.75 50.35 66.01

PET 89.16 PET 42.99 44.16 54.96 51.01

PLA 85.41 PLA 53.42 64.40 67.33 47.61 50.50

SAN 86.55 SAN 50.90 54.18 61.42 50.61 45.56 38.35

PESTUR 88.08 PESTUR 52.25 58.44 55.24 52.74 40.94 38.93 42.71

PVC 86.51 PVC 49.60 60.90 60.01 43.54 51.09 42.02 53.97 42.25

Glass 84.35 Glass 54.77 60.40 60.10 49.79 45.01 38.97 45.56 35.11 43.89

HDPE 88.63 HDPE LDPE PP PS PET PLA SAN PESTUR PVC

Fig S1 Barplot of the cell numbers per mm2 evaluated at different time and pressures. The bars represent the

different pressures with 2, 3, 4 bar at 2, 3, 4 minutes respectively. The vertical bars denote the Standard Error of

the data.

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Fig S2 Abundance profiles of the source (short-term) and re-colonized (long-term) communities on the family

level on different synthetic polymers and glass. OTUs with a mean relative abundance of at least 0.1% in one

substrate type (nsource=1; nre-col=5) were analysed. Displayed are taxonomic families with abundances of > 1% in at

least one substrate type. The group `others` was made up of families with abundances < 1%. A * indicates the term

“unclassified”.

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Fig S3 Discriminative OTUs of the nine different plastics (n=5). OTUs with a mean relative abundance of at

least 0.1% (n=5) in at least one substrate type were analysed. Displayed are OTUs jointly contributing to the total

dissimilarity of at least 3% between plastic or with relative abundance of at least 1% on one substrate type. OTUs

with a mean relative abundance of at least 0.1% present on both, plastics and glass, were rejected. The amount of

contribution is indicated by the colour of cells, darker colours represent higher contributions. A * indicates the

term “unclassified”, # indicates the term “uncultured”.

Supplement material for Chapter III

Dangerous Hitchhikers?

Evidence for potentially pathogenic Vibrio spp. on microplastic particles

Six tables giving detailed information about sampling stations sampling dates and

corresponding geographic coordinates of sampling sites, water volume which passed through

the Neuston net, environmental parameters, collected particle identity at corresponding stations,

MALDI-TOF Vibrio identification results & species-specific and virulence-associated-gene

PCR results.

SUPPLEMENT CHAPTER III

152

Table S1: Sampling stations with sampling dates and corresponding geographic coordinates of sampling sites.

HE409 2013 HE430 2014

Station

No. Sampling Date Latitude N Longitude E

Station

No. Sampling Date Latitude N Longitude E

1 19.09.2013 54,0822 7,4608 39 31.07.2014 53,8252 7,7673

2 19.09.2013 53,9931 6,9928 40 31.07.2014 53,7947 7,3492

3 19.09.2013 53,8681 6,4367 41 01.08.2014 53,7475 6,9987

4 20.09.2013 53,7061 6,6381 42 01.08.2014 53,7177 6,6760

5 20.09.2013 53,4842 6,8097 43 01.08.2014 53,6513 6,3315

6 20.09.2013 53,3183 7,0392 44 02.08.2014 53,6130 6,1380

7 21.09.2013 53,8256 7,1300 45 02.08.2014 53,5530 5,5923

8 21.09.2013 53,8897 7,6250 46 02.08.2014 53,4747 5,1825

9 21.09.2013 53,6847 8,0892 47 03.08.2014 53,3033 4,8048

10 21.09.2013 53,5269 8,1800 48 03.08.2014 53,1422 4,6017

11 22.09.2013 53,5539 8,5547 49 03.08.2014 52,9177 4,4325

12 22.09.2013 53,7222 8,2764 50 04.08.2014 52,4260 4,3475

13 22.09.2013 53,8344 8,1394 51 04.08.2014 52,1702 4,0132

14 22.09.2013 54,0000 8,0264 52 04.08.2014 51,8667 3,6258

15 22.09.2013 54,1489 7,8858 53 05.08.2014 51,5395 3,1822

16 23.09.2013 54,3328 7,7178 54 05.08.2014 51,2847 2,5150

17 23.09.2013 54,6958 7,9758 55 05.08.2014 51,0777 1,9037

18 23.09.2013 54,4928 8,0947 56 06.08.2014 50,4965 1,1654

19 23.09.2013 54,2667 8,2956 57 06.08.2014 51,5836 2,4426

20 24.09.2013 54,1056 8,3936 58 07.08.2014 52,1503 2,8428

21 24.09.2013 53,9439 8,6719 59 07.08.2014 52,9783 3,2288

22 24.09.2013 53,8819 9,0658 60 08.08.2014 53,9062 3,1847

23 25.09.2013 54,3433 10,1742 61 08.08.2014 54,8117 3,3883

24 25.09.2013 54,6528 10,1697 62 09.08.2014 55,8355 3,5624

25 25.09.2013 54,7356 10,1739 Helgoland drift line 2013

26 25.09.2013 54,8333 9,8628 63 01.08.2013 54,2875 7,9000

27 26.09.2013 54,5550 10,8672

28 26.09.2013 54,5822 11,0358

29 26.09.2013 54,3889 11,5358

30 26.09.2013 54,0842 11,1842

31 27.09.2013 54,2861 12,0853

32 27.09.2013 54,6108 12,3831

33 27.09.2013 54,8261 13,0408

34 27.09.2013 54,8333 13,7525

35 28.09.2013 54,7058 14,3600

36 28.09.2013 54,5117 14,2575

37 28.09.2013 54,2375 14,2839

38 28.09.2013 53,9975 14,2272

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Table S2: Water volume which passed through the Neuston net (300 µm), determined by the use of a mechanical

flowmeter.

HE409 2013 HE430 2014

Station No. Start Flow End Flow Liter m³ Station No. Start Flow End Flow Liter m³

1 49649 57284 68715 68,72 39 92509 100113 68436 68,44

2 62367 67824 49113 49,11 40 4196 14306 90990 90,99

3 72922 77213 38619 38,62 41 18813 27785 80748 80,75

4 83701 91832 73179 73,18 42 32919 44963 108396 108,40

5 96327 101807 49320 49,32 43 49906 57772 70794 70,79

6 3433 7619 37674 37,67 44 60029 69737 87372 87,37

7 9638 16657 63171 63,17 45 73637 79552 53235 53,24

8 19047 27835 79092 79,09 46 82292 90188 71064 71,06

9 31024 43069 108405 108,41 47 94068 106096 108252 108,25

10 46675 54364 69201 69,20 48 10651 21495 97596 97,60

11 57752 67578 88434 88,43 49 25453 36755 101718 101,72

12 68990 77696 78354 78,35 50 39892 48927 81315 81,32

13 80181 86948 60903 60,90 51 50817 60723 89154 89,15

14 89189 97154 71685 71,69 52 63436 75348 107208 107,21

15 1698 7924 56034 56,03 53 77074 88854 106020 106,02

16 14722 21229 58563 58,56 54 93352 105088 105624 105,62

17 28134 36277 73287 73,29 55 7596 18520 98316 98,32

18 39255 47167 71208 71,21 56 22866 34363 103473 103,47

19 50437 57539 63918 63,92 57 38202 50200 107982 107,98

20 61413 71174 87849 87,85 58 53857 68068 127899 127,90

21 73193 84028 97515 97,52 59 72092 86125 126297 126,30

22 85496 95390 89046 89,05 60 91829 106432 131427 131,43

23 119 5203 45756 45,76 61 11632 23306 105066 105,07

24 6415 15374 80631 80,63 62 28821 40817 107964 107,96

25 19624 27148 67716 67,72

26 31600 38955 66195 66,20

27 42023 51727 87336 87,34

28 56054 60557 40527 40,53

29 66420 74847 75843 75,84

30 79712 82547 25515 25,52

31 84794 95416 95598 95,60

32 928 10663 87615 87,62

33 16057 24210 73377 73,38

34 27942 35303 66249 66,25

35 40476 47486 63090 63,09

36 53208 60592 66456 66,46

37 66177 75699 85698 85,70

38 82645 89783 64242 64,24

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Table S3: Environmental parameters. Temperatures and salinities recorded at each station.

HE 409 2013 HE 430 2014

Station No. °C PSU Station No. °C PSU

1 16,9 32,31 39 19,70 32,6

2 17,25 32,57 40 20,43 32,32

3 17,14 32,85 41 21,13 32,18

4 15,77 31,86 42 21,65 31,66

5 14,90 30,36 43 20,73 32,29

6 15,07 25,14 44 20,70 32,56

7 16,72 31,48 45 20,95 33,08

8 16,83 31,50 46 20,50 33,18

9 15,15 30,68 47 20,08 32,77

10 15,17 31,19 48 19,85 33,00

11 15,70 14,23 49 19,96 34,00

12 15,10 25,30 50 20,54 30,96

13 15,74 30,82 51 20,04 31,62

14 16,88 32,16 52 20,45 32,48

15 16,64 32,21 53 20,71 32,40

16 16,19 31,74 54 19,56 34,16

17 16,23 29,45 55 18,83 34,33

18 16,08 29,64 56 18,34 33,77

19 15,82 28,47 57 18,79 34,46

20 15,38 27,64 58 18,38 33,53

21 15,66 16,89 59 18,89 33,51

22 16,85 3,03 60 17,73 34,00

23 15,24 15,93 61 19,07 34,00

24 15,27 15,73 62 18,70 34,20

25 15,41 15,5 Helgoland drift line 2013

26 15,35 16,61 63 16,60 30,23

27 14,93 16,8

28 14,79 15,27

29 14,95 12,99

30 14,84 12,64

31 14,07 11,65

32 14,56 8,75

33 15,11 7,59

34 14,52 7,43

35 15,25 7,26

36 14,67 7,27

37 14,85 7,06

38 14,46 5,67

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Table S4: Occurrence of visible particles collected during the North and Baltic Sea cruises and on Helgoland

beach. Stations and corresponding collected particle samples, identity of the material and the corresponding HIT-

Score of ATR-FT IR analysis are display here (Hit-Scores of ≥700 were accepted. Any matches with quality index

<700 were individually inspected and interpreted based on the closeness of their absorption frequencies to those

of chemical bonds in the known polymers, N.i. = Not identified).

HE 409 2013 HE 430 2014

Station

No.

Sample

No. Material

ATR-FT IR

HIT-Score

Station

No.

Sample

No. Material

ATR-FT IR

HIT-Score

1 1P1 Varnish 646 39 1P1 Acrylnitril-Butadien-

Styrol 917

1 1P2 Keratin 185 39 1P2 Polystyrene 994

1 1P3 Polyethylene 794 39 1P3 Polystyrene 915

2 2P1 Polystyrene 996 39 1P4 Varnish 653

2 2P2 Polyethylene 998 40 2P1 Polyethylene 795

2 2P3 Polypropylene 903 40 2P2 Polyethylene 995

2 2P4 Polystyrene 997 40 2P3 Polyethylene 993

2 2P5 Polystyrene 840 40 2P4 Polypropylene 920

2 2P6 N.i. 382 40 2P5 Polyethylene 784

3 3P1 Polyvinylalcohol 329 41 3P1 Polyethylene 817

3 3P2 Varnish 504 41 3P2 Polyethylene 997

3 3P3 Polyethylene 298 41 3P3 Polyethylene 998

3 3P4 Polypropylene 747 41 3P4 Polyethylene 993

3 3P5 Polypropylene 795 41 3P5 Polyethylene 990

3 3P6 Polystyrene 901 41 3P6 Polyethylene 993

4 4P1 Polyethylene 986 41 3P7 Polyethylene 846

4 4P2 Ethylen-vinylalcohol 968 41 3P8 Polypropylene 884

4 4P3 Polyethylene 997 42 4P1 Polypropylene 696

4 4P4 Polypropylene 933 42 4P2 Varnish 560

4 4P5 Ethylen-vinylalcohol 947 43 5P1 Polyethylene 989

4 4P6 Polyethylene 997 43 5P2 Polyethylene 797

4 4P7 Polyethylene 997 43 5P3 Polypropylene 812

5 5P1 Polypropylene 755 47 9P2 Polyethylene 398

5 5P2 Polyethylene 790 49 11P1 Polystyrene 986

5 5P3 Polyethylene 695 49 11P2 Polyethylene 796

6 6P1 Chitin 542 49 11P3 Polyethylene 944

6 6P2 Chitin 723 49 11P4 Chitin 608

6 6P3 Polyethylene 987 50 12P1 Polyethylene 823

6 6P4 Polypropylene 877 52 14P1 Polypropylene 854

6 6P5 Polypropylene 563 52 14P2 Polyethylene 819

6 6P6 Chitin 612 52 14P3 Polyethylene 707

7 7P1 Polypropylene 871 52 14P4 Polyethylene 796

7 7P2 Polyethylene 695 53 15P1 Polypropylene 852

7 7P3 Polyethylene 837 55 17P1 Polyethylene 644

7 7P4 Polyethylene 723 55 17P2 Polyethylene 609

8 8P1 Polyethylene 838 56 18P1 Polyethylene 786

SUPPLEMENT CHAPTER III

Table S4: continued

156

8 8P2 Polyethylene 838 56 18P2 Polypropylene 795

8 8P3 Polyethylene 996 56 18P3 Polyethylene 701

8 8P4 Polyethylene 998 56 18P4 Polyethylene 529

8 8P5 Polyamide 487 56 18P5 Stearic Acid 291

9 9P1 Ethylen-vinylalcohol 645 56 18P6 Polypropylene 450

9 9P2 Polypropylene 852 56 18P7 Polypropylene 688

9 9P3 Polyethylene 564 56 18P8 Polyethylene 466

9 9P4 Polypropylene 841 56 18P9 Polyethylene 891

9 9P5 Polyethylene 778 56 18P10 Polyethylene 820

10 10P1 Polyethylene 988 56 18P11 Chitin 447

11 11P1 Polyethylene 796 56 18P12 Polypropylene 848

13 13P1 Polyethylene 837 56 18P13 Polystyrene 984

14 14P1 Polyethylene 836 57 19P1 Polypropylene 562

15 15P1 Polyethylene 820 57 19P2 Polyethylene 703

15 15P2 Polyethylene 839 57 19P3 Polyethylene 663

15 15P3 Polyethylene 837 57 19P4 Polyethylene 406

15 15P4 Polyethylene 725 57 19P5 Polyethylene 387

15 15P5 Polyethylene 996 58 20P1 N.i.

15 15P6 Polyethylene 726 58 20P2 N.i.

15 15P7 Polyethylene 726 58 20P3 N.i.

16 16P1 Polyethylene 692 58 20P4 N.i.

16 16P2 Polyethylene 993 58 20P5 N.i.

16 16P3 Polyethylene 516 58 20P6 N.i.

16 16P4 Polypropylene 890 58 20P7 N.i.

16 16P5 Polyethylene 837 58 20P8 N.i.

16 16P6 Polyethylene 725 58 20P9 N.i.

16 16P7 Polyethylene 815 58 20P10 N.i.

16 16P8 Chitin 449 59 21P1 N.i.

17 17P1 Polypropylene 695 59 21P2 N.i.

17 17P2 Polyethylene 985 59 21P3 N.i.

17 17P3 Polyethylene 997 59 21P4 N.i.

17 17P4 Polyethylene 541 59 21P5 N.i.

17 17P5 Polypropylen 701 60 22P1 N.i.

17 17P6 Polyethylene 800 60 22P2 N.i.

17 17P7 Polypropylen 422 60 22P3 N.i.

17 17P8 Polyethylene 995 60 22P4 N.i.

17 17P9 Polyethylene 993 61 23P1 N.i.

17 17P10 Polyethylene 837 61 23P2 N.i.

18 18P1 Polyethylene 574 61 23P3 N.i.

18 18P2 Polyethylene 641 61 23P4 N.i.

18 18P3 Polyethylene 726 61 23P5 N.i.

18 18P4 Polyethylene 839 61 23P6 N.i.

18 18P5 Polyethylene 469 61 23P7 N.i.

18 18P6 Polyethylene 994 61 23P8 N.i.

19 19P1 Polystyrene 595 61 23P9 N.i.

21 21P1 Polyethylene 568 61 23P10 N.i.

21 21P2 Polyethylene 564 Helgoland drift line 2013

SUPPLEMENT CHAPTER III

Table S4: continued

157

21 21P3 Polystyrene 697 63 63P1 Polyethylene 794

21 21P4 Polyethylene 751 63 63P2 Polyvinylchloride 494

22 22P1 Polypropylen 669 63 63P3 Polyethylene 994

22 22P2 Polypropylen 909 63 63P4 Polypropylene 990

22 22P3 Polypropylen 652 63 63P5 Polyethylene 793

23 23P1 Polystyrene 669 63 63P6 Polypropylene 929

23 23P2 Polyethylene 994 63 63P7 Polypropylene 605

26 26P1 Keratin 550 63 63P8 Polyamide 917

29 29P1 Chitin 529 63 63P9 Polyethylene 903

30 30P1 Polypropylene 994 63 63P10 Polyurethane 311

32 32P1 Polyethylene 724 63 63P11 Polyethylene 773

32 32P2 Keratin 364 63 63P12 Varnish 175

33 33P1 Keratin 285 63 63P13 Polyvinylchloride 478

35 35P1 Keratin 654 63 63P14 Polystyrene 994

35 35P2 Keratin 460 63 63P15 Polyethylene 999

36 36P1 Keratin 597

37 37P1 Polyethylene 658

37 37P2 Polyethylene 724

37 37P3 Polyethylene 726

38 38P1 N.i. 299

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Table S5: MALDI-TOF Vibrio identification results & species-specific and virulence-associated-gene PCR results

(+ = positive, - = negative) of V. parahaemolyticus obtained from microplastic samples. Meaning of MALDI HIT-

Score: 2.300-3.000 highly probable species identification, 2.000-2.299 secure genus – probable species

identification, 1.700-1.999 probable genus identification, 0.000-1.699 not reliable identification.

Station

No. Particle No.

Polymer

type

Isolate

label

Species

identification

MALDI

HIT-Score toxR tdh trh

63 63P1 PE 1A V. parahaemolyticus 2,59 + - -

63 63P4 PP 4B V. parahaemolyticus 2,61 + - -

63 63P6 PP 6A V. parahaemolyticus 2,58 + - -

63 63P9 PE 9A V. parahaemolyticus 2,62 + - -

5 5P2 PE VN-4252 V. parahaemolyticus 2,40 + - -

5 5P2 PE VN-4253 V. parahaemolyticus 2,55 + - -

9 9P3 PE VN-4225 V. parahaemolyticus 2,62 + - -

11 11P1 PE VN-4229 V. fluvialis 2,53

21 21P2 PE VN-4237 V. parahaemolyticus 2,54 + - -

30 30P1 PP VN-4239 V. parahaemolyticus 2,62 + - -

30 30P1 PP VN-4240 V. fluvialis 2,55

39 39P3 PS VN-3234 V. parahaemolyticus 2,47 + - -

41 41P1 PE VN-3228 V. parahaemolyticus 2,22 + - -

41 41P3 PE VN-3231 V. parahaemolyticus 2,38 + - -

41 41P4 PE VN-3225 V. parahaemolyticus 2,42 + - -

41 41P6 PE VN-3232 V. alginolyticus 2,35

55 55P2 PE VN-3227 V.spp 1,91

55 55P2 PE VN-3229 V.spp 1,96

58 58P9 NI VN-3224 V. fluvialis 2,57

58 58P10 NI VN-3226 V. fluvialis 2,44

58 58P7 NI VN-3233 V. fluvialis 2,53

59 59P4 NI VN-3230 V. fluvialis 2,28

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159

Table S6: MALDI-TOF Vibrio identification results & species-specific and virulence-associated-gene PCR results

(+ = positive, - = negative) of V. parahaemolyticus, V. cholerae and V. vulnificus obtained from water samples.

Meaning of MALDI HIT-Score: 2.300-3.000 highly probable species identification, 2.000-2.299 secure genus –

probable species identification, 1.700-1.999 probable genus identification, 0.000-1.699 not reliable identification.

Station

No.

Isolate

label

Species

identification

MALDI

HIT-Score toxR tdh trh O1 O139 ctxA

1 VN-4208 V. diazotrophicus 2,47

1 VN-4209 V. diazotrophicus 2,54

2 VN-4210 V. vulnificus 2,62 +

2 VN-4211 V. vulnificus 2,60 +

3 VN-4212 V. parahaemolyticus 2,61 + + -

3 VN-4213 V. parahaemolyticus 2,64 + - -

4 VN-4214 V. mimicus 2,56

4 VN-4215 V. mimicus 2,57

5 VN-4216 V. cholerae 2,64 + - - -

5 VN-4217 V. parahaemolyticus 2,67 + - -

5 VN-4218 V. parahaemolyticus 2,62 + - -

6 VN-4219 V. cholerae 2,52 + - - -

6 VN-4231 V. cholerae 2,52 + - - -

6 VN-4220 V. parahaemolyticus 2,67 + - -

8 VN-4221 V. vulnificus 2,62 +

8 VN-4222 V. fluvialis 2,50

9 VN-4223 V. cholerae 2,62 + - - -

9 VN-4224 V. parahaemolyticus 2,64 + - -

10 VN-4226 V. cholerae 2,65 + - - -

10 VN-4227 V. parahaemolyticus 2,55 + - -

11 VN-4228 V. parahaemolyticus 2,56 + - -

12 VN-4230 V. parahaemolyticus 2,61 + - -

12 VN-4254 V. mimicus 2,41

12 VN-4255 V. parahaemolyticus 2,70 + - -

12 VN-4261 V. cholerae 2,63 + - - -

13 VN-4262 V. parahaemolyticus 2,67 + - -

13 VN-4263 V. parahaemolyticus 2,72 + - -

16 VN-4256 V. vulnificus 2,52 +

16 VN-4243 V. diazotrophicus 2,43

17 VN-4257 V. fluvialis 2,53

17 VN-4264 V. mechnikovii 2,26

18 VN-4258 V. fluvialis 2,57

19 VN-4265 V. parahaemolyticus 2,68 + - -

19 VN-4259 V. parahaemolyticus 2,63 + - -

20 VN-4232 V. parahaemolyticus 2,67 + - -

21 VN-4233 V. cholerae 2,58 + - - -

21 VN-4234 V. mimicus 2,48

21 VN-4235 V. parahaemolyticus 2,46 +

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Table S6: continued

160

21 VN-4236 V. parahaemolyticus 2,66 + - -

25 VN-4238 V. diazotrophicus 2,45

31 VN-4241 V. cholerae 2,65 + - - -

32 VN-4242 V. fluvialis 2,49

35 VN-4248 V. diazotrophicus 2,61

36 VN-4244 V. vulnificus 2,48 +

36 VN-4245 V. vulnificus 2,43 +

36 VN-4246 V. vulnificus 2,53 +

36 VN-4247 V. cholerae 2,48 + - - -

37 VN-4249 V. vulnificus 2,59 +

38 VN-4250 V. cholerae 2,64 + - - -

38 VN-4251 V. cholerae 2,66 + - - -

39 VN-3253 V. fluvialis 2,43

39 VN-3257 V. parahaemolyticus 2,49 + - -

39 VN-3280 V. vulnificus 2,60 +

40 VN-3268 V. parahaemolyticus 2,41 + - -

41 VN-3265 V. parahaemolyticus 2,60 + - -

42 VN-3255 V. parahaemolyticus 2,34 + - -

42 VN-3266 V. parahaemolyticus 2,24 + - -

42 VN-3245 V.spp 1,78

42 VN-3250 V. spp. 2,21

42 VN-3275 V. parahaemolyticus 2,60 + - -

43 VN-3262 vulnificus 2,34 +

43 VN-3236 V.spp 1,66

43 VN-3251 V. parahaemolyticus 2,49 + - -

43 VN-3252 V. vulnificus 2,48 +

43 VN-3269 V. fluvialis 2,43

44 VN-3244 V. parahaemolyticus 2,47 + - -

45 VN-3282 V. parahaemolyticus 2,66 + - -

45 VN-3271 V. vulnificus 2,48 +

47 VN-3261 V. parahaemolyticus 2,41 + - -

48 VN-3277 V. vulnificus 2,33 +

48 VN-3278 V. parahaemolyticus 2,53 + - -

48 VN-3273 V. parahaemolyticus 2,60 + - -

48 VN-3256 V. mimicus 2,63

49 VN-3270 V. fluvialis 2,37

49 VN-3260 V. fluvialis 2,41

49 VN-3239 V. fluvialis 2,37

51 VN-3284 V. fluvialis 2,39

51 VN-3285 V. parahaemolyticus 2,60 + - -

51 VN-3286 V. fluvialis 2,45

51 VN-3272 V. parahaemolyticus 2,50 + - -

51 VN-3263 V. parahaemolyticus 2,32 + - -

51 VN-3274 V. parahaemolyticus 2,45 + - -

51 VN-3259 V. vulnificus 2,41 +

51 VN-3276 V. vulnificus 2,55 +

SUPPLEMENT CHAPTER III

Table S6: continued

161

51 VN-3254 V. parahaemolyticus 2,33 + - -

51 VN-3249 V. parahaemolyticus 2,38 + - -

51 VN-3279 V. vulnificus 2,64 +

51 VN-3246 V. fluvialis 2,45

51 VN-3241 V. mimicus 2,51

51 VN-3242 V. fluvialis 2,38

51 VN-3237 V. mimicus 2,52

52 VN-3240 V. parahaemolyticus 2,44 + - -

52 VN-3247 V. parahaemolyticus 2,64 + - -

52 VN-3264 V. vulnificus 2,55 +

52 VN-3287 V. fluvialis 2,60

53 VN-3258 V. parahaemolyticus 2,69 + - -

58 VN-3235 V. fluvialis 2,42

58 VN-3281 V. fluvialis 2,53

58 VN-3288 V. fluvialis 2,58

62 VN-3267 V. fluvialis 2,58

62 VN-3238 V. fluvialis 2,50

62 VN-3283 V. fluvialis 2,43

62 VN-3243 V. fluvialis 2,52

62 VN-3248 V. fluvialis 2,42

Supplementary material for Future Perspectives

Detailed information about isolation of plastic-associated bacteria and fungi, including media

preparation, enrichment, isolation, dereplication, DNA extraction and sequencing is provided.

Furthermore, two tables give information about the taxonomic classification of representative

bacterial and fungal strains.

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Isolation of plastic-associated bacteria and fungi

Medium preparation

Artificial seawater and all media were prepared with sterile filtered (0.2 µm polycarbonate

filter) ultrapure water (Millipore, Germany). Artificial seawater was prepared as described by

(Winkelmann and Harder, 2009) containing the following basal salts dissolved in 1 l ultrapure

water: 26.37 g NaCl, 0.19 g NaHCO3, 1.47 g CaCl2 x 2 H2O, 0.72 g KCl, 0.10 g KBr, 0.02 g

H3BO3, 0.02 g SrCl2 0.003 g NaF. The artificial seawater was autoclaved at 121°C for 25 min,

passively cooled to room temperature and supplemented with 1 ml sterile filtered SeW (Widdel

and Bak, 1992) solution and 2 ml autoclaved trace element solution containing 2.1 g FeSO4 x

7 H2O, 5.2 g Na2-EDTA, 30 mg H3BO3, 100 mg MnCl2 x 4 H2O, 190 mg CoCl2 x 6 H2O, 24 mg

NiCl2 x 6 H2O, 10 mg CuCl2 x 2H2O, 144 mg ZnSO4 x 7 H2O, 36 mg Na2 MoO4 x 2 H2O per

litre ultrapure water and the pH was adjusted to 6.0 with 5 M NaOH (Pfennig and Trüper, 1981).

For the enrichment HaHa_100 medium (Hahnke et al., 2015) and magnesium subtracted

HaHa_100-Mg medium was used. Therefore the artificial seawater was supplemented with 7.9

ml autoclaved MgCl2 x 6 H2O (500 g l-1), 9.5 ml MgSO4 x 7 H2O (500 g l-1), with 10 ml

autoclaved KH2PO4 (2 g l-1), 4 ml NH4Cl (0.2 g l-1) and the sterile filtered carbon sources

glucose, cellobiose, yeast extract, casamino acids and typtone – peptone at a concentration of

0.1 g l-1 each providing a final concentration of 16.8 mM organic carbon. The magnesium

subtracted HaHa_100-Mg was supplemented with the same ammonium, phosphate and carbon

sources but without magnesium sources. The HaHa_100 agar was prepared as described

previously by (Hahnke et al., 2015) with slight modifications. Washed agar (18 g l-1, BactoTM)

and artificial seawater were mixed and autoclaved at 121°C for 25 min in conventional glass

bottles. The medium was passively cooled to 55°C and then supplemented with sterile filtered

HEPES (50 mM, pH 7.5). The HaHa_Hexane agar was prepared in the same way but without

the carbon sources. In lieu thereof 200 µl of n-hexane (86.18 g/mol) were add on a sterile filter

cellulose-nitrate filter (Sartorius) in a sterile petri dish and immediately overlaid with the

“carbon free” HaHa_100 agar.

For the enrichment of plastic-associated fungi Wickerham medium consisting of 10 g glucose

x H2O, 5 g soya peptone, 3 g malt extract, 3 g yeast extract and 30 g NaCl dissolved in 1 l ultra

pure water was used. As solid medium glucose-peptone-yeast extract agar consisting of 1 g

glucose x H2O, 0.5 g peptone, 0.1 g yeast extract and 15 g agar dissolved in 1 l artificial

seawater (described above) was used. The pH was adjusted to 7.2 – 7.4 with 1 M HCl. Both

SUPPLEMENT FUTURE PERSPECTIVES

165

media were autoclaved at 121°C for 25 min and passively cooled and supplemented with Strep-

Pen (50µg/ml).

Enrichment and isolation of plastic-associated bacteria and fungi

Five synthetic polymers (HDPE, PS, PET, SAN, PESTUR) were chosen for bacterial and fungal

enrichment, glass served as control. For the enrichment of plastic-associated microbes the re-

colonized substrate strips (Chapter II) were transferred into Erlenmyer flasks providing 75ml

HaHa_100 medium (bacterial enrichment cultures) or 75ml Wickerham medium (fungal

enrichment cultures) and incubated shaking at 18°C in the dark. After three and five days

respectively dilutions of samples were plated by using Drigalski spatula or Spiral-plater

(easySpiral® Dilute; Interscience, France) on HaHa_100, HaHa_Hexane or glucose-peptone-

yeast extract agar. All inoculated agar plates were incubated at 18°C in the dark and daily

screened for growth. The appearing colonies were checked with respect to distinct colony

colorations, shape and size. Ten representative colonies of each colony type were picked and

differentially streaked out on respective medium and incubated under same conditions.

De-replication by MALDI-TOF MS of unveiled bacteria

For rapid de-replication all isolates grown on HaHa-medium were measured in triplicate by

Intact-Cell MALDI-TOF mass spectrometry as previously described by (Dieckmann et al.,

2005). All isolates were analysed via the direct transfer procedure according to manufacturers`

recommendations (Bruker Daltonics Inc., Germany, Bremen). This involved picking colonies

with sterile toothpicks which were directly spotted onto the target plate (MSP 96 target polished

steel) as thin layer. Each sample spot was overlaid with 1 µl formic acid (70% v/v) followed by

an overlay with 1 µl matrix solution (saturated solution of α-cyano-4-hydroxycinnamic acid in

50% acetonitrile and 2.5% trifluoroacetic acid) and allowed to air dry prior to analysis. Mass

spectra were aquired using the microflex LT/SH system (Bruker Daltonics Inc., Germany,

Bremen). All generated mass spectra were first processed with the baseline correction and

smoothed with the MALDI BiotyperTM software (Bruker Daltonics Inc., Germany, Bremen,

version 3.1). Cluster analysis (PCA dendrogram) was performed based on the comparison of

the resulting spectra of the isolates analysed. The parameter settings were, distance measure

Euclidian, linkage complete, and a cuttoff of 2. In the first step, each created spectrum of the

dataset was compared with each of the other spectra resulting in a PCA dendrogram with main

and sub-clusters. Based on this dendrogram ten, if possible, representative isolates of each

SUPPLEMENT FUTURE PERSPECTIVES

166

substrate type and from each sub-cluster were chosen for further analysis. In the second step,

proteins of the representative strains were extracted using a previously described formic

acid/acetonitrile extraction method (Mellmann et al., 2008) to create high quality mass spectra.

Those were clustered against each other (Fig S1). In order to check the reliability of the cluster

assignment via Intact-Cell MALDI-TOF MS five generated spectra of different isolates of V.

cholerae, V. vulnificus and V. parahaemolyticus each were included in the cluster analysis (Fig

S1).

SUPPLEMENT FUTURE PERSPECTIVES

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Fig

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SUPPLEMENT FUTURE PERSPECTIVES

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DNA extraction and Sanger Sequencing of bacterial and fungal isolates

DNA extraction of the bacterial isolates de-replicated by MALDI-TOF MS and for the fungal

isolates IK_Pi68, IK_Pi70, IK_Pi74 and IK_Pi75 was carried out using lysozyme/SDS lysis

and phenol/chloroform extraction, followed by isopropanol precipitation as described

previously by Oberbeckmann et al. (2011a). DNA extraction of the fungal isolates IK_Pi03,

IK_Pi05, IK_Pi10, IK_Pi11, IK_Pi12, IK_Pi14, IK_Pi07 and IK_Pi13 was carried out using

DNA extraction kit (Power biofilm MoBio Laboratories, Inc.). Prior to PCR experiments, DNA

quantity and quality was determined photometrically (TECAN infinite M200, Switzerland).

PCR was performed with the primer 27F (5`-AGA GTT TGA TCC TGG CTC AG-3`)

(Weisburg et al., 1991), 1492R (5`-GGT TAC CTT GTT ACG ACT T-3`) (Suzuki and

Giovannoni, 1996). The 18S gene was amplified using the fungal-specific primerset Euk-1A

(5′-AACCTGGTTGATCCTGCCAGT-3) (Medlin et al., 1988) and FR1 (5′-

AICCATTCAATCGGTAIT-3′) (Vainio and Hantula, 2000). Amplified PCR products were

purified using QiaQuick reagents (Qiagen, Germany) and PCR products were then sequenced

using Sanger sequencing techniques at Qiagen Genomic Services (Hilden, Germany).

Sequencing was performed by the use of the primersets 27F (5`-AGA GTT TGA TCC TGG

CTC AG-3`) (Weisburg et al., 1991), 1492R (5`-GGT TAC CTT GTT ACG ACT T-3`) (Suzuki

and Giovannoni, 1996) and 907R (5′-CCG TCA ATT CCT TTR AGT TT-3′) (Lane et al., 1985)

for bacteria, and Euk-1A (5′-AACCTGGTTGATCCTGCCAGT-3) (Medlin et al., 1988) and

FR1 (5′-AICCATTCAATCGGTAIT-3′) (Vainio and Hantula, 2000) for fungi.

All rRNA sequences were submitted to ENA via the GFBio data submission service. The

prokaryotic 16S and 18S sequences are available under the accesison numbers LR218064-

LR218111 and LR536736-LR536747.

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Table S1 Bacterial Isolates. Taxonomic classification of representative isolates after MALDI-TOF MS de-

replication, based on 16S sequence analysis (BLAST).

Sample ID Substrate Closest relative (BLAST) Class

Accession

Number

IK_P42 PS Thalassospira lucentensis VBW014 Alphaproteobacteria KC534149.1

IK_P67 HDPE Thalassospira sp. DG1243 Alphaproteobacteria DQ486488.1

IK_P38 Glass Thalassospira lohafexi 139Z-12 Alphaproteobacteria NR_136875.1

IK_P40 PESTUR Thalassospira lohafexi 139Z-12 Alphaproteobacteria NR_136875.1

IK_P71 PESTUR Thalassospira lucentensis VBW014 Alphaproteobacteria KC534149.1

IK_P41 PET Thalassospira sp. DG1243 Alphaproteobacteria DQ486488.1

IK_P69 SAN Thalassospira lucentensis VBW014 Alphaproteobacteria KC534149.1

IK_P39 HDPE Thalassospira lucentensis VBW014 Alphaproteobacteria KC534149.1

IK_P44 Glass Marinobacter sp. NBRC 101711 Gammaproteobacteria AB681536.1

IK_P45 PET Marinobacter sediminum R65 Gammaproteobacteria NR_029028.1

IK_P92 SAN Marinobacter similis A3d10 Gammaproteobacteria KJ547704.1

IK_P77 HDPE Marinobacter salarius R9SW1 Gammaproteobacteria KJ547705.1

IK_P36 PS Pseudoalteromonas carrageenovora NBRC 12985 Gammaproteobacteria NR_113605.1

IK_P01 Glass Alteromonas stellipolaris LMG 21861 Gammaproteobacteria CP013926.1

IK_P30 PESTUR Alteromonas stellipolaris PQQ-44 Gammaproteobacteria CP015346.1

IK_P59 HDPE Alteromonas stellipolaris LMG 21856 Gammaproteobacteria CP013120.1

IK_P64 SAN Alteromonas stellipolaris PQQ-42 Gammaproteobacteria CP015345.1

IK_P32 PET Alteromonas stellipolaris LMG 21861 Gammaproteobacteria CP013926.1

IK_P54 SAN Alteromonas stellipolaris Gammaproteobacteria CP015346.1

IK_P15 PS Muricauda ruestringensis DSM 13258 Flavobacteria NR_074562.1

IK_P17 HDPE Sporosarcina sp. NBRC 100704 Firmicutes AB681231.1

IK_P24 PET Sporosarcina sp. Lc50-2 Firmicutes GU733475.1

IK_P06 SAN Sporosarcina sp. Lc50-2 Firmicutes GU733475.1

IK_P07 SAN Sporosarcina sp. NBRC 100704 Firmicutes AB681231.1

IK_P03 PESTUR Paenisporosarcina sp. Firmicutes JX949201.1

IK_P05 PS Sporosarcina sp. DRB15 Firmicutes JF778686.1

IK_P02 HDPE Sporosarcina sp. Lc50-2 Firmicutes GU733475.1

IK_P04 PET Sporosarcina sp. NBRC 100704 gene Firmicutes AB681231.1

IK_P79 HDPE Sporosarcina sp. NBRC 100704 Firmicutes AB681231.1

IK_P20 PET Jeotgalibacillus marinus ATCC 29841 Firmicutes NR_112057.1

IK_P22 SAN Jeotgalibacillus marinus 581 Firmicutes NR_025351.1

IK_P09 PESTUR Jeotgalibacillus marinus ATCC 29841 Firmicutes NR_112057.1

IK_P66 PS Micrococcus luteus JGTA-S5 Actinobacteria KT805418.1

IK_P91 HDPE Sulfitobacter sp. S11-B-4 Alphaproteobacteria EU016167.1

IK_P11 Glass Celeribacter baekdonensis L-6 Alphaproteobacteria NR_117908.1

IK_P76 PESTUR Celeribacter sp. Ar-141 Alphaproteobacteria JX844513.1

IK_P13 PET Celeribacter sp. Ar-141 Alphaproteobacteria JX844513.1

IK_P88 PET Celeribacter sp. R-52665 Alphaproteobacteria KT185135.1

IK_P83 PESTUR Celeribacter sp. Ar-141 Alphaproteobacteria JX844513.1

IK_P81 SAN Celeribacter sp. Ar-141 Alphaproteobacteria JX844513.1

IK_P12 PET Bacillus sp. KSM-KP43 Firmicutes AB055093.1

IK_P14 PS Bacillus sp. KSM-KP43 Firmicutes AB055093.1

IK_P55 PS Bacillus halmapalus Firmicutes LN867283.1

IK_P48 HDPE Bacillus sp. KSM-KP43 Firmicutes AB055093.1

IK_P47 Glass Bacillus sp. KSM-KP43 Firmicutes AB055093.1

IK_P49 PESTUR Bacillus sp. B055-44 Firmicutes KJ191007.1

IK_P65 SAN Bacillus sp. JSM 101020 Firmicutes KM199862.1

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Table S2 Fungal Isolates. Taxonomic classification of fungal strains based on phylogenetic analysis using ARB®.

The used tree (FungiRef111.newick) includes tree-based taxonomic information and 9366 sequences (Yarza et al.,

2017).

Sample ID Substrate Phylum Class

IK_Pi68 HDPE Basidiomycota Tremellomycetes

IK_Pi70 PS Basidiomycota Cystobasidiomycetes

IK_Pi74 PS Basidiomycota Microbotryomycetes

IK_Pi75 PESTUR Basidiomycota Microbotryomycetes

IK_Pi3 PESTUR Ascomycota Leotiomycetes

IK_Pi5 Glass Ascomycota Sordariomycetes

IK_Pi7 Glass Ascomycota Eurotiomycetes

IK_Pi10 PESTUR Ascomycota Eurotiomycetes

IK_Pi11 Glass Basidiomycota Exobasidiomycetes

IK_Pi12 Glass Ascomycota Eurotiomycetes

IK_Pi13 Glass Ascomycota Eurotiomycetes

IK_Pi14 HDPE Ascomycota Eurotiomycetes

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ACKNOWLEDGEMENTS

187

ACKNOWLEDGEMENTS

First and foremost, I would like to thank the members of my PhD Thesis committee, PD Dr.

Bernhard Fuchs (Max Planck Institute for Marine Microbiology), Dr. Georg Krohne

(University Würzburg), Dr. Gunnar Gerdts, and Dr. Antje Wichels (Alfred Wegener Institute

Helmholtz Centre for Polar Marine Research) for their continuous and excellent supervision,

for all the fruitful discussions and valuable inputs which guided my work.

I would like to thank the Alfred-Wegener-Institute Helmholtz Center for Polar and Marine

Research for funding my PhD project. I am also grateful for scientific and financial support

from the International Max Planck Research School of Marine Microbiology (MarMic).

Especially, I would like to thank Dr. Christiane Glöckner for her assistance with all MarMic

related issues.

I am truly grateful for the privilege of learning so much and for meeting so many people who

encouraged and inspired me on my PhD journey. In particular, I want to express my gratitude

to my direct supervisors Gunnar Gerdts and Antje Wichels, for their scientific guidance,

patience, trust, and support, and for helping me to find my way. I also want to thank you for

your understanding and your kindness, when life was though and I needed it most.

I would like to thank all the colleagues on Helgoland for the good and constant cheerful working

atmosphere and for all the amazing moments we shared. I am grateful to Prof. Dr. Maarten

Boersma for his support and for enabling me to broaden my scientific expertise by working on

a project not related to my PhD thesis. Thanks to all former and current members of the working

group “Microbial Ecology” for all the support and fruitful discussions. Especially, I want to

thank Hilke Döpke and the master students Maike, Lizzy, Tabea and Merle for all the assistance,

and for their contribution and support in the lab.

Special thanks to my “Pizza Pizza girls” – Ale, Sidi, Claudi & Birte – for all the time we shared,

for all the discussions we had, for sharing the load of life, and for their endless motivational

support. Meeting so many amazing people on this tiny island broadened my mind and made my

PhD time very special. Thank you so much for the countless hours we spent together laughing,

talking, dancing, cooking, and savouring the moment.

ACKNOWLEDGEMENTS

188

I am grateful to all my dear mainland friends for their support during my studies and for proving

that distance means so little when someone means so much. Thank you for helping me out so

many times in so many ways. I would like to thank Maike for such being a good friend and for

being my mainland base in the past years during every North Sea storm event. Björn and Jani,

thank you for our long-lasting friendship over the past two decades and for believing in me.

This always kept me going.

Ced, my partner, friend, and colleague, thank you for all the love, moral and professional

support you have given me during the last years. Thank you for your constant help, including

proof reading, all the feedback and fruitful discussions, but also for listening over and over

again to all my ideas and thoughts. I will be always hugely grateful for you being there, not

only for the good times, but also that in dark and difficult times you always found the right way

to encourage me. Words are not enough to express you how grateful I am to have you in my

life.

Name: Inga Vanessa Kirstein Ort, Datum: Bremen, den 11. März 2019

Anschrift: Jadestr. 566, 27498 Helgoland

ERKLÄRUNG

Hiermit erkläre ich, dass ich die Doktorarbeit mit dem Titel:

It`s all about the base

Marine biofilms in the plastic age

selbstständig verfasst und geschrieben habe und außer den angegebenen Quellen keine weiteren Hilfsmittel verwendet habe. Ebenfalls erkläre ich hiermit, dass es sich bei den von mir abgegebenen Arbeiten um drei identische Exemplare handelt.

_______________________________

(Unterschrift)