Pishchany, G. Fake plastic planets of the Sippewissett Universe

Fake plastic planets of the Sippewissett Universe
Gleb Pishchany
Microbial Diversity 2013
Marine Biological Laboratory
Introduction
Plastics are high molecular weight moldable organic materials. Throughout the
20th and 21st century humans have increasingly utilized plastic due to its cheapness,
durability, lightweight, and versatility. Massive production, poor recycling efforts, and
slow rates of biodegradation have resulted in contamination of natural environments with
plastics [1-3]. Ironically, while plastics are negatively affecting large ecosystems, they
also create niches for microbial life [4]. This is at least in part due to the hydrophobic
properties of plastic polymers that favor microbial attachment. Because plastic is a rich
potential source of organic carbon, plastic debris may also serve as source of energy for
microorganisms. In this work I wanted to get a glimpse into the communities that
colonize plastic in the environment. I also wanted to investigate how plastic shapes
microbial communities and how microbial communities residing on plastic change their
habitat. Below is the outline of these efforts.
Results and discussion
Samples of plastic were collected from the Little Sippewissett Marsh (figure 1A
and 1B). From here on I will refer to these samples as Bag #1 and Bag #2. To visualize
microbial communities, I stained plastic with DAPI and CARD-FISH probes specific for
detection of Bacteria (Eub I-III) and Eukaryotes (Euk516). A variety of microbes were
detected through these methods with a large presence of both Eukaryotic and Bacterial
cells (Figure 2). Interestingly, Eukaryotic and Bacterial cells were frequently observed in
close proximity to each other each other. Co-localization measurement indicated that
green and red fluorescence did not overlap in the images confirming specificity of the
probes (Figure 2F).
To investigate the surface of plastic colonizers in more detail I performed
scanning electron microscopy. Low magnification images revealed that the surface of
Bag #1 was deteriorating (Figure 3A). Higher magnification revealed the presence of
diverse particles of shapes and sizes consistent with those of living microbial organisms
(Figure 3B-G). These included spheres of various sizes, curved and straight rods, and
cylindrical shapes. Notably some of the smaller microorganisms appeared to be located in
the pits raising the possibility that they were formed through the activity of the microbe
(Figure 3F and 3G). This is consistent with a previous report, noting the pit formation in
plastic particles found in the oceans [4]. Scanning electron micrographs of bag 2 revealed
a dense and diverse microbial community (Figure 3H and I). Notable is the
overwhelming presence of pennate diatoms, which have previously been identified on
plastic debris recovered from the oceans [4]. Additionally, cells that are likely of bacterial
origin were also observed (Figure 3I-3K).
Microbial communities of Plastic bags #1 and #2 were analyzed using 454
pyrosequencing of a region of 16S rRNA gene. Additionally, the communities of the soil
samples collected in immediate proximity to the bags were analyzed for comparison. It
should be noted that due to limitations of methodology, 18S rRNA from Eukaryotic
organisms was not analyzed. Community analysis indicated that a big fraction of the
communities in both bags was represented by proteobacteria and bacteroidetes (Figure
4A). Search against RDP database indicated a large presence of cyanobacteria in Bag #2
sample (Figure 4A), however further analysis indicated that more than 80% of the
sequences that were assigned as cyanobacteria, were chloroplasts (data not shown). This
classification misnomer is due to the assignment of Eukaryotic chloroplast 16S sequences
as those of cyanobacteria. Among proteobacteria found within Bag #2 sample, 60% was
represented by gammaproteobacteria (Figure 4B). A large fraction of
gammaproteobacterial sequences were classified as Chromatialis, which are
photosynthetic purple sulfur bacteria. These results indicate a high number of
photosynthetic organisms found on the surface of plastic bag #2. Notably, cyanobacteria
(and chloroplasts), gammaproteobacteria in general, and Chromatialis specifically
constituted a much lower fraction of the total microbiome of the control sand sample
collected at the site where bag #2 was found (Figure 4C). This indicates that the plastic
provided a niche that was distinct from that of the sand found in its direct proximity.
These data confirm that a diverse community of microorganisms colonizes plastic found
in the environment [4].
To investigate plastic colonization in vitro, I used SaranTM Wrap. After cutting
plastic wrap into squares I irradiated it with UV light to simulate natural damage induced
by solar radiation. Pieces of the plastic wrap were then placed inside glass tubes (Figure
1C). Following media was then added to the tubes: Condition I. Carbon free medium,
described in the methods, and inoculum from Bag #2. Condition II. Filtered water
collected from Sippewissett salt marsh and inoculum from Bag #2. Condition III.
Unfiltered water collected from Sippewissett. Half of the samples were incubated under
hypoxic conditions (Fig. 1C, left) and another half under aerobic conditions (Figure 1C,
right). Negative controls included “no inoculum” and “no plastic” controls.
After 14 days of incubation, one of the samples from each of the conditions was
used for confocal and scanning electron microscopy. DAPI staining followed by confocal
imaging indicated that plastic was colonized with microorganisms in all tested conditions
(Figure 5). Microbial cells of various shapes were present on plastic, although in contrast
to Bag #2, which was used for inoculation, larger cells were notably absent from in vitro
samples. Scanning electron micrographs of in vitro samples did not confirm the presence
of microbial cells on the surface of plastic. In contrast to the samples collected in the
environment, the surface of the in vitro samples remained smooth after the incubations in
all conditions (data not shown, representative is depicted in Figure 3L).
To measure potential change in the mass of plastics, duplicates of the in vitro
samples were recovered from the tubes after 20 days of incubation. Weight of the plastic
samples after the incubation was compared to their original weight. Mass of the samples
incubated anaerobically in the presence of the inoculum in all three conditions was up to
5% lower than the mass of the same samples prior to incubation (Figure 6A). Plastics that
were incubated in sterile media displayed a lower change in weight (Figure 6A). In
contrast, samples that were incubated under aerobic conditions, displayed an opposite
trend. In this case, the presence of the inoculum appeared to lower the change in weight
of plastic samples (Figure 6B). These results raise the possibility that environmental
microbes alter the rates of plastic deterioration. It should be noted that the experiment
was performed once and the data represent an average and standard deviation of two
samples. Any conclusions about the effect of the microbial communities on integrity of
the plastic samples would be premature.
In addition to plastic microbiome found in the environment, we analyzed
communities colonizing plastic in vitro. After 7 days of incubation at the same conditions
as disrobed for plastic weight change experiments, samples were analyzed with 454
pyrosequencing. In addition to sequencing of plastic-associated microbes, we analyzed
unattached communities that were found in the media.
Rarefaction analysis indicated that the communities inoculated from unfiltered
marsh water growing in the presence of plastic (Condition III) and environmental
samples were most diverse (Figure 7A). Communities that were inoculated from Bag #2
and were incubated with plastic in filtered marsh water were less diverse (Condition II).
Finally, communities that were inoculated from Bag #2 and growing in the presence of
plastic in Carbon free medium were least diverse Condition III). This held true for either
anaerobic or aerobic conditions, and both plastic-attached and unattached communities. It
should be noted that the shapes of rarefaction curves also indicated that many of highly
diverse communities were not sequenced with a sufficient depth.
Strikingly, in carbon-free medium, where the only presumed source of organic
carbon is plastic, proteobacteria was an overwhelmingly dominant phylum (Figure 7B).
Even more surprisingly, in these conditions sequences of Alteromonas genus occupied a
large fraction of all microbial sequences (Figure 7C). Notably, Alteromonas was
occupying a larger fraction of the microbial community found attached to the plastic than
those that was in the medium. This was in stark contrast to the number of Alteromonas
found in Bag #2 communities, which were used as inoculum (Figure 7D). Thus
incubation in the conditions where plastic is the only presumed source of carbon selected
for Alteromonas. I cannot conclude that the presence of plastic contributed to the
selection of Alteromonas in these conditions, as it is possible that Alteromonas were
selected by the conditions presented in carbon-free medium independent of plastic. Due
to limited number of sequencing reactions I did not assess the community of bacteria
present under the same conditions in the absence of plastic.
Despite these limitations, it is tempting to speculate as to the conditions that
allowed for the selection of Alteromonas in the presence of plastic as a sole carbon
source. A number of recent reports have identified Alteromonas as key agents of cyclic
hydrocarbon degraders that are enriched in the sites of anthropogenic oil spills [5-8].
Although the composition of plastic, which was used in the enrichment experiments, is
unknown, cyclic hydrocarbons are frequently used as plasticizers. Plasticizers are
compounds that are added to plastic polymers to provide the material with desired
physical properties (rigidity, plasticity etc.) During plastic deterioration plasticizers
escape into the environment and can potentially serve a source of organic carbon. It is
conceivable that in the conditions where no other source of carbon is available,
Alteromonas, which are able to consume cyclic hydrocarbons leaking from plastic, gain
an advantage over other species. Possible effects of microbes on plastic in vitro may
inform strategies on the development of plastic biodegradation technologies and
detoxification of the environments contaminated with plasticizers.
Methods
Sample collection
The samples where collected during low tide at the Little Sippewissett Marsh salt by
Falmouth, MA, USA. Plastic bag #1 was found on a damp surface, which gets covered
with water during high tide and is exposed to air during low tide. Plastic bag #2 was
found partially covered with sand under water. Along with the plastic bags, the soil that
was covering the bags was collected. Water around plastic bag #2 was collected into a
sterile flask. Samples where processed for DNA extraction and inoculum preparation as
described below within a few hours after collection.
Plastic degradation experiments
Plastic
Commercially available plastic wrap was cut into rectangular pieces and weighed. Using
UV crosslinker (UV Stratalinker 1800, Strategene) plastic wrap was irradiated with
ultraviolet light (254 nm) at 1 J/cm2 on each side.
Glassware
All glassware used in the experiments was soaked in sodium hypchlorite (bleach) for at
least 24 hours to remove organic matter, extensively rinsed (at least 5 times) with double
deionized water, and autoclaved.
Media
Enrichment experiments were carried out ineither filtered (0.2 m) Sippewissett seawater
(see Sample Collection above), unfiltered Sippewissett water or Carbon free medium
(CFM). CFM contained the following: 342.2 mM NaCl, 14.8 mM MgCl2, 1.0 mM CaCl2,
6.71 mM KCl, 10 mM NH4Cl, 1 mM Na2SO4, 1 mM KH2PO4/K2HPO4 and 1X Wolfe’s
trace elements mineral solution. CFM medium was sterilized by autoclave.
Preparation of plastic inoculum
To prepare the inoculum a rectangle about 5 by 10 cm from Bag #2 was cut with sterile
scissors and placed into a 50 ml falcon tube. Thirty ml of sterile sea base (342.2 mM
NaCl, 14.8 mM MgCl2, 1.0 mM CaCl2, 6.71 mM KCl) was added to the tube. The tube
with the bag was vortexed for about 2 hours. The liquid was used for inoculation.
Anaerobic growth conditions:
Pieces of plastic (40-50 mg) were placed into 10 ml of CFM, filtered or unfiltered
Sippewissett water in 30 ml Hungate glass tubes. CFM and filtered swamp seawater was
inoculated with 100 l of the plastic inoculum. Unfiltered seawater was not inoculated.
Nitrogen was bubbled through each sample for 2 minutes and the tubes were immediately
sealed using Hungate technique. Tubes were incubated in the upright position at 30°C.
Negative controls included: uninoculated plastic in CFM, uninoculated plastic in filtered
seawater; inoculated CFM, and filtered/unfiltered seawater without plastic.
Aerobic growth conditions:
Pieces of plastic (20-30 mg) were placed into 5 ml of CFM or filtered and unfiltered
Sippewissett water in 15 ml glass tubes. CFM and filtered Sippewissett seawater was
inoculated with 100 l of the plastic inoculum. Unfiltered seawater was not inoculated.
Tubes were incubated in the shaker at 30°C. Negative controls included: uninoculated
plastic in CFM, uninoculated plastic in filtered seawater; inoculated CFM, and
filtered/unfiltered seawater without plastic.
DNA isolation and processing for community 16S rDNA sequencing
DNA was isolated from the bags collected in the Sippewissett marsh, the soil collected in
the vicinity of the bags and from plastic degradation experiments. To purify DNA from
the bags, 2-5 columns of Biofilm DNA Isolation Kit (MO-BIO) were used per sample as
necessary to acquire amount that was detectable by Nanodrop. To purify DNA from soil
1-5 columns of PowerSoil DNA Isolation Kit (MO-BIO) were used per sample. DNA
from the media was isolated using the Biofilm DNA isolation kit after filtering the media.
16S rDNA was PCR amplified using barcoaded 515F and 907R primers. The presence of
the PCR product was confirmed with gel electrophoresis. PCR samples were normalized
based on DNA concentration, combined and gel purified. Pooled sample was submitted
for 454 pyrosequencing (Life Sciences). Sequences were analyzed with QIIME pipeline
[9]. For quality control, sequences longer than 500 bp or shorter than 300 bp were
removed; any sequences containing ambiguous characters were removed as well.
Chimeric sequences were identified with USEARCH 6.1, using only sequences within
the sequencing files as references, and removed [10]. Clustering was performed with
UCLUST [11]. Operational taxonomic units (0.97) were annotated based on a
representative sequences. Taxonomy was assigned based on RDP database classification
[12]. For rarefaction and alpha diversity measurements interval of resampling was set to
500 with 10 repetitions for resampling.
Epifluorescence and confocal microscopy.
To stain the microbial communities the plastics were cut into ~0.5 cm x 0.5 cm rectangles
with sterile scissors. The plastic rectangles were then placed onto glass microscope slides
and mounted with ~10 ul mounting medium containing DAPI stain. The images were
then analyzed using Zeiss LSM-700 confocal microscopes and Zen (Zeiss) software.
Eukaryotic and Bacterial cells (Figure 2) were stained with catalyzed reporter deposition
fluorescence in situ hybridization probes Euk516 and Eub I-III respectively. Euk516 was
detected with tyramide 595, and Eub I-III with tyramide 488. Confocal microscopy was
carried out at the core facility of the Marine Biological Laboratory, Woods Hole, MA.
Scanning electron microscopy
Plastics were cut into ~0.5 cm x 0.5 cm rectangles with sterile scissors and placed
overnight into 4% PFA. Samples were then dehydrated by consecutive incubations in
70%, 85%, 95% ethanol for 10 minutes, followed by three incubations in 100% ethanol
on ice. Samples were dried using Samdri 780A critical point drier (Tousimis) and coated
with 5 nm platitum using Leica EM MED020 (Leica Microsystems). Samples were
visualized with Zeiss Supra 40VP scanning electron microscope. All procedures were
carried out at the microscopy core facility of the Marine Biological Laboratory, Woods
Hole, MA.
Acknowledgements.
I would like to thank the directors of the 2013 Summer Course in Microbial Diversity
Daniel Buckley and Stephen Zinder (both Cornell) for their dedication and support. This
work was a part of the course. I would also like to thank Ashley Campbell (Cornell) who
was the course coordinator. I am indebted to Sara Kleindienst (University of Georgia,
Athens) who helped with CARD-FISH; Erik Zettler (Sea Education Association), Louis
Kerr (Marine Biological Laboratory) and Kyle Fisk for expertise in microscopy and
Charles Pepe-Ranney for invaluable help with bioinformatics. I would also like to thank
all Microbial Diversity course faculty and assistants for creating an incredible
environment, which allowed for this work to be carried out. Microbial Diversity was
course was funded by Gordon and Betty Moore Foundation, Howard Hughes Medical
Institute, NASA, National Science Foundation and U.S. Department of Energy. I received
individual scholarships from Gary N. Calkins Memorial Scholarship, Thomas B. Grave
and Elizabeth F. Grave Scholarship, and Arthur Klorfein Scholarship and Fellowship
Fund.
Figures
Figure 1. Environmental and in vitro samples. A. and B. Collection site and
appearance of Bag #1 and Bag #2. C. Pieces of SaranTM wrap placed into inoculated
media and incubated in the absence (right) or presence (left) of oxygen.
Figure 2. Confocal microscope analysis of the microbial communities colonizing
plastic bag #2. A. DAPI staining of the surface of Bag #2. B-F. CARD-FISH staining of
the surface. Green fluorescence indicates bacterial cells, red fluorescence is staining for
eukaryotic organisms. G. Co-localization analysis of F. Y-axis represents red
fluorescence of individual pixels, X-axis represents green fluorescence.
Figure 3. Scanning electron micrographs of plastic surfuces. Panels A-G are images
acquired from Bag #1. Panels H-K are images from Bag #2. Panel L is an image of a
sample from in vitro incubations.
Figure 4. Community analyses of microbial consortia colonizing Bag #1 and Bag #2
recovered from Little Sippewissett. A. Phylum level composition of communities
isolated from Bag #1,Bag #2 and adjacent sand samples. B. Classes of proteobacteria,
and orders of gammaproteobacteria identified in Bag #2. C. Classes of proteobacteria,
and orders of gammaproteobacteria identified in Sand sample #2 (collected at the same
site as Bag #2).
Figure 5. DAPI stained confocal images of plastic samples incubated in vitro. A-D
Incubated anaerobically, E-I Incubated aerobically. A, E. Samples incubated in CFS
medium, inoculated from Bag #2. B, F, and G. Samples incubated in filtered
Sippewissett water, inoculated from Bag #2. D, H and I. Samples incubated in unfiltered
Sippewissett water. Red light is due to autofluorescence.
Figure 6. Change in weight of plastic samples incubated in vitro. A. Samples
incubated anaerobically. B. Samples incubated aerobically. Columns represent the
average of two replicates. Error bars represent standard deviation.
Figure 7. Community analysis of in vitro plastic communities. A. Rarefaction curves
for in vitro communities and communities established in vitro. Group 1 contains curves
for environmental communities (Bag #1 and #2, Sand #1 and #2) and communities found
in the samples incubated in unfiltered sea water, including supernatant and attached cells.
Group 2 includes curves for communities from samples incubated in filtered seawater and
inoculated from Bag #2. Group 3 includes curves for communities incubated in CFM and
inoculated from Bag #2. B. Community composition on phylum level of in vitro samples.
Color coding is identical to that of Figure 4. C. Fraction of Alteromonas in CWS samples
as compared to entire community. D. Fraction of Alteromonas in Bag #2 community,
which was used to inoculate in vitro samples.
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