RESEARCH ARTICLE
Bacterial communities associated with the ctenophores
Mnemiopsis leidyi and Beroe ovata
Camille Daniels & Mya Breitbart
College of Marine Science, University of South Florida, St. Petersburg, FL, USA
Correspondence: Mya Breitbart, College of
Marine Science, University of South Florida,
St. Petersburg, FL 33701, USA. Tel.:
727 553 3520; fax: 727 553 1189; e-mail:
[email protected]
Received 25 November 2011; revised 1 May
2012; accepted 6 May 2012.
Final version published online 13 June 2012.
DOI: 10.1111/j.1574-6941.2012.01409.x
Editor: Gary King
MICROBIOLOGY ECOLOGY
Keywords
ctenophore; bacterial community; 16S;
T-RFLP; temporal.
Abstract
Residing in a phylum of their own, ctenophores are gelatinous zooplankton
that drift through the ocean’s water column. Although ctenophores are known
to be parasitized by a variety of eukaryotes, no studies have examined their
bacterial associates. This study describes the bacterial communities associated
with the lobate ctenophore Mnemiopsis leidyi and its natural predator Beroe ovata in Tampa Bay, Florida, USA. Investigations using terminal restriction fragment length polymorphism (T-RFLP) and cloning and sequencing of 16S
rRNA genes demonstrated that ctenophore bacterial communities were distinct
from the surrounding water. In addition, each ctenophore genus contained a
unique microbiota. Ctenophore samples contained fewer bacterial operational
taxonomic units (OTUs) by T-RFLP and lower diversity communities by 16S
rRNA gene sequencing than the water column. Both ctenophore genera contained sequences related to bacteria previously described in marine invertebrates, and sequences similar to a sea anemone pathogen were abundant in
B. ovata. Temporal sampling revealed that the ctenophore-associated bacterial
communities varied over time, with no single OTU detected at all time points.
This is the first report of distinct and dynamic bacterial communities associated with ctenophores, suggesting that these microbial consortia may play
important roles in ctenophore ecology. Future work needs to elucidate the
functional roles and mode of acquisition of these bacteria.
Introduction
Ctenophores are gelatinous zooplankton that are ubiquitous throughout the world’s oceans. Although they are
often mistaken for jellyfish, ctenophores comprise a phylum of their own, and a recent reorganization within the
animal tree of life places ctenophores as the earliestdiverging extant form of multicellular animals (Dunn
et al., 2008). Composed of > 95% water (Bailey et al.,
1995; Scolardi et al., 2006), these carnivorous animals
glide through the oceans via ciliary action and/or currents
(Mills & Haddock, 2007). Their position in the oceanic
food web was initially thought to be an ecological dead
end because of their low biomass content (Vinogradov
et al., 1992; Dumont et al., 2004), but reports have now
shown that ctenophores can be consumed by various fish,
turtles, jellyfish, and even other ctenophores (Arai, 2005).
Recent work has demonstrated that the overall effect of
gelatinous organisms such as ctenophores is to generate
ª 2012 Federation of European Microbiological Societies
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dissolved organic carbon that promotes microbial respiration, further fueling the microbial loop rather than being
incorporated into biomass, which can be transferred up
the food chain (Condon et al., 2011). Because of the
effects of global climate change and the exploitation of
global fish stocks, ctenophore blooms have increased in
number and intensity, and it has been suggested that
these gelatinous animals may replace top predators in the
world’s oceans (Richardson et al., 2009). This scenario is
especially problematic for fisheries, as ctenophores consume pelagic fish eggs and larvae (Purcell & Arai, 2001),
outcompete fish under low oxygen conditions (Decker
et al., 2004; Seibel & Drazen, 2007), and can be introduced by ballast water as invasive species (Ivanov et al.,
2000; Oguz et al., 2008). It has been hypothesized that
ctenophores and jellyfish can serve as vectors for fish
pathogens (Purcell & Arai, 2001), and recent findings
revealed that bacteria associated with jellyfish served as
the source of secondary infections in farmed salmon
FEMS Microbiol Ecol 82 (2012) 90–101
91
Ctenophore-associated bacterial communities
(Ferguson et al., 2010; Delannoy et al., 2011). Such a
compounding set of factors puts ctenophores in a unique
position to severely alter marine food webs (Lynam et al.,
2006), as anthropogenic eutrophication of coastal waters
promotes increasing numbers of dead zones (Diaz &
Rosenberg, 2008) and the number of ctenophore blooms
continues to increase globally (Richardson et al., 2009;
Purcell, 2012).
Studies characterizing the bacteria associated with marine invertebrates have largely focused on benthic organisms such as corals, sponges, and hydrothermal vent
worms. These aforementioned animals all contain microbiota that are distinct from the water they inhabit, with
these bacteria playing critical roles for host ecology. Corals and sponges contain diverse, often species-specific
bacterial communities that fill niches to support functional roles involving carbon, nitrogen, and sulfur cycling,
as well as protection from potential pathogens (Webster
et al., 2001; Hentschel et al., 2002; Rohwer et al., 2002;
Ritchie & Smith, 2004; Ritchie, 2006; Taylor et al., 2007;
Wegley et al., 2007). Conversely, other marine invertebrates harbor specific microbiota that are constrained to
a limited number of bacterial groups. For example,
specific chemosynthetic bacterial symbionts allow the
annelids Riftia pachyptila and Alvinella pompejana to
thrive in the extreme environments near hydrothermal
vents (Cavanaugh et al., 1981; Haddad et al., 1995). In
pelagic invertebrates, the best-studied bacterial symbiosis
is the Hawaiian bobtail squid Euprymna scolopes, whose
light organ is colonized exclusively by Vibrio fischeri (Nyholm & McFall-Ngai, 2004). Whether the microbiota is
extremely specific (such as in the squid-Vibrio system) or
a diverse community similar to that seen in corals and
sponges, it is clear that bacterial associates play crucial
roles in the ecology of marine invertebrates.
Although ctenophores are known to be parasitized by a
variety of eukaryotes including amoebae, dinoflagellates, and
sea anemones (Moss et al., 2001; Hay, 2006; Reitzel et al.,
2007; Smith et al., 2007), no published studies have
described the bacterial communities associated with ctenophores. Given ctenophores’ potential impact on fisheries,
ease of movement across the oceans, and the fact that environmental conditions are changing to favor their persistence, it is vital to determine whether ctenophores contain
specific bacterial communities. This study examined the bacterial communities associated with two ctenophore genera:
Mnemiopsis leidyi, a lobate ctenophore widely distributed
throughout the Gulf of Mexico, Caribbean, and along the
eastern coasts of North and South America (Purcell et al.,
2001), and its natural predator Beroe ovata. The aims of this
study were to determine whether the bacterial communities
associated with ctenophores are distinct from those in the
surrounding water column, compare microbiota between
FEMS Microbiol Ecol 82 (2012) 90–101
the two ctenophore genera, and examine the temporal variability in ctenophore-associated bacterial communities.
Materials and methods
Sample collection
Ctenophore specimens (ranging in size from 2 to 30 mL
total volume) were carefully collected from Tampa Bay
using a dip net off the Bayboro Harbor seawall in St. Petersburg, Florida, USA (27.76°N, 82.63°W). Samples were collected on a single day of the following months: April 2009,
February 2010, March 2010, May 2010, June 2010, August
2010, October 2010, and April 2011. Sampling dates were
chosen based on the visual observation of ctenophores from
the seawall and were sporadic due to the ephemeral nature
of ctenophore blooms. Ctenophore blooms contained
uneven distributions of the two ctenophore genera, and
specimen recovery varied for each time point (numbers of
samples collected from each genus on each sampling date
are shown in Fig. 2). Intact ctenophores were immediately
placed into separate sterile (100-kD-filtered) seawater baths
to remove loosely associated microorganisms. To allow the
ctenophores sufficient time to clear their guts, ctenophores
were incubated in the sterile seawater baths for 4 h, with
complete water changes performed every hour. Specimens
were subsequently given a final rinse with sterile seawater,
flash-frozen with liquid nitrogen, and stored at 80 °C. A
bulk seawater sample (1 L) collected at the same time was
filtered onto a 0.2-lm Sterivex filter (Millipore, Billerica,
MA) to obtain bacteria from the water column for comparison with ctenophore-associated bacteria.
DNA extraction
Individual whole-ctenophore specimens were defrosted
and then pelleted by centrifugation at 2000 g for 30 min at
4 °C. The supernatant was discarded, and 1 mL of the
ctenophore pellet was transferred to a clean microcentrifuge tube. The Sterivex filter containing the water column
bacteria was removed from its housing, cut with a sterile
scalpel from its internal spool, and placed into a separate
microcentrifuge tube with sterile forceps. DNA was
extracted from ctenophore samples and Sterivex filters
using the All Prep DNA/RNA kit (Qiagen, Valencia, CA),
according to the manufacturer’s instructions, with an additional 1-min spin prior to elution into 50 lL of PCR-grade
water. DNA was stored at 20 °C until further processing.
Species confirmation of ctenophore specimens
Visual identification of ctenophores can be difficult,
given the resemblance of many groups and fragility of
ª 2012 Federation of European Microbiological Societies
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92
ctenophore tissues. A molecular phylogenetic approach has
been established for the Phylum Ctenophora (Podar et al.,
2001), and others have since advocated the importance of
combining morphological data with DNA-based identification (Gorokhova et al., 2009; McManus & Katz, 2009).
Universal
eukaryotic
primers
18S-F
(5′-ACCTGGTTGATCCTGCCA-3′) and 18S-R (5′-TGATCC
TTCYGCAGGTTCAC-3′) were used to amplify ctenophore
18S rRNA genes to confirm visual species identifications
(Medlin et al., 1988; Podar et al., 2001). PCRs (50 lL volume) contained final concentrations of 1 lM for each primer, 10 mg mL 1 bovine serum albumin (ThermoFisher,
Waltham, MA), 0.2 mM Apex dNTPs, 19 Apex Taq Buffer, 2.5 U Apex Taq polymerase (Genesee Scientific, San
Diego, CA), and 5 lL of DNA template. PCR conditions
were adjusted from Medlin et al. (1988) with denaturation
at 94 °C for 5 min, 30 cycles of 94 °C for 1 min, 55 °C for
2 min, 72 °C for 3 min, and a final extension at 72 °C for
10 min. The 18S PCR products were cleaned with the
UltraClean PCR Kit (MO-BIO, Carlsbad, CA), sequenced
bi-directionally, trimmed in Sequencher (Gene Codes, Ann
Arbor, MI), and assembled in SeqMan (Lasergene, Madison, WI). A BLASTN (Altschul et al., 1990) query against the
GenBank nonredundant database confirmed the identity of
both M. leidyi and B. ovata (99% identity, Genbank accession numbers: JN653094 and JN653095).
Bacterial clone library construction and
sequencing
16S rRNA gene clone libraries were constructed and
sequenced to analyze the composition of the bacterial communities associated with selected ctenophore specimens and
compare the ctenophore-associated bacteria with those in
the surrounding water column. PCR mixtures (50 lL total
volume) contained 5 lL of template DNA for each ctenophore or water sample and final concentrations of 1 lM of
each primer (27F: 5′-AGAGTTTGATCMTGGCTCAG-3′
and 1492R: 5′-TACGGYTACCTTGTTACGACTT-3′) (Lane,
1991), 1.5 mM MgCl2, 0.2 mM dNTPs, 1 U Apex Red Taq,
and 19 Apex Red Taq Buffer (Genesee Scientific). Touchdown PCR was performed by denaturation at 95 °C for
5 min, followed by 30 cycles of 94 °C for 1 min, 65 °C
( 0.5 °C each cycle) for 1 min, 72 °C for 3 min, and a final
extension at 72 °C for 10 min. Amplicons were confirmed
on a 1% agarose gel containing ethidium bromide. TOPO
TA cloning (Invitrogen, Carlsbad, CA) was used to ligate
PCR products into the PCR4-TOPO vector, which was
transformed into chemically competent Escherichia coli, and
grown on LB-AMP (50 lg mL 1) plates containing
20 mg mL 1 X-Gal. Clones were subsequently screened by
PCR with M13 primers to confirm inserts and then
sequenced by Beckman Genomics (Danvers, MA). Clone
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C. Daniels & M. Breitbart
library dereplication and diversity index values were
obtained using FastGroupII (Yu et al., 2006). Each clone
library was analyzed separately based on 97% similarity with
gaps, which considers single-base insertions or deletions that
might occur during sequencing. Dereplicated sequences
were queried by BLASTN (Altschul et al., 1990), and the
CLASSIFIER module in the Ribosomal Database Project
(RDP) was used to provide robust taxonomic identifications
of the bacterial 16S rRNA gene sequences (Cole et al., 2009).
Sequences were subsequently deposited in NCBI’s GenBank
database (JN653096–JN653274).
Bacterial community profiling (terminal
restriction fragment length polymorphism)
Terminal restriction fragment length polymorphism
(T-RFLP) is a DNA community profiling technique that
has been used to characterize microbial communities from
a wide range of environments (Liu et al., 1997; Schutte
et al., 2008). T-RFLP was selected for this study based on
its reproducibility, ability to discriminate taxa in mixed
communities, and the availability of robust databases for
sequence comparison. The PCR composition was identical
to that used for 16S rRNA gene sequencing, except that the
primers were each labeled with different fluorophores
(27F-TAM and 1492-HEX). PCR conditions consisted of
initial denaturation at 94 °C for 3 min, 30 cycles of 94 °C
for 1 min, 55 °C for 1 min, 72 °C for 3 min, and a final
extension at 72 °C for 10 min (Danovaro et al., 2006).
Products were pooled from triplicate reactions performed
on each sample and cleaned using the DNA Cleaner/Concentrator Kit-5 (Zymo Research, Irvine, CA). Cleaned PCR
products from each ctenophore and water sample were
digested with 10 U of either HhaI, HaeIII, or MspI (New
England BioLabs, Ipswitch, MA) for 3 h at 37 °C in three
separate 20-ll reactions, followed by incubation at 65 °C
for 20 min to stop the digest. Digests were cleaned with the
Zymo DNA Cleaner/Concentrator Kit-5, quantified with
Pico Green (Invitrogen), and normalized to 5 ng lL 1.
Capillary runs on a DNA 3730xl genetic analyzer (Applied
Biosystems, Foster City, CA) at the University of Illinois
Urbana-Champaign Core Facility separated fragments by
size (operational taxonomic units; OTUs) and detected fluorescence values (relative abundance) for each OTU in the
community profiles. Instrument precision was validated by
a series of control runs that included samples ranging from
a monoculture to complex bacterial communities (Daniels
et al., 2011).
T-RFLP data analysis
The Local Southern algorithm (Southern, 1979) in PEAKSv 1.0 software (Applied Biosystems) was used to
CANNER
FEMS Microbiol Ecol 82 (2012) 90–101
Ctenophore-associated bacterial communities
confirm OTU sizes in the raw T-RFLP profiles for each
enzyme and fluorophore. Data were filtered to remove
shoulder peaks and only retain peaks > 5 times the baseline
height (Schutte et al., 2008). An R script binner (Ramette,
2009) was subsequently employed to place the T-RFLP data
into 2-base-pair (bp) bins to account for instrument error
(±1 bp) and normalize the data to reflect the percent contribution of each OTU (peak area) to the total profile fluorescence. Visual confirmation of proper binning was
performed prior to importing the binned T-RFLP profiles
into Microsoft Excel, and OTUs 0.16% of the total fluorescence were retained for downstream analysis (Danovaro
et al., 2006; Luna et al., 2006).
Statistical tests were performed in PRIMER-E v.6.1 (Clarke,
1993) and MATLAB v. 7.8 (MathWorks, Natick, MA) software packages to determine variation among sample types
and across time. Bray–Curtis distances (Bray & Curtis,
1957), which reflect similarity based on OTU composition
and abundance, were calculated from square-root-transformed profile data. Similarity matrices were produced via
pairwise sample comparisons and used to generate multidimensional scaling (MDS) plots. Analysis of similarity
(ANOSIM, ranks of distances) and permutational analysis of
variance (PERMANOVA, means of distances, which is less
prone to type I errors) were performed to determine
whether sample types were significantly different. Nonparametric dispersion analyses were conducted (Anderson,
2006) to confirm that the T-RFLP data did not violate the
PERMANOVA assumption of equal variances (Anderson,
2001). All statistical tests were run with five thousand permutations, and a P-value 0.05 was considered significant. OTUs responsible for driving differences among
samples or across time were identified using the similarity
percentages (SIMPER) module in PRIMER.
Linking clone libraries to T-RFLP profiles
To link T-RFLP OTUs to ctenophore bacterial community
16S rRNA gene sequences, selected clones were sequenced
bidirectionally with the M13 primers to identify dominant
bacterial ribotypes and those that influenced community
variation. After vector trimming, separate in silico digests
of these sequences were performed in Geneious (Biomatters, Auckland, New Zealand) with the same enzymes used
for T-RFLP to match sequences to specific T-RFLP OTUs.
Results
Comparison of bacterial communities from
ctenophores and the water column
In March 2010, specimens from both M. leidyi (n = 7) and
its predator B. ovata (n = 2) were simultaneously collected
FEMS Microbiol Ecol 82 (2012) 90–101
93
along with a sample of the surrounding water for bacterial
community analyses. Significant differences were observed
between T-RFLP profiles from the ctenophores and water
samples for all three enzymes employed (average ANOSIM
R = 0.91, P-value = 0.02; average PERMANOVA F = 4.81,
P-value = 0.01, Fig. 1a). Bray–Curtis values revealed that
the bacterial community profile in the water was on average 71% dissimilar to the M. leidyi bacterial profiles and
86% dissimilar to the B. ovata profiles. The presence of distinct bacterial communities in the ctenophores vs. the
water column was confirmed by 16S rRNA gene sequencing, which revealed very few shared sequences between the
ctenophore and water column samples (Table 1). Figure 1a
further illustrates that the two ctenophore genera contained
different bacterial communities, with M. leidyi and B. ovata specimens clearly separated from each other, regardless of
the restriction enzyme chosen (average ANOSIM R = 0.94,
P-value = 0.07; average PERMANOVA F = 6.3, P-value
= 0.07). The M. leidyi bacterial profiles were on average
82% dissimilar to the B. ovata profiles. In contrast to the
distinct communities observed in the two ctenophore genera, comparisons revealed that bacterial communities from
individual M. leidyi specimens were only 44% dissimilar to
each other, and bacterial communities from B. ovata specimens were 32% dissimilar to each other.
The average number of OTUs detected in the M. leidyi
profiles was 16, while B. ovata profiles contained an average
of 10 OTUs, and the water samples contained 30 OTUs
on average. The lower number of OTUs in the ctenophore
T-RFLP profiles compared with the water column was supported by results obtained from 16S rRNA gene sequencing
of clone libraries. As measured by the Shannon–Weiner
index, bacterial community diversity was lower in M. leidyi
and B. ovata specimens (H′ = 1.9 and H′ = 2.4, respectively)
compared with the water column sample (H′ = 3.8). Rarefaction analysis indicated that further sequencing is necessary
to capture the diversity in all samples, but the water column
curve is much steeper than the ctenophore curves (Fig. 1b).
Proteobacteria, a bacterial supergroup composed of many
taxa with diverse metabolisms, dominated all 16S rRNA
gene libraries regardless of sample type, comprising 50–
83% of the clones (Fig. 1c). Overall, Alphaproteobacteria,
Gammaproteobacteria, and Bacteroidetes were the most
abundant groups detected (Table 1 and Fig. 1c). Further
resolution of the Alphaproteobacteria revealed that the
water column contained the Order Rickettsiales and Rhodobacterales (Supporting Information, Table S1), while the
ctenophore-associated Alphaproteobacteria primarily consisted of the Order Rhodospirillales (Table 1). An uncultured Marinomonas sp. (Gammaproteobacteria belonging to
the Order Oceanospirillales) dominated the M. leidyi library
(52% of sequences), but was not detected in the water column or the B. ovata specimen. When linked to T-RFLP proª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
94
C. Daniels & M. Breitbart
(a)
n=7
M. leidyi
n=2
B. ovata
n=1
Water
0.05
2D Stress:2D Stress:
0.05
(b)
50
HaeIII
M. leidyi
B. ovata
Water
ANOSIM: R = .98, P = .003
2D Stress: 0
2D Stress: 0
Number of Ribotypes
45
40
35
30
25
20
15
10
5
0
0
20
40
60
80
100
Number of Sequences
HhaI
(c)
100
2D Stress: 0
2D Stress: 0
MspI
Clones (% of library)
90
ANOSIM: R = .74, P = .006
Alphaproteobacteria
Betaproteobacteria
Deltaproteobacteria
Gammaproteobacteria
Actinobacteria
Bacillariophtya
Bacteroidetes
Chlorophtya
Cyanobacteria
Firmicutes
Tenericutes
Unclassified
80
70
60
50
40
30
20
10
0
ANOSIM: R = 1, P = .04
M.leidyi B.ovata Water
(n = 92) (n = 91) (n = 61)
files through in silico digests of clones, the Marinomonas
OTU was responsible for 9% of the variation between the
M. leidyi and water column profiles. The B. ovata sample
was dominated by an uncultured Alphaproteobacteria from
the Rhodospirillaceae family, which accounted for 22% of
the clones in the library. Interestingly, a sequence 98% identical to Tenacibaculum aiptasiae a5, which was previously
isolated from diseased sea anemones (Wang et al., 2008),
accounted for the second highest number of clones (21%) in
the B. ovata sample and was exclusive to this ctenophore.
Both ctenophore libraries contained sequences similar to
cultured representatives of the Alphaproteobacteria capable
of hydrocarbon degradation (Thalassospira and/or Alcanivorax). In addition, the M. leidyi library also contained
sequences similar to Nisaea (Table 1), a bacterial genus
known to be involved in nitrogen metabolism (Urios et al.,
2008).
Temporal variation of ctenophore-associated
bacteria
Because of the episodic nature of ctenophore blooms,
March 2010 was the only time point at which M. leidyi
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
Fig. 1. (a) MDS plots comparing T-RFLP
profiles of bacterial communities sampled from
individual Mnemiopsis leidyi (n = 7), Beroe
ovata (n = 2), and a water sample acquired
during a single collection in March 2010.
Three separate enzymes (HaeIII, HhaI, MspI)
and two different fluorophores (only TAM is
shown here) were used to compare bacterial
communities. (b) Rarefaction curve for the 16S
rDNA libraries from the March 2010
ctenophore and water samples. (c) Community
composition of bacterial 16S rDNA libraries
sequenced from March 2010 ctenophore and
water samples. Designations are based on RDP
CLASSIFIER results. n refers to the number of
representative sequences for each library.
and B. ovata were found simultaneously, allowing for
direct comparison between the two genera. However, to
expand our understanding of ctenophore-associated bacterial communities, specimens of M. leidyi were collected
at six time points (April 2009, February 2010, March
2010, June 2010, August 2010, April 2011), and B. ovata
specimens were collected at three time points (March
2010, May 2010, October 2010). Despite high levels of
variability in the M. leidyi bacterial communities over
time, these ctenophores always contained a microbiota
distinct from that of the water column (Fig. 2a). Consistent bacterial T-RFLP patterns were not found across all
M. leidyi temporal samples; however, most T-RFLP
profiles from specimens collected in a specific
month grouped together (average ANOSIM R = 0.69,
P-value < 0.001; average PERMANOVA F = 12.5, P-value
< 0.001, Fig. 2a). Bray–Curtis values demonstrated that
the M. leidyi specimens collected closest to each other in
time (i.e. February/March 2010 and June/August 2010)
had the most similar bacterial communities, while samples collected further apart had more dissimilar bacterial
communities (Fig. 2a). For example, the M. leidyi February 2010 vs. March 2010 bacterial community comparison
FEMS Microbiol Ecol 82 (2012) 90–101
95
Ctenophore-associated bacterial communities
Table 1. Bacterial 16S rDNA sequences from Mnemiopsis leidyi and Beroe ovata specimens collected during a single sampling effort in March
2010
AN
RDP CLASSIFIER (80% threshold)
M. leidyi (n = 92)
JN653096
Gammaproteobacteria, Oceanospirillales
JN653097
Alphaproteobacteria, Rhodospirillales
JN653098
Alphaproteobacteria, Rhodospirillales*
JN653099
Alphaproteobacteria, Rickettsiales
JN653100
Bacteroidetes, Flavobacteria
JN653101
Bacteroidetes, Flavobacteria
JN653102
Actinobacteria
JN653103
Cyanobacteria
JN653104
Tenericutes, Mollicutes
JN653105
Alphaproteobacteria, Rhodobacterales
JN653106
Bacteroidetes, Flavobacteria
JN653107
Bacteroidetes, Flavobacteria
JN653108
Betaproteobacteria, Methylophilales
JN653109
Deltaproteobacteria
JN653110
Gammaproteobacteria
JN653111
Gammaproteobacteria
JN653112
Gammaproteobacteria
JN653113
Bacteria
B. ovata (n = 91)
JN653114
Alphaproteobacteria, Rhodospirillales
JN653115
Bacteroidetes, Flavobacteria
JN653116
Alphaproteobacteria, Rhodospirillales†
JN653117
Alphaproteobacteria, Rhodospirillales*
JN653118
Bacteria
JN653119
Alphaproteobacteria, Rickettsiales
JN653120
Gammaproteobacteria, Chromatiales
JN653121
Gammaproteobacteria, Oceanospirillales
JN653122
Alphaproteobacteria, Rhodobacterales
JN653123
Bacteroidetes, Flavobacteria
JN653124
Actinobacteria
JN653125
Alphaproteobacteria, Rhodospirillales
JN653126
Bacillariophyta
JN653127
Bacteroidetes, Flavobacteria
JN653128
Bacteroidetes, Flavobacteria
JN653129
Firmicutes
JN653130
Gammaproteobacteria, Enterobacteriales
JN653131
Gammaproteobacteria, Pseudomonadales
JN653132
Gammaproteobacteria, Xanthomonadales
JN653133
Tenericutes, Mollicutes
Identity
(%)
Sample source (GenBank top hit)
# of clones
represented
100
99
100
100
100
99
100
99
98
99
100
97
100
100
89
94
99
91
Sapelo Island Microbial Observatory salt marsh
Xiaomaidao Island sewater (China)
Yellow Sea water
Seawater (North Carolina, USA)
Desalination reverse osmosis membrane
Monterey Bay Station MO plankton extracts
Seawater (Chesapeake Bay, USA)
N.E. Pacific Ocean (10 m)
Mnemiopsis leidyi (Baltic Sea)
Gymnodinium catenatum (Dinoflagellate)
NW Mediterranean Sea water
Geoje island seawater (South Korea)
Sapelo Island Microbial Observatory salt marsh
Saanich Inlet, 10 m depth
Zebra mussel Loon Lake (Michigan)
Bantou freshwater reservoir
Sapelo Island Microbial Observatory salt marsh
Hadopelagic sediments (Japan)
48
10
7
5
4
3
2
2
2
1
1
1
1
1
1
1
1
1
99
98
100
100
90
100
97
100
100
100
99
98
99
100
100
91
100
100
100
90
Xiaomaidao Island seawater (China)
Aiptasia pulchella (Sea anemone)
Station Aloha seawater (HOTS, Hawaii)
Nabeta Bay seawater (Japan)
Erythropodium caribaeorum (Octocoral)
Beaufort Inlet seawater (NC, USA)
Hydrothermal vent (SW Indian Ocean Ridge)
Enrichment culture with heavy fuel oil
South China Sea water
Newport Harbour seawater (Rl, USA)
Urban watershed (Brazil)
Emiliania huxleyi culture
Anoxic fjord sediment
Oura Bay seawater (Japan)
Monterey Bay Station MO plankton extracts
Nematostella vectensis (Sea anemone)
Drinking water
Jiaozhou Bay seawater (China)
Catheter biofilm
Penaeus vannamei (Pacific white shrimp)
22
19
12
11
4
3
3
3
2
2
1
1
1
1
1
1
1
1
1
1
Detected
in H20
sample
✓
✓
✓
n represents the total number of 16S rDNA clones in each library. Each representative sequence from FastGroupII dereplication is described by its
GenBank accession number (AN), annotation from the RDP CLASSIFIER, percent nucleotide identity and source of the top Genbank hit, and the
number of clones that the sequence represents. The final column indicates if the sequence was also detected in a clone library from the surrounding water collected simultaneously (Table S1).
*Clones related to Thalassospira spp.
†
Clones related to Nisaea spp.
produced an average dissimilarity value of 59.6% (average
ANOSIM R = 0.28), while the February 2010 samples were
78% dissimilar to those from June 2010 (average ANOSIM
R value = 0.85). Beroe ovata bacterial profiles were also
distinct from the water column at all sampling time
FEMS Microbiol Ecol 82 (2012) 90–101
points. Although B. ovata samples from May and October
2010 collections consistently clustered together and displayed greater overlap (63% average dissimilarity) in bacterial community profiles than the March 2010 vs.
October 2010 B. ovata profile comparisons (88% average
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
96
C. Daniels & M. Breitbart
Apr 2009 Feb 2010 Mar 2010 Jun 2010 Aug 2010 Apr 2011
(a)
n=3
n = 10
n=7
n = 22
n=8
n = 21
(b)
M. leidyi
n=2
Mar 2010
May 2010
Oct 2010
n=2
n=9
n=2
B. ovata
Water
Water
2D Stress: 0.16
2D Stress: 0.07
HaeIII
ANOSIM: R = .75, P < .001
ANOSIM: R = .54, P = .005
2D Stress: 0.2
2D Stress: 0
HhaI
ANOSIM: R = .46, P = .03
ANOSIM: R = .60, P < .001
2D Stress:
0.07
2D Stress:
0.07
2D Stress: 0.19
2D Stress:
0.19
MspI
ANOSIM: R = .74, P < .001
ANOSIM: R = .46, P = .03
M. leidyi
Apr 2009 Feb 2010 Mar 2010 Jun 2010 Aug 2010 Apr 2011
Apr 2009
Feb 2010
Mar 2010
Jun 2010
Aug 2010
Apr 2011
–
0.57
–
0.69
0.28
–
0.88
0.85
0.79
–
0.59
0.71
0.6
0.35
–
0.78
0.73
0.7
0.72
0.73
–
Fig. 2. Temporal T-RFLP MDS plots for Mnemiopsis leidyi (a) and Beroe ovata (b) bacterial communities. ANOSIM values shown on the MDS plots
reflect comparison of the profile data across months. n represents the numbers of ctenophore individuals analyzed at each time point. An ANOSIM
matrix containing average R values for the M. leidyi temporal T-RFLP profiles is shown below the plots. Dark grey shading represents P-values
< 0.01, while light grey shading refers to P-values < 0.05. The ANOSIM matrix for B. ovata is not included since the statistics are not robust due to
the small sample size.
dissimilarity, Fig. 2b), no temporal trends can be inferred,
given the small sample size.
16S rRNA gene sequencing of representative individual
M. leidyi specimens over time verified the temporal variª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
ability observed from the T-RFLP profiles (Fig. 3).
Although Proteobacteria consistently represented > 48%
of the bacterial community, the types of Proteobacteria
varied greatly between months. For example, Gammapro-
FEMS Microbiol Ecol 82 (2012) 90–101
97
Ctenophore-associated bacterial communities
100
90
Alphaproteobacteria
Clones (% of library)
80
Betaproteobacteria
Deltaproteobacteria
70
Epsilonproteobacteria
Gammaproteobacteria
60
Actinobacteria
50
Bacteroidetes
40
Chlorophtya
tinal bacteria from various bivalves constituted 4% of the
clones in the April 2011 library (Table S2). All Betaproteobacteria clones (18%) from the August 2010 library were
similar to Polynucleobacter sp., which have previously only
been reported in freshwater environments (Jezbera et al.,
2011) and were also detected in M. leidyi from June 2010
(Fig. 3, Table S2).
Cyanobacteria
30
Firmicutes
Plantomycetes
20
Tenericutes
Verrucomicrobia
10
Unclassified
0
Apr 2009* Feb 2010~ Mar 2010 Jun 2010 Aug 2010 Apr 2011
(n = 157) (n = 181) (n = 92) (n = 81) (n = 76) (n = 84)
Fig. 3. Community composition of bacterial 16S rRNA gene libraries
sequenced from Mnemiopsis leidyi over time. Annotations are based
on RDP CLASSIFIER results. n refers to the number of clones
sequenced from each library. Sequences originated from a single
ctenophore specimen in each month, except April 2009 (*) which
originated from three separate ctenophores, and February 2010 (~),
which originated from two ctenophores). Clone library composition
for the individual ctenophore samples from April 2009 and February
2010 is shown in Fig. S1.
teobacteria dominated the M. leidyi bacterial community
in March 2010, whereas this group was not detected in
June 2010. In addition, Epsilonproteobacteria made up
18% of the bacterial community associated with M. leidyi
in April 2009, but were not detected at any other time
points. Other bacterial groups also displayed temporal
variation, such as the Bacteroidetes, which comprised 45%
of the sequences in February 2010, but were not recovered in April 2009. Representatives from the Tenericutes
phylum were only present at half of the time points (February 2010, March 2010, August 2010 libraries).
Although the majority of the M. leidyi sequences were
most similar to bacteria previously identified in seawater,
temporal 16S rRNA gene libraries contained many
sequences closely related to bacteria previously described
in various marine invertebrates, including corals (healthy
and diseased), sponges, anemones, and bivalves (Table
S2). Bacteria with order- to species-level identity to
sequences previously described from various corals
(Acropora, Diploria, Montastraea, Porites, and Gorgonia)
were found at multiple time points, peaking at 11% of
the sequences in April 2009. Sponge-associated sequences
represented a small number (< 10%) of the clones at
most time points, except March 2010, and were similar to
bacteria previously described from Callyspongia, Cymbastela, Gelloides, or Tethys spp. (Table S2). In addition, 34%
of the sequences from the August 2010 M. leidyi library
were 99% identical to Gammaproteobacteria previously
identified as gill symbionts of the bivalve Phacoides pectinaus (Table S2). Sequences related to parasites and intesFEMS Microbiol Ecol 82 (2012) 90–101
Discussion
Although microbial communities associated with several
marine invertebrates have been characterized, this is the
first published study to describe bacteria associated with
representatives from Phylum Ctenophora. The high ANOSIM R values based on T-RFLP profiles indicate little to
no overlap (Ramette, 2007) among bacterial communities
from the ctenophores and the surrounding water column,
or between the two ctenophore genera. Given that these
gelatinous animals are composed primarily of water, it is
striking that tissue samples of M. leidyi and B. ovata collected simultaneously contained bacterial communities
different from the water column and distinct from each
other (Fig. 1a). The ANOSIM and PERMANOVA P-values for
both taxa were slightly greater than the significance cutoff; however, this is a function of sample size (only two
B. ovata specimens were obtained during the March 2010
collection). Ctenophore samples contained fewer bacterial
OTUs by T-RFLP and lower diversity communities by
16S rRNA gene sequencing, when compared with the
water column. The low diversity of the ctenophore-associated bacterial communities is different than the trend
previously observed in corals and sponges, where bacterial
diversity is typically higher than that of the surrounding
water column (Rohwer et al., 2002; Taylor et al., 2007;
Webster et al., 2009; Lee et al., 2011).
Ctenophore and water 16S rRNA gene libraries were
dominated by sequences similar to uncultured Alphaproteobacteria, Gammaproteobacteria, and Bacteroidetes
(Fig. 1c). Unlike the water samples, which were primarily
composed of free-living bacteria from the Ricksettsiales
(specifically Pelagibacter) and Rhodobacterales orders
(Table S1), a majority of the Alphaproteobacteria in the
M. leidyi and B. ovata samples from March 2010 were
from the Order Rhodospirillales (Table 1), which are the
purple nonsulfur, photosynthetic bacteria (Kersters et al.,
2006; Gupta & Mok, 2007). These bacteria could potentially be involved in carbon cycling for ctenophores, providing an additional source of fixed carbon to the animal,
especially when prey items are scarce. In addition, the
detection of clones 99% identical to Thalassospira spp. in
both M. leidyi and B. ovata samples from March 2010 is
interesting, as these bacteria are known to be capable of
chemotaxis to inorganic phosphate (Hutz et al., 2011).
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
98
Sequences similar to bacteria described from other
marine invertebrates were present in both M. leidyi and
B. ovata samples. Marinomonas sp., Gammaproteobacteria
from the Order Oceanospirillales, accounted for approximately half of the clones in the M. leidyi 16S rRNA gene
library from March 2010 and has previously been
described in corals and sponges (Cassler et al., 2008;
Thompson et al., 2011). Marinomonas sp. contain genes
for the breakdown of dimethylsulfoniopropionate, suggesting a potential functional role in the biogeochemical
cycling of sulfur (Todd et al., 2007; Johnston et al.,
2008). Mycoplasma and Spiroplasma sp., representatives
from the Phylum Tenericutes appeared at low numbers in
both ctenophore libraries from March 2010 (Table 1),
but were not detected in the surrounding water. These
bacteria are known to parasitize animal and plant tissues
to obtain nutrition and compensate for their reduced
genome size (Rottem, 2003; Pitcher & Nicholas, 2005).
Mycoplasma sp. have previously been reported as associates of other marine invertebrates, such as deep-water
corals (Penn et al., 2006; Kellogg et al., 2009; Neulinger
et al., 2009). Some of the Mycoplasma sequences identified in M. leidyi in this study were 98% identical to
Mycoplasma sp. from M. leidyi specimens collected from
the Baltic Sea (S. Hammann, unpublished GenBank
record), suggesting that there may be a consistent association of this bacterial genus with M. leidyi.
A large component of the B. ovata 16S rRNA gene
library was 99% identical to T. aiptasiae a5, a member of
the Bacteroidetes that was previously isolated from diseased sea anemones (Wang et al., 2008). Ctenophores are
known to harbor eukaryotic parasites and are used as a
refuge by other animals (Gasca et al., 2007); however, this
study also suggests that ctenophores can potentially serve
as vectors for bacteria that are pathogenic to other marine organisms. The genus Tenacibaculum also contains
other pathogenic species that have been detected on jellyfish and implicated in fish gill disease (Ferguson et al.,
2010; Delannoy et al., 2011). Sequences related to the
freshwater-restricted Betaproteobacteria genus, Polynucleobacter sp., were found in the June and August 2010
M. leidyi libraries and may be indicative of a freshwater
input into Bayboro Harbor, where these ctenophores were
collected.
Although the ctenophores in this study always contained bacterial communities that were distinct from the
surrounding water column, the ctenophore-associated
bacterial communities varied significantly over time
(Figs 2 and 3). This type of community flux has been
described by Littman et al. (2009) in juvenile acroporid
corals; however, adult corals and most sponges ultimately
contain conserved bacterial communities (Hentschel
et al., 2006; Taylor et al., 2007; Littman et al., 2009).
ª 2012 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
C. Daniels & M. Breitbart
Unlike these sessile invertebrates, the ctenophores analyzed in this study are pelagic animals, and the changing
bacterial community may represent a lifestyle strategy to
take advantage of the bacteria available in the environment at any given time, rather than spending resources
for maintenance and protection of a conserved consortia.
Although the individual community members (i.e. OTUs
or ribotypes) varied over time, it is possible that the different bacterial communities contained members fulfilling
a given set of metabolic functions for the ctenophore.
However, further work is needed to assess the functional
roles of the ctenophore-associated microbiota. The distribution of M. leidyi populations is linked to temperature
and food availability, and their life span is on the order
of months (Ghabooli et al., 2011; Shiganova & Dumont,
2011); therefore, it is likely that several distinct populations were sampled during this study. If the bacterial
communities are dependent on factors in early life stages
(e.g. the bacterial community present in the water column at the time of initial colonization), the sampling of
different ctenophore populations may explain the temporal variation observed in their bacterial communities.
Alternatively, the bacterial community associated with
individual ctenophores may change over time. If this is
the case, then perhaps the composition of the surrounding bacterial community in the water column or food
availability may influence ctenophore-associated microbiota. Cydippid larvae of M. leidyi primarily feed on protistan prey; however, an ontogenetic shift occurs such that
the diet of lobate adults is dominated by metazoan prey
including copepods, cladocera, and larvae of fish and
mollusks (Rapoza et al., 2005). Although dietary diversity
is similar among M. leidyi > 3 cm in length, the composition of the diet varies based on prey availability. Total
food availability as well as dietary composition may affect
the ctenophores’ bacterial community. Future studies
should consider analyzing zooplankton prey community
composition while examining the ctenophore-associated
bacterial community to determine whether diet is driving
the temporal variability reported here.
Condon et al. (2011) recently demonstrated that the
release of dissolved organic matter by jellyfish could alter
food web dynamics and restructure planktonic microbial
communities, and ctenophores are known to adversely
impact marine food webs (Purcell et al., 2001; Oguz et al.,
2008). The apparent increase in frequency and intensity of
ctenophore blooms, combined with recent invasions of
ctenophores into new areas, enforces the need for a detailed
study of factors influencing ctenophore ecology. This is the
first report of distinct and dynamic bacterial communities
associated with ctenophores, suggesting that these microbial consortia may play important roles in ctenophore biology. While the 16S rRNA gene data in this study provide
FEMS Microbiol Ecol 82 (2012) 90–101
Ctenophore-associated bacterial communities
some clues on the potential metabolic capability of ctenophore-associated bacteria, future work needs to be performed to elucidate the functional roles of microorganisms
associated with ctenophores (e.g. through a metagenomic
survey). Future studies should also examine the location of
the bacteria within the ctenophore body plan, assess the
mode of acquisition of bacteria by the ctenophores, and
attempt to determine the factors driving the temporal variability in ctenophore-associated bacterial communities
observed in this study.
Acknowledgements
This work was funded by a grant to M.B. from the Alfred
P. Sloan Foundation (BR-4772). C.D. was funded by the
McKnight Doctoral Fellowship from the Florida Education Fund. Thanks to Mary Beth Decker, Paul Turner,
and David Jones for helpful discussions and the Breitbart
Lab, especially to Karyna Rosario Cora and Darren
Dunlap, for their assistance during sample collection.
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Supporting Information
Additional Supporting Information may be found in the
online version of this article:
Fig. S1. Community composition of bacterial 16S rDNA
libraries sequenced from M. leidyi individuals from the
same collections in April 2009 and February 2010.
Table S1. Bacterial 16S rDNA sequences from the water
sample collected during the March 2010 sampling effort.
Table S2. Bacterial 16S rDNA sequences from the April
2009, February 2010, June 2010, August 2010, and April
2011 M. leidyi samples.
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