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Life in the “Plastisphere”: Microbial Communities on Plastic Marine
Debris
Erik R. Zettler,
†,∥
Tracy J. Mincer,
‡,
*
,∥
and Linda A. Amaral-Zettler
§,
*
,∥
†
Sea Education Association, P.O. Box 6, Woods Hole, Massachusetts 02543, United States
‡
Marine Chemistry & Geochemistry, Woods Hole Oceanographic Institution, 266 Woods Hole Rd., MS#51, Woods Hole,
Massachusetts 02543, United States
§
The Josephine Bay Paul Center for Comparative Molecular Biology and Evolution, Marine Biological Laboratory, 7 MBL Street,
Woods Hole, Massachusetts 02543, United States
*
SSupporting Information
ABSTRACT: Plastics are the most abundant form of marine
debris, with global production rising and documented impacts
in some marine environments, but the influence of plastic on
open ocean ecosystems is poorly understood, particularly for
microbial communities. Plastic marine debris (PMD) collected
at multiple locations in the North Atlantic was analyzed with
scanning electron microscopy (SEM) and next-generation
sequencing to characterize the attached microbial commun-
ities. We unveiled a diverse microbial community of
heterotrophs, autotrophs, predators, and symbionts, a
community we refer to as the “Plastisphere”. Pits visualized
in the PMD surface conformed to bacterial shapes suggesting
active hydrolysis of the hydrocarbon polymer. Small-subunit
rRNA gene surveys identified several hydrocarbon-degrading bacteria, supporting the possibility that microbes play a role in
degrading PMD. Some Plastisphere members may be opportunistic pathogens (the authors, unpublished data) such as specific
members of the genus Vibrio that dominated one of our plastic samples. Plastisphere communities are distinct from surrounding
surface water, implying that plastic serves as a novel ecological habitat in the open ocean. Plastic has a longer half-life than most
natural floating marine substrates, and a hydrophobic surface that promotes microbial colonization and biofilm formation,
differing from autochthonous substrates in the upper layers of the ocean.
■INTRODUCTION
Plastic has become the most common form of marine debris
since it entered the consumer arena less than 60 years ago, and
presents a major and growing global pollution problem.
1−3
The
current global annual production, estimated at 245 million
tonnes
1
represents 35 kg of plastic produced annually for each
of the 7 billion humans on the planet, approximating the total
human biomass. Some fraction of the increasing amount of
postconsumer plastic trash inevitably escapes the recycling and
waste streams and makes its way to the global oceans.
Additionally, tsunamis and storms can result in large pulses
of plastic entering the ocean from coastal areas. Plastic
accumulates not only on beaches worldwide, but also in
“remote”open ocean ecosystems.
1
Drifter buoys and physical
oceanographic models have shown that surface particles such as
PMD can passively migrate from Eastern Seaboard locations all
the way to the interior of the North Atlantic Subtropical Gyre
in less than 60 days,
4
illustrating how quickly human-generated
debris can impact the gyre interior that is more than 1000 km
from land. Plastic debris in the North Atlantic Subtropical
Gyre
4
and North Pacific Subtropical Gyre is well-docu-
mented
5−9
and models and limited sampling confirm that
accumulations of PMD have formed in all five of the world’s
subtropical gyres.
10,11
The effects of plastic debris on animals such as fish, birds, sea
turtles, and marine mammals as a result of ingestion,
12−15
and
marine entanglement
3,16−18
are well documented, but studies of
plastic-associated microbial communities are lacking, and we
know little about the impact of this anthropogenic substrate
and its attached community on the oligotrophic open ocean. As
a relatively new introduction into the marine ecosystem, plastic
debris provides a substrate for microbes that lasts much longer
than most natural floating substrates and has been implicated as
a vector for transportation of harmful algal species
19
and
persistent organic pollutants (POPs).
20,21
With a hydrophobic
surface rapidly stimulating biofilm formation in the water
column, PMD can function as an artificial “microbial reef”.
PMD at concentrations of up to 5 ×105pieces/km2in the
North Atlantic Subtropical Gyre
4
represents a new floating
Received: March 26, 2013
Revised: May 26, 2013
Accepted: June 7, 2013
Published: June 7, 2013
Article
pubs.acs.org/est
© 2013 American Chemical Society 7137 dx.doi.org/10.1021/es401288x |Environ. Sci. Technol. 2013, 47, 7137−7146
substrate for microbial colonization and transportation, and the
presence of particles in aquatic systems is known to stimulate
microbial productivity and respiration.
22−25
Once trapped in
central ocean gyres, there are very few avenues for export, and
buoyant plastic particles accumulate and may persist for
decades. Increases in PMD have been documented in the
North Pacific Gyre,
6
but despite increases in plastic production,
use, and presumably input into the ocean, other studies show
no significant trend in plastic accumulation in the North
Atlantic Subtropical Gyre
4
or in the waters from the British
Isles to Iceland since the 1980s.
26
Physical shearing and
photodegradation are proposed mechanisms of plastic degra-
dation.
27,28
These physical mechanisms may result in
fragmentation into pieces small enough to pass through
standard sampling nets.
29
In addition, biofilm formation and
colonization by invertebrates can decrease plastic buoyancy
allowing some of the plastic debris to sink to deeper waters,
with eventual seafloor deposition.
1,30
However, plastic debris is
absent from sediment traps
4
suggesting that density-mediated
transport of small pieces is relatively low.
PMD has been reported by a number of studies starting in
the 1970s
31,32
where authors mention diatoms and other
microbes on the debris. However, our study presents the first
comprehensive characterization of microbial communities living
on PMD in the open ocean with an emphasis on bacteria. We
hypothesized that this new man-made substrate is physically
and chemically distinct from surrounding seawater and
naturally occurring substrates such as macroalgae, feathers,
and wood, with the potential to select for and support distinct
microbial communities. Using pyrotag sequencing and SEM,
we investigated representative microbial communities on pieces
of polyethylene (PE) and polypropylene (PP) plastic from
geographically distinct areas from the North Atlantic
Subtropical Gyre and compared them to the microbial
communities in the surrounding seawater. Our analyses
unveiled for the first time the breadth of PMD microbes that
make up what we call the “Plastisphere”.
■EXPERIMENTAL SECTION
Sample Collection. Plastics were collected in a 1 ×0.5 m
rectangular neuston net with 333-μm mesh towed at the surface
from the Sea Education Association (SEA) vessel SSV Corwith
Cramer as part of SEA Semester research cruises C-230 and C-
241 (Supporting Information (SI) Table S1). Individual pieces
of plastic were sorted with sterile forceps and rinsed with sterile
seawater prior to subdivision using a sterile razor blade and
preservation for DNA extraction and SEM. While the net was
in the water, we filtered 4 L from a clean seawater system
(periodically freshwater-flushed nonmetallic line drawing water
from 3 m below the surface) through a 0.2 μm Sterivex
cartridge filter (Millipore, Billerica, MA) to collect micro-
organisms suspended in the ambient surface water.
Sample Preservation. Plastic and seawater filters for
downstream DNA analysis were immediately flooded with
Puregene lysis buffer (Qiagen, Valencia, CA) and frozen at −20
°C. PMD samples for SEM were fixed in 4% paraformaldehyde
for 2−23 h, then transferred to 50% ethanol in Phosphate
Buffered Saline (PBS) and kept at −20 °C.
SEM. Preserved plastic samples for SEM were dehydrated on
ice through an ethanol series: 10 min each in 50%, 70%, 85%,
95%, followed by 3 ×15 min in 100% ethanol. Samples were
immediately critical point dried using a Samdri 780A
(Tousimis, Rockville, MD), sputter coated with 5 nm of
platinum using a Leica EM MED020 (Leica Microsystems, Inc.
Buffalo Grove, IL), then visualized and imaged on a Zeiss Supra
40VP SEM (Carl Zeiss Microscopy, Thornwood, NY). Cell
measurements were made from digital images using ImageJ
software (Rasband, W.S., ImageJ, U.S. National Institutes of
Health, Bethesda, MD http://imagej.nih.gov/ij/, 1997-2012).
Raman Spectroscopy. The resin composition of plastic
pieces was identified using a PeakSeeker Pro Raman
spectrometry system (Agiltron, Woburn, MA) with a micro-
scope attachment that enabled measurement of spectra from
very small pieces of plastic. Each sample was compared with
reference scans from plastics of known composition. We
selected three pieces of polypropylene and three pieces of
polyethylene that were large enough to subdivide for SEM and
DNA extraction.
Amplicon Pyrotag Sequencing. DNA was extracted
using a modified bead-beating approach
33
in combination
with the Puregene Tissue DNA extraction kit (Qiagen,
Valencia, CA). We amplified bacterial V6−V4 hypervariable
regions of the small-subunit rRNA (SSU rRNA) gene using
primers targeting Escherichia coli positions 518 and 1046.
34
We
multiplex-sequenced the resulting amplicons with a barcoded
primer strategy
35
on a 454 Genome Sequencer FLX (Roche,
Basel, Switzerland) using the manufacturer’s suggested protocol
for the GS-FLX-Titanium platform. We trimmed sequences of
adapter and primer sequences and removed low-quality reads as
described previously.
36
We further filtered low quality base calls
by applying “anchor trimming”to search for conserved V5
priming regions and trimmed to these conserved regions. We
assigned Operational Taxonomic Units (OTUs) to clusters
using the UCLUST v3.0.617 de novo clustering algorithm
37
at
four percent cluster widths (96% similarity) to further minimize
OTU inflation associated with pyrosequencing errors of longer
V6−V4 amplicons. Eukaryotic amplicon sequencing targeted
the V9 hypervariable region and followed protocols established
in Amaral-Zettler et al. (2009).
33
Sequence data are deposited
in NCBI’s Sequence Read Archive (SRP026054) and conform
to the Minimum Information about a MARKer gene Sequence
(MIMARKS) standard (SI Table S2).
38
Data Analyses and Statistical Methods. We used the R
package limma
39
to calculate Venn diagrams and plotted the
resulting figures using the Venn Diagram Plotter (http://ncrr.
pnnl.gov/). Data for subsequent analyses were resampled down
to the lowest number of reads recovered (6,102 reads) to
standardize for sampling effort. R package routines gplots and
heatmap.2 helped to generate the heatmap summary of all
OTUs that were encountered with a frequency of greater than
2% in a given sample. OTU bar graphs were generated using
Global Alignment Sequence Taxonomy (GAST) algorithms
40
and graphical output in QIIME v1.3.0.
41
We used linear
discriminant analysis (LDA) effect size (LEfSe)
42
to identify
biomarkers for plastics versus seawater and substrate specific
(PP vs PE vs seawater) analyses. To ascertain the closest
relative of our dominant Vibrio OTU sequence found on the C-
230 polypropylene substrate (SI Figure S2), we used the quick-
add-sequence-to-tree parsimony feature in the SILVA- 111
reference tree.
43
We then retained only the named type species
to summarize the result. We examined co-occurrence patterns
using network analysis and significant linear Pearson
correlations. For the input matrices we only considered
OTUs that occurred in at least 30% of the samples. We used
Cytoscape (http://www.cytoscape.org) to visualize the result-
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ing network and only considered significant correlations with
an R-value >0.9.
■RESULTS AND DISCUSSION
Identification and Selection of PMD. The majority of
plastic pieces recovered in all net tows were fragments of less
than 5 mm as has been reported in other studies.
29,44
Even the
centimeter-sized pieces chosen to extract DNA and image the
same piece with SEM were fragments without identifiable
markings of undetermined origin. Pieces we examined with
SEM ranged in size from sub-mm diameter monofilament
(Figure 1, piece C230_02) to flat fragments that were several
cm long before subdivision (Figure 1, piece C230_01). All
showed signs of degradation including cracks and pitting as
shown in Figure 2. With the microscope-based Raman
spectrometer, most fragments collected were positively
Figure 1. Raman spectroscopy spectra of the plastics collected from the Sargasso Sea that were imaged and sequenced. The bottom scan on each
panel is of a known standard. Images along the top are light-micrographs of the plastic samples extracted for DNA analyses ((1-mm gradations; note
different magnification on sample C230_01).
Figure 2. SEM images showing examples of the rich microbial community on PMD: (a) pennate diatom on sample C241_07 with possible
prosthecate filaments produced by Hyphomonas-like bacteria; (b) filamentous cyanobacteria on sample C230_01; (c) stalked predatory suctorian
ciliate in foreground covered with ectosymbiotic bacteria (inset) along with diatoms, bacteria, and filamentous cells on sample C230_01; (d)
microbial cells pitting the surface of sample C241_12. All scale bars are 10 μm.
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identified as polyethylene and polypropylene based on spectra
compared to known standards (Figure 1). This was expected
since these two resins are commonly used in packaging and
other single-use plastic applications. They are also less dense
than seawater so consequently float and accumulate in surface
waters.
The Plastisphere Community. Microscopic (phenotypic)
and molecular sequence (genotypic) data provided comple-
mentary evidence for microbial phototrophy, symbiosis,
heterotrophy (including phagotrophy), and predation in our
analyses of PP and PE PMD samples. SEM photomicrographs
revealed the presence of a rich eukaryotic and bacterial
microbiota living on both PP and PE samples (Figure 2, a−
d). Cell counts of random images identified over 50 distinct
morphotypes covering between 0 and 8% of the surface area of
the plastic. Especially intriguing were round cells about 2 μmin
diameter embedded in pits in the surface of the PMD (abstract
image and Figure 2d). Often occurring in rows or patches, the
pits conform closely to the shape of the contained cells, and
included dividing cells that suggest active growth. We have not
identified these cells but they were the third most common
type of morphotype seen, after diatoms and filaments. DNA
Figure 3. Bar chart showing similarity between all three seawater samples and dominance of a relatively small number of abundant OTUs, versus
plastic samples with greater variability between samples and greater evenness indicated by more groups representing smaller proportions of the total
population. The most abundant OTUs are listed as follows: (1) Bacteria, Verrucomicrobia, Verrucomicrobiae, Verrucomicrobiales, Rubritaleaceae,
Rubritalea; (2) Bacteria, Proteobacteria, Gammaproteobacteria, Vibrionales, Vibrionaceae, Vibrio; (3) Bacteria, Proteobacteria, Gammaproteobac-
teria, Pseudomonadales, Moraxellaceae, Psychrobacter; (4) Bacteria, Proteobacteria, Gammaproteobacteria; (5) Bacteria, Proteobacteria,
Alphaproteobacteria, Sphingomonadales, Erythrobacteraceae, Erythrobacter; (6) Bacteria, Proteobacteria, Alphaproteobacteria, Rhodobacterales,
Rhodobacteraceae, Thalassobius; (7) Bacteria, Proteobacteria, Alphaproteobacteria, Rhodobacterales, Rhodobacteraceae; (8) Bacteria,
Proteobacteria, Alphaproteobacteria, Parvularculales, Parvularculaceae, Parvularcula; (9) Bacteria, Proteobacteria, Alphaproteobacteria,
Caulobacterales, Hyphomonadaceae, Hyphomonas; (10) Bacteria, Cyanobacteria, Cyanobacteria, Subsection III, Unassigned, Phormidium; (11)
Bacteria, Cyanobacteria, Cyanobacteria, Subsection III; (12) Bacteria, Bacteroidetes, Sphingobacteria, Sphingobacteriales, Saprospiraceae, Lewinella;
(13) Bacteria, Bacteroidetes, Sphingobacteria, Sphingobacteriales, Flammeovirgaceae, Fulvivirga; (14) Bacteria, Bacteroidetes, Sphingobacteria,
Sphingobacteriales, Chitinophagaceae; (15) Bacteria, Proteobacteria, Deltaproteobacteria, Myxococcales; (16) Bacteria, Chloroflexi, Anaerolineae,
Anaerolineales, Anaerolinaceae; (17) Bacteria, Bacteroidetes, Sphingobacteria, Sphingobacteriales, Saprospiraceae, Saprospira; (18) Bacteria,
Bacteroidetes, Sphingobacteria, Sphingobacteriales, Flammeovirgaceae; (19) Bacteria, Bacteroidetes, Sphingobacteria, Sphingobacteriales,
Flammeovirgaceae, Marinoscillum; (20) Bacteria, Proteobacteria, Gammaproteobacteria, Alteromonadales, Alteromonadaceae, Alteromonas; (21)
Bacteria, Proteobacteria, Alphaproteobacteria, Rhodobacterales, Rhodobacteraceae, Rhodovulum; (22) Bacteria, Proteobacteria, Gammaproteobac-
teria, Oceanospirillales, SAR86; (23) Bacteria, Proteobacteria, Gammaproteobacteria, Alteromonadales, Pseudoalteromonadaceae, Pseudoalter-
omonas; (24) Bacteria, Proteobacteria, Alphaproteobacteria, Rickettsiales, SAR116; (25) Bacteria, Proteobacteria, Alphaproteobacteria, Rickettsiales,
SAR11, Pelagibacter; (26) Bacteria, Proteobacteria, Alphaproteobacteria, Rhodospirillales, Rhodospirillaceae; (27) Bacteria, Bacteroidetes,
Sphingobacteria, Sphingobacteriales, Flammeovirgaceae, Amoebophilus; (28) Bacteria, Bacteroidetes, Sphingobacteria, Sphingobacteriales,
Chitinophagaceae, Sediminibacterium; (29) Bacteria, Bacteroidetes, Flavobacteria, Flavobacteriales, Flavobacteriaceae; (30) Bacteria, Proteobacteria,
Gammaproteobacteria, Oceanospirillales, Oceanospirillaceae, Oceaniserpentilla; (31) Bacteria, Cyanobacteria, Cyanobacteria, Subsection I,
Unassigned, Prochlorococcus; (32) Bacteria, Actinobacteria, Actinobacteria, Acidimicrobiales.
Environmental Science & Technology Article
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sequence analyses confirmed that these communities were
consistently distinct between plastics and the surrounding
seawater. For example, photosynthetic filamentous cyanobac-
teria including Phormidium and Rivularia OTUs occurred on
plastics but were absent from seawater samples where
unicellular Prochlorococcus dominated the bacterial phototroph
community in the seawater samples (Figure 3, SI Figure S1).
The presence of cyanobacteria representing Plectonema-like
genera was also evident in our SEM photomicrographs (Figure
2b).
45
Other conspicuous phototrophs included diatoms
(Figure 2a and background of 2b and 2c) that were assigned
to a number of bacillariophyte genera including Navicula,
Nitzschia,Sellaphora,Stauroneis, and Chaetoceros based on DNA
sequence data. The genera Navicula,Nitzschia, and Sellaphora
are commonly substrate-associated and known biofilm formers
in aquatic environments.
46
In addition to diatoms, representa-
tive OTUs from other protists with known photosynthetic
representatives included prasinophytes, rhodophytes, crypto-
phytes, haptophytes, dinoflagellates, chlorarachniophytes,
chrysophytes, pelagophytes, and phaeophytes.
While we only considered eukaryotic diversity for two of our
samples, there was corroboration between SEM images and
DNA sequence data including the stalked suctorian ciliates
(Figure 2c) covered with bacteria (Figure 2c, inset), and
sequence data confirming that one of the major ciliates was the
genus Ephelota, a marine suctorian known to colonize marine
surfaces and to harbor ectosymbiotic rod-shaped bacteria.
47
The relationship between the ectosymbiotic bacteria and the
Ephelota is unknown, but ectosymbiontic bacteria on the
surface of other stalked ciliates have been identified as sulfide-
oxidizing Gammaproteobacteria
48
of the genus Thiobios, and
we identified sequences corresponding to members of the same
genus in our samples.
Surprisingly, DNA sequences derived from polycystine
colonial radiolaria were present on both plastic types and
dominated one polypropylene sample but were not identified in
our SEM imaging. The discovery of radiolarian OTUs
associated with PMD is somewhat unusual in that they are
planktonic protists that are not understood to be substrate
associated. However, there is precedence for other free-living
taxa such as planktonic foraminifera becoming associated with
PMD.
49
Furthermore, it is highly likely that radiolarians
become passively associated with PMD both in the water
column and in our nets given that colonial forms can reach
meters in length and have a somewhat gelatinous nature. This
may also explain the presence of other traditionally free-living
taxa appearing to be associated, typically at low abundance, with
our PMD substrates (i.e., Pelagibacter,Prochlorococcus).
Heterotrophic bacteria in seawater samples were dominated
by Pelagibacter and other free-living picoplanktonic bacterial
groups
50
but showed very different abundance patterns in the
plastics samples. A striking example was the dominance of a
member of the genus Vibrio that constituted nearly 24% of the
C230_01 polypropylene sample. This is noteworthy because
members of this genus are rarely found in concentrations
greater than 1% of the community
51
although members of the
species V. harveyi are a notable exception.
52,53
To the best of
our knowledge, blooms of vibrios have not been associated with
particles per se although they can dominate phytoplankton and
zooplankton surfaces.
54
Vibrios are also known to have
extremely fast growth rates
55
so this may explain their ability
to dominate members of the Plastisphere on occasion.
Based on its taxonomic placement (SI Figure S2), the Vibrio
sequence we recovered in high abundance on the poly-
propylene plastic sample C230_01 was related to the type
species of V. natriegens, a known nitrogen fixer.
56
However, this
sequence also shared 100% identity with other nontype strain
vibrios assigned to the species V. harveyi,V. alginolyticus,V.
owensii,V. azureus,V. parahemolyticus,V. campbellii,V.
diabolicus,V. communis, and V. rotiferianus, all recent additions
to GenBank.
We are unable to assign our dominant Vibrio OTU to a
specific species based on rRNA sequence data alone so we
cannot rule out the possibility that Plastisphere microbes such
as vibrios could be animal or human pathogens. Plastic could
serve as a vector of infectious diseases since both birds and
fishes ingest PMD and a recent study found fishes contain
human pathogenic Vibrio strains.
57
Because PMD persists
longer than natural substrates (e.g., feathers, wood, and
macroalgae), it can traverse significant distances, and it has
been shown to transport invasive species.
58
Harmful dino-
flagellate species belonging to the genus Alexandrium were
reported from PMD in the Mediterranean,
19
and we also
detected several dinoflagellate species including members of
this genus on our Atlantic PMD. One property that these
groups share is their propensity to adhere to surfaces. In the
case of former reports of HAB-associated PMD, the authors
specifically hypothesize that it was the “sticky”nature of the
vegetative cysts that allowed them to adhere to plastic and may
facilitate their dispersal beyond their typical range.
19
This final
point reiterates the properties of PMD that set it apart from
other types of marine debris: PMD is a selective environment
with hydrophobicity that stimulates early colonizers, rapidly
driving biofilm formation and succession of other microbes.
25
Additionally, the stimulation of microbial respiration and
growth by inert surfaces is a well-characterized phenomenon
in which dilute nutrients are concentrated creating a favorable
environment for microbial colonization.
59
Termed the “ZoBell
effect”(after Claude ZoBell who first thoroughly described the
phenomenon
59
), this concentration of micronutrients by
abundant PMD in oligotrophic areas of the ocean could play
Figure 4. Venn diagram showing bacterial OTU overlap for pooled
PP, PE, and seawater samples; n= number of sequenced reads per
group. Numbers inside the circles represent the number of shared or
unique OTUs for a given substrate.
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a significant role in increasing microbial activity in the upper
layer of ocean gyres.
Alpha Diversity. Over a thousand species equivalents, or
OTUs were observed from single fragments of PE and PP (SI
Table S1). Overall, there were two notable differences between
diversity patterns between the plastics samples and the
surrounding seawater: (1) average observed richness was
much higher in surrounding seawater; but, (2) plastic substrates
showed greater evenness than seawater, that is the community
was not dominated by a small number of abundant organisms.
It is difficult to directly compare richness between seawater and
plastics because of sample size considerations. Seawater samples
had the highest average richness and polyethylene the lowest,
but when normalized with respect to sampling effort (number
of reads recovered), the greatest richness in a single sample was
associated with polypropylene. Although the wide range of
richness values obtained from plastic pieces cautions against
drawing general conclusions about richness and plastics, we
expect richness to be related to substrate area and observed the
highest richness on the largest piece of plastic analyzed (see
image of C230_01 in Figure 1). Evenness, on the other hand,
was consistently higher on plastics (mean Simpson evenness
0.95) compared to seawater (mean Simpson 0.89) and the
brown alga Sargassum (mean Simpson 0.90) (data not shown).
In other words, seawater was characterized by many more rare
taxa that contributed to the richness in these samples.
60,61
This
Figure 5. Taxonomic tree generated using the LEfSe online software highlighting the biomarkers that statistically differentiate PP, PE, and seawater
samples. Circle diameter is proportional to taxon abundance. The tree highlights both high-level (Class) and genera-specific taxonomic trends. Refer
to the legend for substrate color-coding.
Environmental Science & Technology Article
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could be due to a greater abundance of standing stock bacteria
in oligotrophic seawater, known to support a high degree of
rare taxa, partly due to lower grazing and viral pressures.
60
Additionally, a lower richness is expected in the more selective
and metabolically active population of bacteria on the plastic
surfaces supported by a relatively higher nutrient microenviron-
ment. The distinctness of microbial communities from PMD
was also reflected in the percentage of shared OTUs across the
different plastic substrates (Figure 4) and Sargassum (data not
shown). Collectively we found 350 bacterial OTUs shared
between the PE and PP samples. Seawater had the largest
number of unique OTUs (n= 1789), but these were mostly
rare. Seawater shared a minor proportion of its OTUs with PE
(8.6%) and PP (3.5%), respectively. In contrast, OTUs in
common between PP and PE represented a substantial
proportion of their overall OTU assemblage with 40% of the
OTUs shared between PE and PP and 30% of the OTUs
shared between PP and PE. Therefore, Plastisphere commun-
ities, despite being quite variable, do appear to have a “core”of
taxa that characterize them.
Community Membership. To further determine the
membership of the “core”Plastisphere community we
performed biomarker analyses.
42
Linear discriminant analysis
(LDA) effect size (LEfSe) revealed Plastisphere OTUs that
characterized PP and PE samples indicating plastic resins may
select for particular microbial community members. Of
particular interest were OTUs found on both plastics but not
in seawater. These included bacteria documented as capable of
degrading hydrocarbons including the filamentous cyanobacte-
rium Phormidium sp. known to settle on benthic substrates
62
and Pseudoalteromonas, a genus frequently associated with
marine algae
63
(Figure 3, SI Figure S1). Additionally, the
alphaproteobacterial family Hyphomonadaceae, known for
forming long holdfast filaments termed prosthecae (which
were common in our SEM micrographs) were unique to PMD
and comprised almost 8% of the OTUs on PP (SI Figure S1).
Members of the Hyphomonadaceae can be methylotrophic,
known to degrade hydrocarbons and present in hydrocarbon
enrichments.
64,65
Figure 5 summarizes the biomarker results
and highlights the differences between each plastic substrate
and seawater. LDA scores are shown in SI Figure S3.
Network Analyses. We can make inferences about
organism associations from SEM observations of physical
location and community architecture on the plastic surface.
Although many bacteria cannot be identified visually, it is
possible to infer interactions between members of the
Plastisphere indirectly via association networks based on
sequence data.
66−68
We conducted network analyses to further
explore co-occurrence patterns between members of the
Plastisphere. Reporting all existing networks is beyond the
scope of this paper, so we present only networks associated
with putative hydrocarbon-degrading bacteria within our overall
network. Figure 6 depicts these networks with the cyanobacte-
rium Phormidium highlighted as green diamonds, Hyphomonas
as blue diamonds, members of the Chloroflexi as purple
hexagons and members of the Myxococcales as yellow triangles.
The figure only depicts first nearest neighbors in the network
with positive correlations having R> 0.9. Noteworthy were the
co-occurrences of several members of putative hydrocarbon
degrading taxa in close proximity to each other in our network.
Figure 6. Network analysis diagram of putative hydrocarbon degrading bacterial OTUs. The cyanobacterium Phormidium is represented in green
diamonds, Hyphomonas is depicted in blue diamonds, members of the Chloroflexi are shown in purple hexagons and members of the Myxococcales
are represented as yellow triangles. SI Table S3 includes the full taxonomy for all the OTUs in the network.
Environmental Science & Technology Article
dx.doi.org/10.1021/es401288x |Environ. Sci. Technol. 2013, 47, 7137−71467143
In addition to the four aforementioned main groups, we further
detected other potential hydrocarbon-degrading OTUs that
were their nearest neighbors. These included Hyphomonas-
associated OTUs such as Haliscomenobacter (OTU C507,
C300, C2094), associated with hydrocarbon contaminated
soils,
69
Devosia (OTU C137) associated with diesel-contami-
nated soils
70
and Oceaniserpentilla (OTU 1470), one the of
major taxa related to OTUs from the Deepwater Horizon oil
spill.
71
While the presence of these taxa alone does not
implicate them in plastic degradation, our network analyses
suggests that consortia of OTUs may be acting in concert to
utilize this recalcitrant carbon source and provides a testable
hypothesis for future investigation.
PMD age and fate are poorly characterized; despite dramatic
increases in plastic production, a 22-year study in the North
Atlantic Subtropical Gyre showed no evidence of increasing
quantities of PMD collected with a neuston net (333 μm
mesh),
4
implying there must be unrecognized sinks to balance
the sources. A recent review by Hidalgo-Ruz et al.
29
summarizes
much of what is known about PMD and emphasizes the need
for further study of how abiotic and biotic factors break it
down. Our SEM images show microbial cells embedded in pits
in the plastic surface, suggesting that microbes may be taking
part in the degradation of plastic via physical or metabolic
means (Figure 2d and abstract). These types of cells were
found on both PE and PP, and include dividing cells (see image
in abstract). Bacteria and fungi are known to degrade refractile
compounds including plastic
62,63
but this has not been
demonstrated in the open ocean. The pits visualized in PMD
surfaces that conform to the shape of cells growing in the pits,
and sequences of known hydrocarbon degraders support the
possibility that some members of the Plastisphere community
are hydrolyzing PMD and could accelerate physical degrada-
tion. Future research directions include understanding the
genetic mechanisms of how microbes attach to PMD and
elucidating the microbes and genes involved in microbially
mediated plastic degradation through assaying our extensive
culture collection, as well as exploring how these processes
influence interactions with larger organisms.
■ASSOCIATED CONTENT
*
SSupporting Information
Figure S1 shows a heatmap with the relative abundances of the
most abundant taxonomic groups from PP, PE and seawater.
Figure S2 shows an ARB tree with the placement of our most
abundant Vibrio OTU. Figure S3 shows the LDA scores for the
OTUs that explain the greatest differences between seawater,
PP and PE communities. Table S1 provides contextual data for
the samples used in this study including observed richness and
evenness. Table S2 provides a MIMARKS table. Table S3
provides the taxonomy for the network analysis shown in
Figure 6.This material is available free of charge via the Internet
at http://pubs.acs.org.
■AUTHOR INFORMATION
Corresponding Author
*Phone: 508-289-7259 (L.A.A.-Z.); 508-289-3640 (T.J.M.).
Fax: 508-457-4727 (L.A.A.-Z.); 508-289-3640 (T.J.M.). E-mail:
amaral@mbl.edu (L.A.A.-Z.); tmincer@whoi.edu(T.J.M.).
Author Contributions
∥
All authors contributed equally to this work.
Notes
The authors declare no competing financial interest.
■ACKNOWLEDGMENTS
We thank Giora Proskurowski for discussions and collecting
samples under NFWF-NOAA Marine Debris Program Award
No. 2009-0062-002, Amy Siuda for collecting samples, and
Emelia Deforce, Greg Boyd, Sonja Uribe and Catherine Stark
for technical assistance. This work was supported by an NSF
Collaborative grant to E.R.Z. (OCE-1155379), T.J.M. (OCE-
1155671) and L.A.A-Z. (OCE-1155571), NSF TUES grant to
E.R.Z. and L.A.A.-Z. (DUE-1043468), and was partially funded
by the Woods Hole Center for Oceans and Human Health
Pilot award (No. 26291503) to T.J.M.
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