it`s all about the base marine biofilms in the plastic age
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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
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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
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(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
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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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40
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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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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77
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
GENERAL DISCUSSION
83
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
GENERAL DISCUSSION
84
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
GENERAL DISCUSSION
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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
GENERAL DISCUSSION
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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.
GENERAL DISCUSSION
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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
GENERAL DISCUSSION
88
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
GENERAL DISCUSSION
89
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).
GENERAL DISCUSSION
90
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
GENERAL DISCUSSION
91
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
GENERAL DISCUSSION
92
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
GENERAL DISCUSSION
93
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.
FUTURE PERSPECTIVES
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.
SUPPLEMENT CHAPTER I
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
114
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
115
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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3
1.3
5
Cyt
op
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
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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).
SUPPLEMENT CHAPTER II
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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
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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
SUPPLEMENT CHAPTER III
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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
SUPPLEMENT CHAPTER III
158
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
SUPPLEMENT CHAPTER III
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 +
SUPPLEMENT CHAPTER III
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
167
Fig
S1
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ial Is
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e th
ree
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. ch
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SUPPLEMENT FUTURE PERSPECTIVES
168
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
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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)
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