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RESEARCH ARTICLE
Disturbance to conserved bacterial communities in the
cold-water gorgonian coral Eunicella verrucosa
Emma Ransome
1,2,3
, Sonia J. Rowley
2,4,5
, Simon Thomas
1
, Karen Tait
1
& Colin B. Munn
2
1
Plymouth Marine Laboratory, Plymouth, UK;
2
School of Marine Science and Engineering, Plymouth University, Plymouth, UK;
3
Smithsonian
National Museum of Natural History, Washington, DC, USA;
4
Bernice Pauahi Bishop Museum, Honolulu, HA, USA; and
5
University of Hawai’i at
Manoa, Honolulu, HI, USA
Correspondence: Emma Ransome,
Department of Invertebrate Zoology, National
Museum of Natural History, Smithsonian
Institution, Washington, DC 20560, USA.
Tel.: (+1) 202 633 9075;
fax: (+1) 202 357 2343;
e-mail: RansomeE@si.edu
Received 3 March 2014; revised 12 July
2014; accepted 27 July 2014. Final version
published online 01 September 2014.
DOI: 10.1111/1574-6941.12398
Editor: Patricia Sobecky
Keywords
coral microbiology; Eunicella verrucosa; coral
disease; Endozoicomonas spp.; DGGE; clone
libraries.
Abstract
The bacterial communities associated with healthy and diseased colonies of the
cold-water gorgonian coral Eunicella verrucosa at three sites off the south-west
coast of England were compared using denaturing gradient gel electrophoresis
(DGGE) and clone libraries. Significant differences in community structure
between healthy and diseased samples were discovered, as were differences in
the level of disturbance to these communities at each site; this correlated with
depth and sediment load. The majority of cloned sequences from healthy coral
tissue affiliated with the Gammaproteobacteria. The stability of the bacterial
community and dominance of specific genera found across visibly healthy colo-
nies suggest the presence of a specific microbial community. Affiliations
included a high proportion of Endozoicomonas sequences, which were most
similar to sequences found in tropical corals. This genus has been found in a
number of invertebrates and is suggested to have a role in coral health and in
the metabolisation of dimethylsulfoniopropionate (DMSP) produced by zoo-
xanthellae. However, screening of colonies for the presence of zooxanthellae
produced a negative result. Diseased colonies showed a decrease in affiliated
clones and an increase in clones related to potentially harmful/transient micro-
organisms but no increase in a particular pathogen. This study demonstrates
that a better understanding of these bacterial communities, the factors that
affect them and their role in coral health and disease will be of critical impor-
tance in predicting future threats to temperate gorgonian communities.
Introduction
Recent marine disease epizootics have reduced the abun-
dance of a variety of endangered, commercially valuable
and habitat-forming species (Harvell et al., 1999). This
includes a number of corals from tropical and subtropical
systems, where an increasing number of coral diseases are
being described (Bourne et al., 2009). Investigations into
bacterial associations with corals have established that
these communities are often species specific (e.g. Ritchie
& Smith, 1997; Littman et al., 2009), spatially and tempo-
rally stable (Knowlton & Rohwer, 2003; Sharp et al.,
2012) and different from surrounding water and sediment
communities (Frias-Lopez et al., 2002; Carlos et al.,
2013). There is strong evidence that these bacterial com-
munities have a beneficial effect, carrying out functions
such as nitrogen fixation, nitrogen cycling and sulphur
cycling (Lesser et al., 2004; Raina et al., 2009). They also
confer resistance to the host by producing antibacterial
agents (Nissimov et al., 2009; Shnit-Orland & Kushmaro,
2009), which are compromised during disease (Ritchie,
2006). Increased host susceptibility (Lesser et al., 2007)
and increased pathogenicity of coral-associated microor-
ganisms (Rosenberg & Ben-Haim, 2002; Bruno et al.,
2007) have both been proposed to be driving incidences
of disease. In addition, disturbance to the fragile relation-
ships between the coral host and their bacterial commu-
nities has been linked to a variety of environmental
factors, including thermal abnormalities (Harvell et al.,
2002; Rosenberg & Ben-Haim, 2002), increased nutrients
(Bruno et al., 2003) and sedimentation (Voss & Richard-
son, 2006). With an increasingly changing marine
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
MICROBIOLOGY ECOLOGY
environment and an increase in the frequency and inten-
sity of disease (Garrabou et al., 2009), it is essential to
understand the progression of disease and its link to envi-
ronmental stress. Documenting the alteration of coral
bacterial communities is necessary to provide accurate
diagnosis of coral disease for researchers and ecosystem
managers (Ainsworth et al., 2007).
Recently, disease outbreaks have been noted in gorgo-
nian corals (Order: Alcyonacea) in temperate waters of
the NW Mediterranean (Cerrano et al., 2000; Martin
et al., 2002; Garrabou et al., 2009) and in SW England
(Hall-Spencer et al., 2007). Mass mortality by tissue
necrosis has been observed for several species and
although the cause of this tissue loss has not been clearly
defined, opportunistic pathogenic bacteria have been
implicated in a number of gorgonians (Cerrano et al.,
2000; Harvell et al., 2001; Martin et al., 2002). Using cul-
ture-based methods Hall-Spencer et al. (2007) found an
increase in the diversity of culturable bacteria from
healthy to diseased Eunicella verrucosa tissue, many of
which were a close match to Vibrio splendidus. Members
of the family Vibrionaceae are associated with disease in
other coral species (e.g. Godwin et al., 2012), including
infections correlated with temperature stress (e.g. Kush-
maro et al., 1996; Ben-Haim & Rosenberg, 2002). A Vib-
rio strain showing thermo-dependent virulence has also
been isolated from diseased colonies of the gorgonian
coral Paramuricea clavata (Bally & Garrabou, 2007) and
has been shown to be involved in mass mortality events
of this coral in the NW Mediterranean (Vezzulli et al.,
2010). Hall-Spencer et al. (2007) demonstrated that these
Vibrio isolates induced tissue necrosis at 20 °C but not at
15 °C in the laboratory, suggesting a possible link
between E. verrucosa disease and temperature.
Details of the nonperturbed microbial communities
thought to associate with temperate anthozoans are scant
in comparison with tropical anthozoans (Bayer et al.,
2013), as is information concerning how these communi-
ties change with the progression of disease. Given a greater
knowledge of these communities we may further our
understanding of multispecies mutualism, the effect that
environmental conditions have on these associates and aid
our identification of species that play a key role in both
maintaining coral health and progressing disease. Further,
documenting shifts in microbiota, if they occur prior to
signs of visible stress, may also allow the use of microbiol-
ogy as a bio-indicator of both environmental change and
disease (Pantos et al., 2003; Bourne & Munn, 2005).
The aim of this study was to investigate microbial com-
munities associated with the pink sea fan, E. verrucosa,
using molecular based methods. This cold-water gorgo-
nian coral is on the international ‘red list’ of threatened
species and is known to be important for the functional
ecology of the benthic environment in which it is found
(Hall-Spencer et al., 2007). Colonies were monitored for
disease at three sites, chosen for differences in depth and
substratum, off the SW coast of England, during 2008,
and healthy and diseased colonies were sampled in June
and September of 2008. Sedimentation, temperature and
irradiance were also recorded.
Materials and methods
Field observation and sample collection
Two sediment traps were deployed at each site for
12 days in June and September 2008 (English et al.,
1997). Seawater temperature and irradiance were mea-
sured at each site every 15 min throughout the monitor-
ing period (HOBO
data loggers, Onset, MA). On 7
June, E. verrucosa colonies at each site were evaluated for
visual signs of necrotic tissue, epibiont cover and fouling
to determine colony health (see Table 1 for site details)
via a stratified videographic survey using closed circuit
rebreather diving technology (AP Diving Inspiration Clas-
sic). Each individual colony encountered within four per-
manent 10 m 92 m belt transects per site was filmed
face on with a scaled back board aligned appropriately
for scale. Fouling was determined by the percentage cover
per colony quantified from videographic frame grabs in
ImageJ64 (Abr
amoff et al., 2004) and was defined as
established, and therefore nontemporary (e.g. not snagged
on colony temporarily due to water current dynamics)
matter covering the coral axis. Abundance data were
quantified in ImageJ64 from scaled frame grabs (using
belt transects, as above) and counts of colonies ≤2cm
tall were used to determine the previous years recruit-
ment (Munro, 2004). Five healthy colonies and two dis-
eased colonies (depending on presence) of similar size
from each site were tagged. Evaluation of visual condition
of colonies was repeated on 16 September. Recruitment
of E. verrucosa was assessed using abundance data from
2007 and 2008 surveys.
Branches 4 cm in length were collected from each of
37 tagged colonies for analysis. In June, five healthy colo-
nies from each site and one diseased colony from site 1
(16 in total) and in September, five healthy colonies and
two diseased colonies from each site (21 in total) were
collected. Only one colony found at site 1 showed signs
of disease during the June survey, and only two diseased
colonies from each site were found in September.
Branches were placed in plastic bags underwater and
immediately taken to the surface where they were washed
in 0.2 lm filtered, sterile phosphate-buffered saline to
remove loosely attached microorganisms. The branches
were then placed in RNAlater (Life Technologies, Paisley,
FEMS Microbiol Ecol 90 (2014) 404–416 ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
Disturbed bacterial communities in E. verrucosa 405
UK) and kept on ice before transporting back to the lab-
oratory and storing at 80 °C until analysis.
Water samples were not collected for microbial analysis
at coral sites during June and September surveys. There-
fore, twenty litres of water collected from the coastal
monitoring site L4 in June and September of 2008
(8 miles from E. verrucosa sites; 50°15.000N, 4°13.020W),
at 10 m depth, was used to assess the microbial commu-
nity in the water column at the time of coral sampling.
There is no record of E. verrucosa at this site. Water was
prefiltered [140 mm diameter, 1.6 lm GF/A filter (What-
man, Maidstone, UK)] and then applied directly to a
0.22 lm Sterivex filter (Millipore, Billerica, MA). Follow-
ing filtration, each Sterivex was pumped dry, frozen in
liquid nitrogen and stored at 80 °C until analysis.
DNA extraction and purification
DNA was extracted using the DNeasy Blood and Tissue
Extraction Kit (QIAGEN, Manchester, UK) according to
the manufacturer’s protocol for animal tissues, with an
extension of the 56 °C incubation to overnight. To
extract DNA from water column samples, the methodol-
ogy of Neufeld et al. (2007) was used. Following the
extraction, total nucleic acids were eluted in 200 lLof
nuclease-free water and total DNA was quantified by
spectrophotometry (Nanodrop 1000, Thermo Scientific)
and diluted to 20 ng lL
1
.
Denaturing gradient gel electrophoresis
(DGGE)
A nested PCR approach was used to amplify the 16S rRNA
gene for DGGE. DNA was amplified using 0.2 lM of prim-
ers 9bfm (50GAGTTTGATYHTGGCTCAG-30) and
1512uR (50ACGGHTACCTTGTTACGACTT-30)(M
€
uhling
et al., 2008), 0.2 mM of each dNTP, 19PCR buffer (Pro-
mega, Southampton, UK), 2 mM MgCl
2
, 0.25 Units of Go
Taq Flexi (Promega) and 0.4 ng of DNA using the follow-
ing conditions: 1 cycle at 96 °C for 4 min; 35 cycles at
96 °C for 1 min, 53 °C for 1 min and 72 °C for 1 min and
one final extension at 72 °C for 5 min. Each PCR was con-
ducted in triplicate. Following successful amplification,
PCR products were diluted 1 : 10 with dH
2
O and primers
341f (50-CCTACGGGAGGCAGCAG-30) with a 40-bp GC
clamp and 907r (50-CCGTCAATTCMTTTGAGTTT-’3)
were used to amplify a 566-bp section of the 16S rRNA
gene (M€
uhling et al., 2008). The reaction mixture con-
tained 0.5 lM each primer, 0.15 mM each dNTP, 1.5 mM
MgCl
2
and 0.15 Units of Go Taq Flexi (Promega) in a total
volume of 60 lL. The amount of diluted DNA added to
each reaction depended on band brightness from agarose
gels from the first PCR round (from 0.1 to 2 lL of diluted
PCR product was added). Temperature cycling for PCR
amplification was 1 cycle at 94 °C for 5 min; 22 cycles at
94 °C for 1 min, 65 °C for 1 min (decreasing by 0.5 °C
every cycle) and 72 °C for 1 min, 15 cycles at 94 °C for
1 min, 55 °C for 1 min and 72 °C for 1 min and one final
extension at 72 °C for 7 min. PCR products were pooled,
cleaned using the QIAquick PCR purification kit (QIA-
GEN) and total DNA quantified by Nanodrop spectropho-
tometry (NanoDrop Technologies, Delaware). DGGE was
performed using the DCode
TM
System (BioRad) with
800 ng of cleaned PCR products on an 8% polyacrylamide
gel with urea and formamide as denaturants (30–60% gra-
dient), at 60 °C and a constant voltage of 60 V for 18 h.
Subsequently, gels were stained with SYBR Green 1 (Molec-
ular Probes, Eugene, OR) for 30 min and then washed with
deionised water for 30 min. This process was repeated
twice to check for reproducible results.
Clone library construction, sequencing and
phylogenetic analysis
PCR products amplified using primers 9bfm and 1512uR
(as above) were pooled by site, month and health status,
Table 1. Site descriptions
Mean value SD (where appropriate)
Site Site 1 Site 2 Site 3
Month June September June September June September
Latitude 03°58.1160W–04°07.2610W–04°08.8810W–
Longitude 50°17.1020N–50°18.1680N–50°20.0210N–
Depth (m) 24–27 –15–25 –6–9–
Sediment type Medium-fine grain –Coarse grain –Fine grain –
Substratum type Wreck –Rocky reef –Artificial reef –
Average temperature (°C) 14.81 0.44 16.72 0.05 14.97 0.41 16.68 0.10 15.58 0.37 16.83 0.08
Temperature (max–min) (°C) 13.85–15.47 16.62–16.81 14.04–16.81 16.52–17.00 14.61–17.48 16.52–17.57
Average light (lux) 119.4 –547.4 –475.5 –
Sedimentation (cm
2
day
1
) 5.84 2.08 2.31 0.64 4.34 0.57 3.08 0.48 12.36 1.81 16.12 7.1
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
406 E. Ransome et al.
(e.g. five replicates from site 1 healthy colonies, in June),
cleaned, quantified and diluted to 20 ng lL
1
. Twelve
clone libraries were constructed, two for water samples
from June and September, and ten from coral tissue
(healthy pooled samples: June site 1, 2 and 3, September
site 1, 2 and 3; diseased pooled samples: June site 1, Sep-
tember site 1, 2 and 3) using a pGEM
-T easy cloning
kit (Promega) and E.coli JM109 (Promega) competent
cells, using the manufacturer’s protocol. Clones contain-
ing inserts were verified using the vector primers M13F
(50-GTAAAACGACGGCCAG-30) and M13R (50
–CAG
GAAACAGCTATGAC-30). Temperature cycling for PCR
amplification was 1 cycle at 94 °C for 3 min; 30 cycles at
95 °C for 30 s, 59 °C for 30 s and 72 °C for 30 s and
one final extension at 72 °C for 5 min. Forty clones were
randomly selected for each coral library and each water
library and directly sequenced using a BigDye Terminator
v3.1 cycle sequencing kit (ABI) and the M13F primer for
the sequencing reaction. Sequences were analysed on an
ABI3100 automated sequencer. Only one strand of the
DNA fragment was sequenced, proving sufficient for tax-
onomic identification. The 16S rRNA gene sequences
were compared to sequences stored in GenBank (NCBI
database) using the BLAST algorithm to identify bacteria
and archaea associated with E. verrucosa and aligned
using CLUSTALW in MEGA 4 (Molecular Evolutionary Genet-
ics Analysis; Tamura et al., 2007), with the closest rela-
tives from BLAST searches. Sequences were submitted to
GenBank under accession numbers KF180619–KF181098.
Presence of zooxanthellae
PCRs were performed using dinoflagellate-specific primers
symITSFP (50-CTCAGCTCTGGACGTTGYGTTGG-30)
and symITSRP (50-TATCGCRCTTCRCTGCGCCCT-30)
as described by Van Oppen et al. (2001) to amplify zoo-
xanthellae ITS1 region from coral tissue DNA samples.
Statistics
GelCompar (Applied Maths) was used to identify DGGE
bands within the bacterial profiles and construct a binary
matrix based on presence and absence of aligned bands.
Levene’s test for equal variances of environmental variables
between sites was performed in MINITAB 6.0 (Minitab) as
were ANOVA’s and Tukey tests to test for differences
between sites. In Primer-E6 v6 (Clarke & Gorley, 2006), a
Bray-Curtis similarity matrix of DGGE bands (presence/
absence data) and of clone libraries (standardised by total,
square root transformed) with nonmetric multidimen-
sional scaling (MDS) enabled visualisation of similarities
between samples. Hierarchical clustering (group average)
with SIMPROF tests tested for structure in each subset of
data corresponding to each branch of a dendogram (1000
permutations, significance level 5%). To assess variability
as a measure of disturbance to coral bacterial communities
multivariate dispersion (MvDISP) analysis was performed
on DGGE and clone library data. PERMANOVA analysis estab-
lished the importance of site, month and health in deter-
mining bacterial community composition for DGGE data
(Primer-E v6). The Shannon–Weaver (H’) test investigated
diversity in DGGE and clone library analyses, Pielou’s (J’)
test investigated evenness in clone libraries and average
taxonomic distinctness (D+) took phylogenetic relatedness
of different classes into account for clone library data.
Results
Environmental perturbations
The environmental conditions at each site are described
in Table 1. In June, seawater temperature at site 3 was
significantly higher (ANOVA F=12.68, P<0.001) and in
September, seawater temperature at site 1 was signifi-
cantly lower (ANOVA F=7, P<0.01) than other sites.
Light intensity (lux) was significantly lower at site 1
(square root transformed data; ANOVA F=25.76,
P<0.001), and sediment trap analysis showed site 3 to
have a significantly higher sediment load in June and
September (ANOVA F=20.70, P<0.005; F=10.63,
P<0.01), compared with site 1 and 2.
Coral recruitment and fouling
Visual signs of disease in E. verrucosa were evident in Sep-
tember at all three sites; however, the numbers of colonies
affected were much smaller compared with previous years,
where 9 per cent of colonies at 7 of 13 sites were found
with disease (Hall-Spencer et al., 2007). In this study, only
one or two colonies were affected at the sites investigated.
Affected colonies had patches of necrotic tissue that were
soft and white with areas of exposed black gorgonian skele-
ton where the tissue was sloughed. Healthy gorgonians typ-
ically had tough, orange–pink coenenchyme (the
mesogloea surrounding and uniting the polyps in anthozo-
ans). In two-factor ANOVAs with Post hoc tests of the effects
of site and season on E. verrucosa recruitment and degree
of fouling, site 1 had significantly higher recruitment and
significantly lower fouling than site 2 and 3 (F=30.985,
P<0.001; F=23.087, P<0.001).
Microbial community changes: DGGE analysis
of the bacterial 16S rRNA gene
Thirty-seven E. verrucosa branches and two water samples
were collected for analysis of their associated microbiota.
FEMS Microbiol Ecol 90 (2014) 404–416 ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
Disturbed bacterial communities in E. verrucosa 407
While the high stress of the 2-dimensional MDS plot of
the DGGE presence/absence data did not emphasise the
distinction between water and coral samples, SIMPROF
groupings showed these bacterial communities to be dis-
tinct (Fig. 1). In June, DGGE fingerprints from colonies
at site 1 and 2 grouped tightly on the MDS and showed
homogeneity with SIMPROF tests (Fig. 1). In contrast,
the bacterial profiles from colonies at site 3 shared similar
structure to that found in coral samples from site 1 and 2
in September. In September, bacterial profiles of colonies
at site 1 and 2 clustered separately to the same colonies
sampled in June, suggesting a shift in community struc-
ture. Site 3 profiles were mostly homogeneous but clus-
tered separately from other sites in September, with one
colony profile clustering with diseased samples. Interest-
ingly, this colony showed no visual signs of disease. Dis-
eased samples from all three sites, whether in June or
September, also grouped closely on the MDS and clus-
tered in the SIMPROF test, away from healthy colonies,
suggesting the presence of a distinct bacterial community
(Fig. 1). While June samples (Site 1 and 2) and diseased
samples showed tight clustering on the MDS, September
samples were more dispersed (Fig. 1). Multivariate dis-
persion analysis (MvDISP) also highlighted this (June: site
1: 0.354, 2: 0.885, 3: 1.056; September: site 1: 1.095, 2:
1.146, 3: 1.464).
PERMANOVA analysis (factors: site, month and health)
showed health of colonies to be the determining factor in
bacterial community composition (Pseudo-F=2.86,
P<0.01). However, significant interactions were found
between site x health and site x month (Pseudo-F=1.87,
P<0.05; Pseudo-F=2.83, P<0.001), indicating the
complex nature of this data set. Without diseased samples
complicating the analysis, significant differences in com-
munity structure were also found between site (F=2.59,
P<0.001) and month (F=4.84, P<0.001) in a two-
factor ANOVA. In pairwise tests, differences between site 1
and 3 and site 2 and 3 were responsible for overall site
differences seen.
Microbial community composition: clone
library analysis
MvDISP based on genus and class level phylogeny con-
firmed DGGE results: June samples were less dispersed than
September samples, indicating higher disturbance in Sep-
tember. MvDISP also indicated an overall greater distur-
bance to bacterial communities at site 3 (1.52) compared
with site 1 (0.60) and site 2 (1.19). Calculated diversity
indices (Shannon–Weaver index; H’) based on class level
data from NCBI closest relatives for sequence data, high-
lighted an increase in diversity from June to September.
Diversity peaked in diseased samples at all sites and was
more similar to that of water samples (Table 2). At site 3,
diversity was also high in September samples. Pielou’s
evenness values (J’) showed June libraries (and site 1 Sep-
tember) to be dominated by few bacterial classes, whereas
diseased colonies, water samples and September site 3 colo-
nies showed a more even distribution of bacterial classes
within their communities (Table 2), with no detection of a
single dominant taxonomic group when the coral suffers
from disease. Average taxonomic distinctness (D+)of
genus level data confirmed high diversity and unrelatedness
of water samples. D+was also lowest in June samples and
highest in September diseased samples, with diversity val-
ues more similar to that of water samples, and with no
detection of a single dominant ribotype under disease con-
ditions (Table 2).
Water libraries were dominated by the Alphaproteobac-
teria and the Bacteroidetes, but also contained a range of
other bacterial groups. Within these groups, no one spe-
cies on the NCBI database was dominant, with the major-
ity of clones representing distinct NCBI hits. In contrast,
healthy coral tissues were dominated by the Gammaprote-
obacteria, representing 80.0–90.0% of clones in June. In
September, this dominance varied, with 80.0%, 67.5%
and 20% of clones at site 1, 2 and 3, respectively. In Sep-
tember, the Alphaproteobacteria increased at sites 2 and 3
and the Bacteroidetes and the Cyanobacteria at sites 1 and
3, in addition to the Firmicutes at site 2 and the Plancto-
mycetes and the Deltaproteobacteria at site 3 (see Table 3
for closest NCBI database affiliations).
Analysis of clone libraries from visually diseased colo-
nies revealed the dominance of clones affiliated to the
Gammaproteobacteria (site 1, June 57.5% and September
Fig. 1. Multidimensional scaling of microbial community DGGE
bacterial profiles in E. verrucosa colonies. Labels represent June (J)
and September (S) sites (1–3) for healthy samples. Diseased (D)
samples and water (W) samples are also represented. Symbols
represent Simprof test results, showing structure in bacterial
communities between samples and clustering represents similarity
between samples (50%).
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
408 E. Ransome et al.
60%; site 2, 42.5%; site 3, 27.5%), although this repre-
sented a drop in number compared to healthy libraries,
with the exception of site 3 in September. In diseased
samples, the Alphaproteobacteria (site 1 June, 15%; site 3,
30%), the Bacteroidetes (site 1 September, 22.5%; site 2,
15%) and the Verrucomicrobia (site 2, 12.5%) also domi-
nated. Further, clones affiliated most closely with the
Betaproteobacteria, the Deltaproteobacteria, the Planctomy-
cetes, the Cyanobacteria and the Chloroflexi, which
appeared in diseased libraries, were not present in their
healthy counterparts (see Table 3 for closest NCBI data-
base affiliations).
Similarities and patterns between clone libraries can be
visualised in Fig. 2 which shows June and September
libraries to the bottom left of the plots, with an exception
of the September site 3 library, which groups more closely
with diseased and water libraries, demonstrating a shift in
community structure. While there is no pattern to the
distribution of many bacterial classes over the MDS plots
(e.g. the Betaproteobacteria),patterns of change can be
seen in the Gammaproteobacteria, the Alphaproteobacteria
and the Flavobacteria (Fig. 2).
Diseased libraries did not show an increase in specific
genera when the community changed; however, healthy
samples had a large proportion of Gammaproteobacteria
clones that affiliated with one of two sequences in the
NCBI database; Spongiobacter sp.(FJ457274) and Endozoi-
comonas montiporae (FJ347758) (Table 4). None of the
clones from water libraries matched these sequences in
the NCBI database. Interestingly, there were differences
between the number of clones affiliated with these two
sequences found in June, September and diseased
libraries, by site, with total numbers originally lower at
site 3 (Table 4). A number of other clones grouped with
sequences retrieved from previous studies of microbial
diversity associated with corals. These included two
unknown bacteria originally cloned from Eunicella cavo-
lini, which increased from 2.5% to 15.0% at site 1, from
7.5% to 20% at site 2 from June to September and were
also present in site 1 diseased samples (June; 12.5%) and
site 3 June and diseased samples (2.5%). Sequences that
most closely affiliated with Delftia sp., found in three of
the libraries (Site 3 September and diseased; Site 2 dis-
eased) were also found in Acropora millepora from the
Great Barrier Reef (Bourne et al., 2008). Similarly,
sequences matching Achromobacter sp. were found in
E. verrucosa as well as A. millepora (Bourne et al., 2008).
Presence of zooxanthellae
The presence of Spongiobacter prompted us to investigate
whether E. verrucosa tissue contained zooxanthellae as it
has been suggested that Spongiobacter species may break
down the high levels of DMSP produced by endosymbi-
otic dinoflagellates in coral tissue (Raina et al., 2009).
However, no sequences were amplified from the tissue of
E. verrucosa using dinoflagellate-specific primers sym-
ITSFP and symITSRP (Van Oppen et al., 2001) for the
zooxanthellae ITS1 (data not shown).
Discussion
Conserved bacterial communities inhabiting
the gorgonian coral E. verrucosa
This study elucidates, for the first time, that bacterial
communities associate with E. verrucosa. It also confirms
that this shallow, cold-water coral has microbial consortia
that differ from the surrounding environment, like those
seen in warm-water species (e.g. Rohwer et al., 2002; Litt-
man et al., 2009), and endorses recent evidence that tem-
perate octocorals also contain specific bacterial genera
(Bayer et al., 2013; La Riviere et al., 2013). Previous stud-
ies have shown that warm-water coral-associated bacteria
can be species specific (Bourne & Munn, 2005) and also
spatially specific, with similar bacterial communities
inhabiting different species of closely related coral in the
same location, but differing between location (Littman
et al., 2009). However, little is known about microbial
diversity associated with temperate gorgonians (La Riviere
et al., 2013). DGGE analysis in this study provides evi-
dence of conserved bacterial genera associating with
E. verrucosa colonies at the same site and demonstrates con-
servation of these bacteria between sites (1 and 2). While
Table 2. Diversity indices for class level and average taxonomic distinctness for genus level clone library analysis of bacteria associated with
water samples, healthy coral colonies in June (J) and September (S) and diseased (D) colonies at three sites
Water Site 1 Site 2 Site 3
J S J S J (D) S (D) J S S (D) J S S (D)
Number of clones 40 40 40 40 40 40 40 40 40 40 40 40
OTUs 30 33 8 7 15 15 14 10 24 12 17 31
Shannon–Weaver diversity (H’) 2.2 2.5 0.4 0.7 1.3 1.4 0.7 1.1 2.1 0.8 2.3 2.2
Pielou’s evenness (J’) 0.77 0.89 0.31 0.47 0.71 0.64 0.37 0.65 0.79 0.43 0.90 0.79
Average taxonomic distinctness (D+) 77.4 77.2 66.7 73.8 71.6 75.2 71.0 72.2 75.2 70.2 77.7 75.7
FEMS Microbiol Ecol 90 (2014) 404–416 ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
Disturbed bacterial communities in E. verrucosa 409
communities at site 3 do not cluster as closely with site 1
and 2 on the MDS plot, similarities can be seen between
replicate colonies at site 3 (Fig. 1 and Table 2). This sug-
gests that although bacterial populations associating with
E. verrucosa are conserved, environmental factors may
cause bacterial populations to differ to some degree by site.
Although the clone libraries used in this study were
small, phylogenetic analysis confirms the presence of con-
served bacterial groups, displaying distinct differences in
sequence affiliation from water samples (dominated by
the Alphaproteobacteria and the Bacteroidetes), to healthy
E. verrucosa libraries, which were dominated by the Gam-
maproteobacteria. Many studies of warm-water coral tis-
sue have found the Gammaproteobacteria to be abundant
(e.g. Bourne & Munn, 2005; Ainsworth et al., 2006;
Ritchie, 2006; Wegley et al., 2007; Correa et al., 2013), as
have recent investigations of temperate gorgonians in the
NW Mediterranean (La Riviere et al., 2013; Vezzulli et al.,
Table 3. Affiliations in the NCBI database for clones from E. verrucosa tissue, shaded if present in June (J), September (S) and diseased (D)
libraries; shades representing ≤10 clones (lightest grey), 1–20 clones (light grey), 20–30 clones (dark grey) and ≥30 clones (darkest grey)
(u) indicates sequence from uncultured bacterium.
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
410 E. Ransome et al.
2013). In culture-based studies, the dominance of the
Gammaproteobacteria amongst isolates is thought to be
related to their ability to produce antibiotics that limit
the growth of other bacteria, in addition to their resis-
tance to antibiotics (Long & Azam, 2001). In coral and
other marine invertebrate hosts their role is still some-
what unclear.
Of the Gammaproteobacteria found within the clone
libraries, a high proportion of sequences affiliated with the
genus Endozoicomonas (also found under the name Spon-
giobacter; Bayer et al., 2013) in all healthy libraries
(Table 3). Members of the family Hahellaceae, order
Oceanospirillales, this genus has been found to associate
with the ascidians Cystodytes dellechiajei (Mart
ınez-Garcia
et al. 2007) and Ciona intestinalis (Dishaw et al., 2014), the
marine sponge Halichondria okadai (Nishijima et al., 2013)
and the nudibrach Elysia ornata (Kurahashi & Yokota,
2007). It has been found to dominate DGGE bands from
the sea anemone Metridium senile (Schuett et al., 2007)
and clone libraries in pre- and post-disease corals, for
example, in Acropora millipora from the Great Barrier Reef
(Bourne et al., 2008), and three other Acropora species
(Littman et al., 2009). It has also has been isolated from
Montipora aequituberculata (Yang et al., 2010) and Pocillo-
pora damicornis (Kurahashi & Yokota, 2007). More
recently is has been found in the Caribbean gorgonian Gor-
gonia ventalina (Sunagawa et al., 2010) and the Mediterra-
nean gorgonians Eunicella cavolini (Bayer et al., 2013) and
Paramuricea clavata (La Riviere et al., 2013).
The dominance of this group in so many marine inver-
tebrates across diverse habitats suggests that they are
important members of marine invertebrate microbial
communities and that they play a key role in the function
of the coral holobiont, supporting the hypothesis that
these animals may shape their microbial partners for
nutritional or protective benefits. Raina et al. (2009,
2010) suggested that this group may play a role in
degrading DMSP, produced by symbiotic zooxanthellae,
which are known to associate with a number of corals.
This prompted an investigation into the presence of zoo-
xanthellae in E. verrucosa, with no positive result. Correa
et al. (2013) similarly found Endozoicomonas spp. domi-
nating the Caribbean octocoral Pseudopterogorgia elisabet-
hae, which also lack zooxanthellae; adding weight to the
possibility of a different or additional role for this genus
in marine invertebrates. One such function may be pro-
tecting the host from potentially invasive microorganisms
(Klaus et al., 2007) by competing for nutrients and niche
allocation (Koh, 1997; Klaus et al., 2007) and producing
antibiotics (Ritchie, 2006). For example, Ritchie (2006)
demonstrated that bacteria within the mucus of healthy
corals inhibit the growth of other bacteria 10-fold. In this
scenario, this genus may be able to inhabit a diverse
range of marine hosts and maintain their communities by
preventing the proliferation of competing or invading
microorganisms, as suggested by Bourne et al. (2008).
La Riviere et al. (2013) recently suggested that these End-
ozoicomonas associates may also have differentiated to form
a stable symbiotic complex specific to a cnidarian taxon.
This was suggested due to the 16S rRNA gene sequence
similarity of Endozoicomonas associates from the gorgo-
nians P. clavata and Gorgonia ventalina (GU118518; 96%)
when compared to Endozoicomonas sequences from hexa-
coral species (93%), suggesting that these associates (e.g.
GU118518) may be a new Hahellaceae genus, that is,
adapted to gorgonian hosts (La Riviere et al., 2013). While
this may be true, this study shows E. verrucosa sequences to
be more closely related to Endozoicomonas montiporae
(FJ347758; 97% similarity), originally isolated from Monti-
pora aequituberculata, an encrusting pore coral (Yang et al.,
2010). This finding suggests that the similarities between
various Endozoicomonas sp. are not always reflected in sim-
ilarities between their coral hosts and calls for further
research to establish the forces that may drive the composi-
tion of these coral-associated microbial communities.
Spatial differences to bacterial diversity
In this study, three sites that differed in depth, and thus
temperature and light intensity, as well as in substratum
and sediment accumulation, were chosen with the aim of
understanding if bacterial populations associated with
E. verrucosa were stable and conserved. While these vari-
ables are insufficient to capture the extent of variation
between the three sites, they give clues as to some of the
environmental factors at work in moulding these bacterial
communities. While sites 1 and 2 have similar bacterial
communities, site 3 communities are less similar, most
diverse (H’) and most disturbed (Fig. 1 and Table 2).
Site 3 had a significantly higher temperature in June and
had significantly higher sediment loads in both months,
when compared to the other two sites (Table 1). Site 3 was
also the shallowest site (at 6–9 m). A number of factors
have previously been shown to correlate with changing
bacterial diversity associated with corals, including
increased nutrients (Bruno et al., 2003) and temperature.
Both have been shown to disrupt bacterial populations and
increase the severity of disease in a number of corals
(Harvell et al., 2007; Ward et al., 2007). However, an
increase in temperature was only seen at site 3 in June of
this study, and all sites had higher temperatures in Septem-
ber, suggesting that temperature was not a contributing
factor to the increased disturbance seen at site 3. Black
band disease has also been shown to be more prevalent at
shallow sites (Kuta & Richardson, 2002), thus a factor
related to depth, such as nutrient load, may contribute
FEMS Microbiol Ecol 90 (2014) 404–416 ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
Disturbed bacterial communities in E. verrucosa 411
towards disturbance at this site. Finally, Voss & Richardson
(2006) have observed increased sedimentation rates at sites
of disease when compared to healthy sites, indicating that
sediments may also play a role in coral infections. Rogers
(1990) estimated that mean sediment rates for reefs not
subject to stresses from human activities are <1to
Fig. 2. MDS bubble plots based on class level clone library data showing relative abundance of bacterial classes at each site. Diameter of circles
(1, 2, 3, 4) represents the level of the variable for Gammaproteobacteria (40, 28, 16, 4), Betaproteobacteria (3, 2.1, 1.2, 0.3),
Alphaproteobacteria (20, 14, 8, 2) and Flavobacteria (8, 5.6, 3.2, 0.8).
Table 4. Two most common affiliations in the NCBI database for clones from E. verrucosa
Closest Relative Spongiobacter sp. Endozoicomonas sp.
Total %
Accession number FJ572274 FJ347758
Similarity (%) % of clone library Similarity (%) % of clone library
Library
Site 1
June ≥96 30.0 ≥97 50.0 80.0
September ≥96 27.5 ≥97 45.0 72.5
Diseased (J) ≥96 17.5 ≥97 22.5 40.0
Diseased (S) ≥96 22.5 ≥97 30.0 52.5
Site 2
June ≥95 52.5 ≥97 10.0 62.5
September ≥96 22.5 ≥97 15.0 37.5
Diseased (S) ≥95 17.5 ≥97 10.0 27.5
Site 3
June ≥96 15.0 ≥97 22.5 37.5
September ≥96 2.5 ≥97 7.5 10.0
Diseased (S) –0≥97 2.5 2.5
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
412 E. Ransome et al.
c. 10 mg cm
2
day
1
, which is lower than sedimentation
seen at site 3. Sediment particles can smother reef organ-
isms, decrease recruitment (Harvell et al., 2007) and
potentially act as vectors for coral disease (Voss & Richard-
son, 2006). However, to date, there have been no compre-
hensive studies on the effect of increased sedimentation on
the function of coral bacterial communities. Although in
environments where corals are found several factors may
contribute to the differences in microbial communities
observed, these results suggest that high sedimentation
(e.g. from dredging) may affect coral bacterial communi-
ties, which warrants further investigation to benefit the
conservation of E. verrucosa in SW England.
Temporal and health-related changes to
bacterial diversity
This study demonstrated a shift in the bacterial commu-
nity with an increase in bacterial diversity and increased
taxonomic distinctness in September samples, suggesting
temporal stress to these communities (Fig. 1 and
Table 2). A further increase in diversity occurred in visu-
ally diseased colonies, with a more prominent shift in the
bacterial community. Previous studies have demonstrated
shifts and increasing diversity in microbial populations
from healthy to bleached or diseased corals (e.g. Bythell
et al., 2002; Sussman et al., 2008; Vezzulli et al., 2013).
Studies into specific coral diseases have also shown the
appearance of certain bacterial groups with the onset of
disease, for example, in Black Band Disease Delta- and
Epsilonproteobacteria appear and Gamma- and Betaproteo-
bacteria are lost (Cooney et al., 2002). This has been hy-
pothesised to change the physiological function of these
communities, affecting coral health. Striking similarity
(36% of sequences) can also be seen between communi-
ties associated with BBD and a white plague-like disease
affecting Monstastrea annularis (Cooney et al., 2002;
Frias-Lopez et al., 2002; Pantos et al., 2003), suggesting
the development of a specific community around a
unique microenvironment. Bourne et al. (2008) also
showed an increase in the retrieval of Vibrio-related
sequences associated with A. millepora colonies during
bleaching. However, there have been few studies carried
out on shallow, cold-water coral species.
In diseased E. verrucosa, dominance of the Gammaprote-
obacteria persisted; however, Endozoicomonas-affiliated
sequences seen in the healthy libraries were reduced in
visually diseased samples (Table 4), suggesting a break-
down in the natural community, potentially disrupting the
coral’s immune system and enhancing susceptibility to bac-
terial infections and disease (Rosenberg et al., 2008).
Bourne et al. (2008) witnessed a similar event during
bleaching of A. millepora, with a decrease in Spongiobacter-
affiliated clones from 41% in prebleached corals to 3% dur-
ing bleaching. Similar results were reported for the temper-
ate gorgonian P. clavata (Vezzulli et al., 2013). This
suggests that this genus may be vulnerable to environmen-
tal stressors, causing an unbalance within the coral–micro-
bial community. However, in comparison to the culture-
based study by Hall-Spencer et al. (2007), where increases
in the numbers of bacteria observed in diseased tissue of
E. verrucosa were most closely matched to Vibrio splendi-
dus, only one Vibrio-affiliated clone was present in any of
the libraries, found in a diseased colony in June at site 1.
Instead of a Vibrio dominated community, the second most
common bacterial group varied across sites, with the
appearance of clones that affiliated with other bacterial
classes. This contrasts the recent study on P. clavata,in
which a clear increase in Vibrio abundance was docu-
mented and Koch’s postulates were proved for a strain of
Vibrio corallilyticus (Vezzulli et al., 2010, 2013). The lack of
Vibrio sequences found in this study could be due to the
point at which sampling occurred in the progression of dis-
ease. However, it is known that culture-based methods,
such as that used by Hall-Spencer et al. (2007), can isolate
Vibrio spp. from corals when they are not detected within
clone libraries (Gray et al., 2011). Nevertheless, this study
also provides evidence that bacterial communities change
prior to visual signs of stress, indicating that bacteria may
play a more primary role in disease. The comparison
between the Hall-Spencer et al. (2007) study and the
results presented here highlights the need for employing a
range of culture-dependent and culture-independent based
techniques when studying microbial diversity in environ-
mental samples (Donachie et al., 2007; Godwin et al.,
2012) and suggests the need for further monitoring of
E. verrucosa populations to rule out the presence of a par-
ticular pathogen.
In conclusion, despite most E. verrucosa populations
being stable, populations around the Southwest of Eng-
land were affected in 2001 by disease (Hiscock et al.,
2005) and again from 2003 to 2007 (Hall-Spencer et al.,
2007). This gorgonian coral is on the international ‘red
list’ of threatened species and is known to be important
for the functional ecology of the benthic environment in
which it is found. As the dynamics of most coral diseases
are likely controlled by a variety of factors (Bruno et al.,
2003), identifying aspects of environmental change, such
as increased sedimentation, that could influence marine
disease dynamics and devising policies to mitigate against
their impacts are new and important challenges for ecol-
ogists and policy makers alike (Bruno et al., 2003). Fur-
ther, the effect that climate change may have on the
current UK distribution of this species is not yet known
and with this study highlighting similarities to bacterial
communities in tropical and subtropical corals that suffer
FEMS Microbiol Ecol 90 (2014) 404–416 ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
Disturbed bacterial communities in E. verrucosa 413
vast disease outbreaks, a better understanding of the role
of Endozoicomonas species in gorgonians needs to be
achieved.
Acknowledgements
Thank you to Jason Hall-Spencer for help in the collec-
tion of coral samples and G. Rowley, M. Rowley & P.
Rowley of the Maureen of Dart for logistical field sup-
port. This work was funded by studentships from The
Natural Environment Research Council (PML/PhD2008/
03KT; ER), Society of General Microbiology (ST) and the
Leverhulme Trust (SR).
References
Abr
amoff M, Magalhaes PJ & Ram SJ (2004) Image processing
with ImageJ. Biophotonics Int 11:36–42.
Ainsworth TD, Fine M, Blackall LL & Hoegh-Guldberg O
(2006) Fluorescence in situ hybridization and spectral
imaging of coral-associated bacterial communities. Appl
Environ Microbiol 72: 3016–3020.
Ainsworth TD, Kramasky-Winter E, Loya Y, Hoegh-Guldberg
O & Fine M (2007) Coral disease diagnostics: what’s
between a plague and a band? Appl Environ Microbiol 73:
981–992.
Bally M & Garrabou J (2007) Thermodependent bacterial
pathogens and mass mortalities in temperate benthic
communities: a new case for emerging disease linked to
climate change. Glob Change Biol 13: 2078–2088.
Bayer T, Arif C, Ferrier-Pages C, Zoccola D, Aranda M &
Voolstra CR (2013) Bacteria of the genus
Endozoicomonas dominate the microbiome of the
Mediterranean gorgonian coral Eunicella cavolini.Mar Ecol
Prog Ser 479:75–84.
Ben-Haim Y & Rosenberg E (2002) A novel Vibrio sp. pathogen
of coral Pocillopora damicornis.Mar Biol 141:47–55.
Bourne DG & Munn CB (2005) Diversity of bacteria
associated with the coral Pocillopora damicornis from the
Great Barrier Reef. Environ Microbiol 7: 1162–1174.
Bourne D, Iida Y, Uthicke S & Smith-Keune C (2008) Changes
in coral-associated microbial communities during a
bleaching event. ISME J 2: 350–363.
Bourne D, Garren M, Work TM, Rosenberg E, Smith GW &
Harvell D (2009) Microbial disease and the coral holobiont.
Trends Microbiol 17: 554–562.
Bruno JF, Petes LE, Harvell CD & Hettinger A (2003) Nutrient
enrichment can increase the severity of coral diseases. Ecol
Lett 6: 1056–1061.
Bruno JF, Selig ER, Casey KS, Page CA, Willis BL, Harvell CD,
Sweatman H & Melendy AM (2007) Thermal stress and
coral cover as drivers of coral disease outbreaks. PLoS Biol
5: e124.
Bythell JC, Barer MR, Cooney RP, Guest JR, O’Donnell AG,
Pantos O & Le Tissier MDA (2002) Histopathological
methods for the investigation of microbial communities
associated with disease lesions in reef corals. Lett Appl
Microbiol 34:359–364.
Carlos C, Torres TT & Ottoboni LMM (2013) Bacterial
communities and species-specific associations with the
mucus of Brazilian coral species. Sci Rep 3: 162.
Cerrano C, Bavestrello G, Bianchi CN et al. (2000) A
catastrophic mass-mortality episode of gorgonians and other
organisms in the Ligurian Sea (NW Mediterranean), summer
1999. Ecol Lett 3: 284–293.
Clarke KR & Gorley RN (2006) PRIMER v6: User Manual/
Tutorial, 2nd edn. PRIMER-E Ltd, Plymouth, UK.
Cooney R, Pantos O, Le Tissier M, Barer M, O’Donnell A &
Bythell J (2002) Characterization of the bacterial
consortium associated with black band disease in coral
using molecular microbiological techniques. Environ
Microbiol 4: 401–413.
Correa H, Haltli B, Duque C & Kerr R (2013) Bacterial
communities of the gorgonian octocoral Pseudopterogorgia
elisabethae.Microb Ecol 66: 972–985.
Dishaw LJ, Flores-Torres J, Lax S et al. (2014) The gut
geographically disparate Ciona intestinalis harbors a core
microbiota. PLoS ONE 9: e93386.
Donachie SP, Fister JS & Brown MV (2007) Culture clash:
challenging the dogma of microbial diversity. ISME J 1:97–99.
English SA, Wilkinson C & Baker VJ (1997) Survey Manual for
Tropical Marine Resources. Australian Institute of Marine
Science (AIDAB), Townsville, Qld, Australia, pp. 390.
Frias-Lopez J, Zerkle AL, Bonheyo GT & Fouke BW (2002)
Partitioning of bacterial communities between seawater and
healthy, black band diseased, and dead coral surfaces. Appl
Environ Microbiol 68: 2214–2228.
Garrabou J, Coma R, Bensoussan N et al. (2009) Mass
mortality in Northwestern Mediterranean rocky benthic
communities: effects of the 2003 heat wave. Glob Change
Biol 15: 1090–1103.
Godwin S, Bent E, Borneman J & Pereg L (2012) The role of
coral-associated bacteria communities in Australian
subtropical White Syndrome of Turbinaria mesenterina.
PLoS ONE 7: e44243.
Gray MA, Stone RP, McLaughlin MR & Kellogg CA (2011)
Microbial consortia of gorgonian corals from the Aleutian
islands. FEMS Microbiol Ecol 76: 109–120.
Hall-Spencer JM, Pike J & Munn CB (2007) Diseases affect
cold-water corals too: Eunicella verrucosa (Cnidaria:
Gorgonacea)necrosis in SW England. Dis Aquat Organ 76:
87–97.
Harvell CD, Kim K, Burkholder JM et al. (1999) Emerging
marine diseases: climate links and anthropogenic factors.
Science 285: 1505–1510.
Harvell CD, Kim K, Quirolo C, Weir J & Smith G (2001)
Coral bleaching and disease: contibutors to 1998 mass
mortality in Briareum asbestinum (Octocorallia,Gorgonacea).
Hydrobiologia 460:97–104.
Harvell DC, Mitchell CE, Ward JR, Altizer S, Dobson AP,
Otsfeld RS & Samuel MD (2002) Climate warming and
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
414 E. Ransome et al.
disease risks for terrestrial and marine biota. Science 296:
2158–2162.
Harvell CD, Jord
an-Dahlgren E, Merkel S, Rosenberg E,
Raymundo L, Smith G, Weil E & Willis B (2007) Coral
disease, environmental drivers and the balance
between coral and microbial associates. Oceanography 20:
172–195.
Hiscock K, Sewell J & Oakley J (2005) Marine Health Check
2005. A Report to Gauge the Health of the UK ‘s Sea-Life.
WWF, Godalming, UK.
Klaus JS, Janse I, Heikoop JM, Sanford RA & Fouke BW
(2007) Coral microbial communities, zooxanthellae and
mucus along gradients of seawater depth and coastal
pollution. Environ Microbiol 9: 1291–1305.
Knowlton N & Rohwer F (2003) Multispecies microbial
mutualisms on coral reefs: the host as a habitat. Am Nat
162: S51–S62.
Koh EGL (1997) Do scleractinian corals engage in chemical
warfare against microbes? J Chem Ecol 23: 379–398.
Kurahashi M & Yokota A (2007) Endozoicomonas elysicola gen.
nov., sp. nov., a c-proteobacterium isolated from the sea
slug Elysia ornata.Syst Appl Microbiol 30: 202–206.
Kushmaro A, Loya Y, Fine M & Rosenberg E (1996) Bacterial
infection and coral bleaching. Nature 380: p. 396.
Kuta KG & Richardson LL (2002) Ecological aspects of black
band disease of corals: relationships between disease
incidence and environmental factors. Coral Reefs 21: 393–398.
La Riviere M, Roumagac M, Garrabou J & Bally M (2013)
Transient shifts in bacterial communities associated
with the temperate gorgonian Paramuricea clavata in
the Northwestern Mediterranean Sea. PLoS ONE 8:
e57385.
Lesser MP, Mazel CH, Gorbunov MY & Falkowski PG (2004)
Discovery of symbiotic nitrogen-fixing cyanobacteria in
corals. Science 305: 997–1000.
Lesser MP, Bythell JC, Gates RD, Johnstone RW &
Hoegh-Guldberg O (2007) Are infectious diseases really
killing corals? Alternative interpretations of the experimental
and ecological data. J Exp Mar Biol Ecol 346:36–44.
Littman R, Willis B, Pfeffer C & Bourne D (2009) Diversities
of coral-associated bacteria differ with location, but not
species, for three acroporid corals on the Great Barrier Reef.
FEMS Microbiol Ecol 68: 152–163.
Long RF & Azam F (2001) Antagonistic interactions among
marine pelagic bacteria. Appl Environ Microbiol 67:
4975–4983.
Martin Y, Bonnefont JL & Chancerelle L (2002) Gorgonians
mass mortality during the 1999 late summer in French
Mediterranean coastal waters: the bacterial hypothesis.
Water Res 36: 779–782.
Mart
ınez-Garcia M, D
ıaz-Vald
es M, Wanner G, Ramos-Espl
a
A & Ant
on J (2007) Microbial community associated with
the colonial ascidian Cystodytes dellechiajei.Environ
Microbiol 9: 521–534.
M€
uhling M, Woolven-Allen J, Murrell JC & Joint I (2008)
Improved group-specific PCR primers for denaturing
gradient gel electrophoresis analysis of the genetic
diversity of complex microbial communities. ISME J 2:
379–392.
Munro L (2004) Determining the reproductive cycle of
Eunicella verrucosa. Reef Research Report 07/2004 ETR 12.
Neufeld JD, Sch€
afer H, Cox MJ, Boden R, McDonald IR &
Murrell JC (2007) Stable-isotope probing implicates
Methylophaga spp and novel Gammaproteobacteria in
marine methanol and methylamine metabolism. ISME J 1:
480–491.
Nishijima M, Kyoko A, Katsuta A, Shizuri Y & Yamasato K
(2013) Endozoicomonas numazuensis sp. nov., a
gammaproteobacterium isolated from marine sponges, and
emended descirption of the genus Endozoicomonas
Kurahashi and Yokota 2007. Int J Syst Evol Microbiol 63:
709–714.
Nissimov J, Rosenberg E & Munn CB (2009) Antimicrobial
properties of resident coral mucus bacteria of Oculina
patagonica.FEMS Microbiol Lett 292: 210–215.
Pantos O, Cooney RP, Le Tissier MDA, Barer MR, O’Donnell
AG & Bythell JC (2003) The bacterial ecology of a
plague-like disease affecting the Caribbean coral Montastrea
annularis.Environ Microbiol 5: 370–382.
Raina JB, Tapiolas D, Willis BL & Bourne DG (2009)
Coral-associated bacteria and their role in the
biogeochemical cycling of sulfur. Appl Environ Microbiol 75:
3492–3501.
Raina JB, Dinsdale EA, Willis BL & Bourne DG (2010) Do the
organic sulfur compounds DMSP and DMS drive coral
microbial associations? Trends Microbiol 18: 101–108.
Ritchie KB (2006) Regulation of microbial populations by
coral surface mucus and mucus-associated bacteria. Mar
Ecol Prog Ser 322:1–14.
Ritchie KB & Smith GW (1997) Physiological comparison of
bacterial communities from various species of scleractinian
corals. Proceeding from 8th International Coral Reef
Symposium 1: 521–526.
Rogers CS (1990) Responses of coral reefs and reef organisms
to sedimentation. Mar Ecol Prog Ser 62: 185–202.
Rohwer F, Seguritan V, Azam F & Knowlton N (2002)
Diversity and distribution of coral-associated bacteria. Mar
Ecol Prog Ser 243:1–10.
Rosenberg E & Ben-Haim Y (2002) Microbial diseases of
corals and global warming. Environ Microbiol 4: 318–326.
Rosenberg E, Kushmaro A, Kramarsky-Winter E, Banin E &
Yossi L (2008) The role of microorganisms in coral
bleaching. ISME J 3: 139–146.
Schuett C, Doepke K, Grathoff A & Gedde M (2007) Bacterial
aggregates in the tentacles of the sea anemone Metridium
senile.Helgol Mar Res 61: 211–216.
Sharp KH, Distel D & Paul VJ (2012) Diversity and dynamics
of bacterial communities in early life stages of the Caribbean
coral Porites astreoides.ISME J 6: 790–801.
Shnit-Orland M & Kushmaro A (2009) Coral
mucus-associated bacteria: a possible first line of defence.
FEMS Microbiol Ecol 67: 371–380.
FEMS Microbiol Ecol 90 (2014) 404–416 ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
Disturbed bacterial communities in E. verrucosa 415
Sunagawa S, Woodley CM & Medina M (2010) Threatened
corals provided unexplored microbial habitats. PLoS ONE 5:
e9554.
Sussman M, Willis BL, Victor S & Bourne DG (2008) Coral
pathogens identified for White Syndrome (WS) epizootics
in the Indo-Pacific. PLoS ONE 3: e2393.
Tamura K, Dudley J, Nei M & Kumar S (2007) MEGA 4:
Molecular Evolutionary Genetics Analysis (MEGA) software
version 4.0. Mol Biol Evol 24: 1596–1599.
Van Oppen MJH, Palstra FP, Piquet AMT & Miller DJ (2001)
Patterns of coral-dinoflagellate associations in Acropora:
significance of local availability and physiology of
Symbiodinium strains and host-symbiont selectivity. Proc R
Soc Lond B268: 1759–1767.
Vezzulli L, Previati M, Pruzzo C, Marchese A, Bourne DG,
Cerrano C & VibrioSea Consortium (2010) Vibrio
infections triggering mass mortality events in a
warming Mediterranean Sea. Environ Microbiol 12:
2007–2019.
Vezzulli L, Pezzati E, Huete-Stauffer C & Cerrano C (2013)
16SrDNA pyrosequencing of the Mediterranean gorgonian
Paramuricea clavata reveals a link among alterations in
bacterial holobiont members, anthropogenic influence and
disease outbreaks. PLoS ONE 8: e67745.
Voss JD & Richardson LL (2006) Coral diseases near Lee
Stocking Island, Bahamas: patterns and potential drivers.
Dis Aquat Organ 69:33–40.
Ward JR, Kim K & Harvell CD (2007) Temperature affects
coral disease resistance and pathogen growth. Mar Ecol Prog
Ser 329: 115–121.
Wegley L, Edwards R, Rodriguez Brito B, Liu H & Rohwer F
(2007) Metagenomic analysis of the microbial community
associated with the coral Porites astreoides.Environ Microbiol
9: 2707–2719.
Yang CS, Chen MH, Arun AB, Chen CA, Wang JT & Chen
EM (2010) Endozoicomonas montiporae sp. nov., isolated
from the encrusting pore coral Montipora aequituberculata.
Int J Syst Evol Microbiol 60: 1158–1162.
FEMS Microbiol Ecol 90 (2014) 404–416ª2014 Federation of European Microbiological Societies.
Published by John Wiley & Sons Ltd. All rights reserved
416 E. Ransome et al.
