Coral disease is one of the major causes of reef degradation. Dark Spot Syndrome (DSS) was described in the early 1990's as brown or purple amorphous areas of tissue on a coral and has since become one of the most prevalent diseases reported on Caribbean reefs. It has been identified in a number of coral species, but there is debate as to whether it is in fact the same disease in different corals. Further, it is questioned whether these macroscopic signs are in fact diagnostic of an infectious disease at all. The most commonly affected species in the Caribbean is the massive starlet coral Siderastrea siderea. We sampled this species in two locations, Dry Tortugas National Park and Virgin Islands National Park. Tissue biopsies were collected from both healthy colonies and those with dark spot lesions. Microbial-community DNA was extracted from coral samples (mucus, tissue, and skeleton), amplified using bacterial-specific primers, and applied to PhyloChip G3 microarrays to examine the bacterial diversity associated with this coral. Samples were also screened for the presence of a fungal ribotype that has recently been implicated as a causative agent of DSS in another coral species, but the amplifications were unsuccessful. S. siderea samples did not cluster consistently based on health state (i.e., normal versus dark spot). Various bacteria, including Cyanobacteria and Vibrios, were observed to have increased relative abundance in the discolored tissue, but the patterns were not consistent across all DSS samples. Overall, our findings do not support the hypothesis that DSS in S. siderea is linked to a bacterial pathogen or pathogens. This dataset provides the most comprehensive overview to date of the bacterial community associated with the scleractinian coral S. siderea.
Citation: Kellogg CA, Piceno YM, Tom LM, DeSantis TZ, Gray MA, Andersen GL (2014) Comparing Bacterial Community Composition of Healthy and Dark Spot-Affected Siderastrea siderea in Florida and the Caribbean. PLoS ONE 9(10): e108767. https://doi.org/10.1371/journal.pone.0108767
Editor: Melanie R. Mormile, Missouri University of Science and Technology, United States of America
Received: June 6, 2014; Accepted: September 3, 2014; Published: October 7, 2014
This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All microarray and metadata files are available from Greengenes, accessible by the following dedicated URL: (http://greengenes.lbl.gov/Download/Microarray_Data/Siderastrea_Kellogg.zip). The data also have been deposited in the NCBI's Gene Expression Omnibus  and are accessible through GEO Series accession number GSE60622 (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE60622). All other relevant data are within the paper and its Supporting Information files.
Funding: This project was supported by the Coral Reef Ecosystems Study (CREST) of the United States Geological Survey's Coastal and Marine Geology Program (http://marine.usgs.gov). A portion of this work was performed under the auspices of the United States Department of Energy under contract DE-AC02-05CH11231 to Lawrence Berkeley National Laboratory. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: Author Christina A. Kellogg is a PLOS ONE Editorial Board member, and this does not alter the authors' adherence to all the PLOS ONE policies on sharing data and materials. The authors also agree to declare co-author Todd DeSantis's affiliation with the company Second Genome, Inc. and confirm that this also does not alter their adherence to all the PLOS ONE policies on sharing data and materials.
Diseases of reef-building corals are now considered a major cause of global coral reef ecosystem decline , . The past two decades have seen a dramatic increase in the number of reports of coral diseases, particularly in the Caribbean –. Most of these diseases are known or suspected to be microbial in origin . Moreover, microbiology is a key part of coral biology, in the same way that human microbiome studies are revealing microbes to be a critical part of human biology .
Siderastera siderea, also known as the massive starlet coral, is a common component of Caribbean reefs, occurring from the Gulf of Mexico to South America . However, there has been little attention focused on the bacterial communities associated with this coral, other than one culture-based study , two clone library studies focused specifically on black band lesions ,  and a recent pyrosequencing study of a white plague-like disease .
Dark spot syndrome (DSS) was first reported as a discoloration observed on S. siderea, Stephanocoenia intercepta, Porites astreoides, and Montastraea cavernosa near Columbia in the early 1990's . Dark spot lesions are described as purple, black, or brown discolored areas of tissue that may be circular, elongate, ring-shaped, or occur lining the coral tissue-algal boundary of an older lesion – (Fig. S1). There is some argument in the literature as to whether to define dark spot as a disease (DSD) or a syndrome (DSS) . This tissue discoloration has been linked to both physical  and microbiological ,  causes, leading some to suggest it may be a non-specific stress response , . Unlike coral diseases such as black band or white plague, dark spot lesions rarely cause whole colony mortality and result in relatively low net tissue loss . Further, dark spot lesions have been observed to disappear in as little as a month , . It has been suggested that dark spots may signal different conditions in different coral species , –. Given that breadth of scope, we have opted to use the more comprehensive term ‘dark spot syndrome’ (DSS), but acknowledge that it can be regarded as synonymous with DSD given the recent push to apply a broader medical definition (“any inhibition of normal function”) to coral diseases , .
Although DSS has been identified in several species of Caribbean corals, S. siderea is the most frequently affected , , –. Previous coral disease work has shown that bacterial communities shift when their hosts are stressed (even in normally pigmented tissues ). Therefore, we hypothesized that there would be a shift in the bacterial communities of DSS-affected tissues compared to healthy colonies, regardless of whether DSS was due to a general immune response or an infection. Because there appear to be multiple possible etiologies for DSS lesions, we wondered if there would be geographic differences or perhaps multiple clusters of DSS bacterial communities rather than the single diagnostic grouping we observed previously in a white plague-like disease . To address these questions, we collected S. siderea samples from Florida and the Virgin Islands and used PhyloChip G3 DNA microarrays to examine the breadth of taxonomic diversity of the bacterial communities associated with both healthy and DSS-affected colonies. This is the first study to apply molecular techniques to the study of DSS in S. siderea.
Materials and Methods
These collections were made under permits VIIS-2008-SCI-0033 (study VIIS-08033) and DRTO-2009-SCI-0018 (study DRTO-00074), granted to the first author by the Virgin Islands National Park and Dry Tortugas National Park, respectively. No ethical approval was required for the experimental research described here.
Sample sites and collections
Healthy (i.e., exhibiting normal pigmentation) and DSS-affected S. siderea samples were collected from Dry Tortugas (DRTO) and Virgin Islands (VIIS) National Parks during the summer of 2009 (Fig. 1). The collection areas at each park were localized such that most coral colonies sampled were within 10 meters of another, and the maximum distance between two coral colonies sampled at one site did not exceed 140 meters. Five healthy and five DSS-affected colonies were sampled from the three-lobed patch reef in Hawksnest Bay (18°20′50 N, 64°46′50 W), St. John, VIIS on June 23, 2009 (Table 1). The five healthy colonies were all located to the north (offshore) of the eastern-most lobe. The dark spot colonies were found on the eastern side of each of the three reef lobes. The water temperature was 29°C with a salinity of 34 ppt. Samples from five healthy and five DSS-affected colonies also were collected from an area on the west side of Loggerhead Key, DRTO (24°38′05 N, 82°55′22 W) on August 3–5, 2009 (Table 1). The water temperature was 31–32°C with a salinity of 35 ppt. Three of the samples, one healthy and two dark spot (DRTOSSD02, DRTOSSH05, and DRTOSSD09) were collected and their bacterial community DNA extracted, but they failed to amplify, so they were not included in Table 1 and do not appear in any subsequent analyses.
Corals were sampled using a bleach-sterilized metal punch of approximately 2-cm diameter to collect two biopsies of tissue with minimal skeletal material attached. DSS-affected corals were sampled with the punch centered over discolored tissue (i.e., in the dark spot lesion). Note that we rarely encountered colonies with discolored tissue spots completely surrounded by normally pigmented tissue (samples DRTOSSD07, DRTOSSD10, VIISSSD07); instead most of our discolored tissue samples were found along the edge of larger, older lesions (Fig. S1). Coral tissue samples were immediately placed into sterile Whirl-pak bags. The punch was cleaned and re-bleached between samples to prevent any transfer of microbes or contamination. Back on shore, the coral samples were briefly rinsed with sterile-filtered seawater to remove loosely associated microbes, wrapped in sterile aluminum foil, placed into a fresh Whirl-pak bag, and flash-frozen in liquid nitrogen. Samples were transported from the field to the USGS St. Petersburg Coastal and Marine Science Center and then transferred to a −80°C freezer for storage.
In the laboratory, each pair of frozen tissue samples from a single coral colony was ground to powder using a sterile mortar and pestle on dry ice and then combined. The microbial community DNA was extracted in duplicate from each combined sample using the MO BIO PowerPlant DNA Isolation Kit with some modification as previously described by Sunagawa et al. . At the end of the protocol, the replicate extractions were combined, resulting in a single microbial DNA extraction representing each coral colony sampled.
PhyloChip G3 microarray
Microbial profiles were generated for the 17 coral samples using PhyloChip G3 microarrays as described previously , , employing the exact PCR conditions and 16S bacterial ribosomal RNA gene primers previously detailed by Kellogg et al. . In some cases the primers amplified both coral and bacterial DNA , requiring gel extraction of the bacterial amplicons for all samples to be consistent. The difference in size between the two products (i.e., coral and bacterial amplicons) was roughly 200 base pairs (bp). To allow clear separation of the target (bacterial) amplicon, gels were run twice as long as usual (25 min) before being stopped to excise the 16S rRNA amplicon.
For each sample, 500 ng of PCR product was applied to an individual PhyloChip G3 DNA microarray. Fragmentation of the 16S rRNA gene amplicons, labeling, hybridization, staining, probe selection, probe scoring of the microarrays and initial data acquisition were conducted according to Hazen et al. .
The taxonomy used for this dataset  was the same as that used for our recent study of a white plague-like disease  (i.e., updated since ). The array fluorescence intensity data were scaled to internal standards and then operational taxonomic units (OTUs) were normalized by dividing the OTU intensity value by the individual array sum and then multiplying by the average of all total array sums. Stage 1 of PhyCA analysis removed OTUs not present in any of the samples using cutoff values: q1 = 0.5, q2 = 0.93, q3 = 0.98. Post-Stage 2 (PhyCA analysis) data were obtained using cutoff values: q1 = 0, q2 = 0, q3 = 0.1. This reduced the total number of OTUs subject to analysis from roughly 60,000 (all possible OTUs) down to 9,700. OTUs only present in a single sample were removed, further decreasing the total number of OTUs to 4,978. The microarray data have been archived online at Greengenes, accessible by the following dedicated URL: (http://greengenes.lbl.gov/Download/Microarray_Data/Siderastrea_Kellogg.zip). The data also have been deposited in the NCBI's Gene Expression Omnibus  and are accessible through GEO Series accession number GSE60622 (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE60622).
A QIIME-formatted OTU table based on the normalized intensity data and a mapping file were created and run through QIIME . The script ‘summarize_taxa_through_plots.py’ was used to generate taxa summary plots of OTU relative intensity (as a proxy for relative abundance) at the phylum through family levels.
PRIMER 6 version 6.1.13 software  was used to conduct non-metric multidimensional scaling (NMDS) analyses. The normalized (as described above), square-root transformed intensity data also were used to calculate the two-way crossed analyses of similarity (ANOSIM) to test for significant differences in the bacterial community composition between predefined sample sets (i.e., based on collection location and health state).
Fungal extraction and amplification
Based on recent findings by Sweet et al. , we also wanted to check our samples for fungal diversity. The DNA from approximately 50 mg of each powdered coral sample was extracted using the Qiagen Blood and Tissue extraction kit, following the manufacturer's Gram- positive protocol. Extractions were amplified using the nested PCR protocol described by Sweet et al.  with the modification that the second set of primers did not include the GC clamp that is specific to denaturing gradient gel electrophoresis (DGGE). The nested amplifications were also attempted using a different version of the ITS1 primer . PCR amplicons were visualized on a 1% agarose gel.
The PhyloChip G3 microarrays detected 9,700 operational taxonomic units (OTUs) across the 17 samples of S. siderea. After removal of OTUs that only occurred in a single sample, there were 4,978 OTUs that represented 70 different phyla. The major phyla detected were Proteobacteria, Firmicutes, Actinobacteria, Bacterioidetes, Cyanobacteria, Planctomycetes, Acidobacteria, Chloroflexi, Spirochaetes, Tenericutes, Verrucomicrobia and Gemmatimonadetes (Fig. 2), partly reflecting the prominence of probes on the array for these major lineages. Regardless of health state or location (Table 1), all coral samples had similar looking bacterial communities at the phylum level (Fig. 2). When examining the samples at the family level (Fig. 3), the same similarity across location and health state was observed. The most variable family across all samples was the Vibrionaceae, which appeared slightly higher in some DSS samples from the Virgin Islands (VIISSSD06, VIISSSD08, VIISSSD09, and VIISSSD10), but also was enriched in a healthy sample (VIISSH05) (Fig. 3, Fig. S2). Less variation was observed across the other major families represented on the PhyloChip G3 and detected in these samples: Aquabacteriaceae, Bacillaceae, Comamonadaceae, Corynebacteriaceae, Enterobacteriaceae, Flavobacteriaceae, Lachnospiraceae, Micrococcaceae, Phyllobacteriaceae, Planctomycetaceae, Pseudoalteromonadaceae, Pseudomonadaceae, Rhizobiaceae, Rhodobacteraceae, Rhodospirillaceae, Rikenellaceae II, Ruminococcaceae, Staphylococcaceae, Streptococcaceae, and Ulvophyceae (Fig. 3). Over 200 additional families represented by fewer OTUs were also detected, resulting in the identification of over 600 bacterial genera (Table S1).
Minor phyla (less than 0.5% of any sample) and unclassified sequences are collectively represented by the ‘other’ category.
The 21 families shown were those representing greater than 1% of at least one sample. The remaining families and unclassified sequences are collectively represented by the ‘other’ category.
NMDS analysis of the normalized intensity data from the 4,978 shared OTUs showed that the samples do not cluster based on geographic location (Fig. 4). Although four DSS samples did group together (Fig. 4), it was determined using a PERL script previously developed to sort microarray OTUs  that there were only 13 OTUs unique to that group, and none were shared by all four samples. Further, ANOSIM conducted on the 4,978 shared OTUs did not find any significant differences between either the locations (ANOSIM Global R = −0.046, p = 0.63) or health states (healthy versus DSS; ANOSIM Global R = 0.162, p = 0.067).
Samples with the prefix ‘DRTO’ were from Dry Tortugas National Park. Samples with prefix ‘VIIS’ were from Virgin Islands National Park.
While there was not a single OTU or a defined group of OTUs with higher relative abundance(s) in all DSS samples compared to healthy coral samples, there were a few OTUs with notable increased relative abundance in several of the DSS samples (Figs. S2, S3, S4). For example, OTU 53657 (containing one sequence aligned with Pseudoscillatoria coralii str. BgP10_4S, GenBank accession FJ210722.2) had intensity values>10,000 units in four DSS samples (DRTOSSD08, VIISSSD06, VIISSSD09, VIISSSD10), whereas the intensity values for all healthy tissue samples and several other DSS samples were <1,000 units. Because P. coralii has been found associated with black band disease (BBD) – and BBD microbial consortia often include sulfate-reducing bacteria , , we also examined relative intensities of Deltaproteobacterial OTUs and appropriate OTUs in classes Clostridia and Nitrospira. No sulfate-reducing OTUs with notably higher relative intensities in DSS samples were found.
All attempts to amplify and clone fungal sequences from S. siderea samples following the method used by Sweet et al.  failed. The fungal primers used (ITS1F, ITS3F, ITS4F ) were not specific enough to differentiate between fungi and coral in S. siderea (similar to co-amplification of coral 18S rRNA genes by bacterial primers ). PCR resulted in strong amplification of a 380 bp amplicon and extremely faint to no amplification of the expected 500 bp fungal amplicon. Subsequent cloning and sequencing of the 380 bp band revealed it to be coral DNA. Attempts were made to optimize the method using gradient PCR, gel extraction and re-amplification, and employing a different fungal primer (ITS1: TCCGTAGGTGAACCTGCGG ). In all cases, the 380 bp coral amplicon dominated, and when present, the possibly fungal 500 bp amplicon was at such a low concentration that it was not possible to clone.
Opinions on the importance of DSS to reef ecology range from those considering it of limited significance due to the low mortality rates and limited tissue loss  to those who feel that the high frequency of occurrence and links to greater susceptibility to subsequent bleaching or disease make it a useful indicator of reef health , . Our initial hypothesis was that we would detect differences in the bacterial communities between healthy and DSS-affected tissues as we had between healthy and white plague-like diseased corals , possibly with a geographic component. While half of the DSS samples formed a cluster at 88% similarity (Fig. 4), this was not statistically significant (healthy versus DSS; ANOSIM Global R = 0.162, p = 0.067), and the remaining DSS samples grouped with healthy corals or alone. We did not detect a difference in overall bacterial diversity nor relative abundance of specific taxa (as in ) between all normally pigmented S. siderea colonies and all those with DSS. Further, there were not substantial geographic differences between the Florida and Virgin Island samples. Most previous microbiological work comparing healthy and DSS-affected S. siderea has been conducted on corals from the southern Caribbean (i.e., Columbia, Venezuela) , , providing an opportunity for biogeographic comparisons to our data from Florida and the Virgin Islands.
Bacterial diversity of healthy Siderastrea siderea
This dataset provides the most comprehensive overview to date of the bacterial community associated with the healthy scleractinian coral S. siderea. Previous work has consisted of a culture-based study of S. siderea mucus  and a recent study that employed pyrosequencing . Gil-Agudelo et al.  cultured bacteria from S. siderea mucus from both healthy and DSS-affected corals. The coral-associated bacterial isolates were identified by metabolic tests (BIOLOG) rather than 16S rRNA sequencing and consisted entirely of Gram-negative Gammaproteobacteria, which probably reflects selection bias of the GASWA culture medium employed . Most metabolic groups were found in both healthy and DSS corals and corresponded to genera identified in this study (e.g., Vibrio, Klebsiella, Aeromonas; Table S1), most notably those in families Enterobacteriaceae and Vibrionaceae (Fig. 3).
Cárdenas et al.  used pyrosequencing of the 16S rRNA gene to examine healthy and white plague-affected corals. Their dataset includes 378 OTUs from healthy S. siderea. These were dominated by Proteobacteria (75% of sequences), followed by less than 10% each of Firmicutes, Actinobacteria, and Bacteroidetes (in decreasing order of relative abundance) . This perfectly matches the order of relative abundance of our four top phyla (Fig. 2), suggesting the microarray data adequately reflects relative distributions of prominent community members. The remaining phyla detected by pyrosequencing  included low percentages of Chloroflexi, Fusobacteria, Verrucomicrobia, Chlorobi, Tenericutes and Cyanobacteria, all of which we also detected (Fig. 2; note that Fusobacteria and Chlorobi were compiled into the ‘other’category). Phyla detected by the PhyloChip G3 in our S. siderea samples (Fig. 2) that were not found by Cárdenas et al.  include Acidobacteria, Gemmatimonadetes, Planctomycetes and Spirochaetes. This may be due to biogeographical differences between the northern and southern Caribbean, but more likely is due to the greater depth of 16S rRNA sequences queried by the microarray .
Two recent clone library studies allowed us to compare S. siderea with its sister species from Brazil, S. stellata , . From these papers we identified 50 sequences, representing 119 clones, from healthy S. stellata. At the phylum level, the two species looked similar, with healthy S. stellata dominated by Proteobacteria (>80% of sequences), followed by much lower percentages of Bacteroidetes, Cyanobacteria, Actinobacteria, and Verrucomicrobia , . However, the two species were markedly different when examined at the family level. Families in common between the two species were Comamonadaceae, Corynebacteriaceae, Flavobacteriaceae, Rhodobacteraceae, and Rhodospirillaceae, as well as Bradyrhizobiaceae, Brucellaceae, Burkholderiaceae, Moraxellaceae, Propionibacteriaceae, Puniceicoccaceae, and Sphingomonadaceae (the latter group being present in S. siderea but summed under ‘other’ in Fig. 3) , . Families present at very low relative abundance in S. stellata but not detected in S. siderea were Cytophagaceae and Ruaniaceae , . While it is likely that some of the variation seen between these two coral species is due to methodological differences between the studies, we expect that some of the bacterial-community differences are genuine indicators of biogeographic and species differentiation.
Bacterial diversity of DSS-affected tissues
Gil-Agudelo et al.  found a group of bacteria metabolically similar to Vibrio carchariae present only in cultures from DSS-affected corals, but subsequent inoculation experiments failed to trigger disease signs. We detected many different Vibrio species using the PhyloChip G3, but V. carchariae was not one of them (Table S1). There were several OTUs affiliated with V. campbellii, V. harveyi, and V. orientalis that had increased relative intensities in many of the Virgin Island samples compared to those from Dry Tortugas (Fig. S2). Perhaps this is an indication that the Virgin Island corals are more stressed (and therefore conducive to hosting higher abundance of this opportunistic taxon). The Virgin Islands National Park hosts an order of magnitude more visitors per year than the Dry Tortugas National Park (e.g., in the 2009 sampling year, VIIS had 415,847 visitors versus 52,011 in DRTO; NPS Visitor Use Statistics https://irma.nps.gov/Stats). Further, while a large proportion of the VIIS visitors will reach the easily accessible Hawknest Bay, only a small percentage of the visitors to DRTO reach Loggerhead Key since most day trips visit Garden Key alone.
The only previous study to employ molecular techniques in examining DSS-affected corals found significant differences between the bacterial communities in healthy versus affected colonies of Stephanocoenia intersepta, both by DGGE and clone library analyses . In contrast, our data did not indicate a significant difference between healthy and DSS-affected corals or between corals from different geographic locations (Figs. 2, 3, 4). This could mean that DSS represents a different condition in St. intersepta than in S. siderea, as has been previously suggested due to different lesion morphologies , . It could also be due to the broader taxonomic coverage achieved using the PhyloChip G3 microarray to examine the coral-associated bacterial communities (4,978 OTUs versus 50 clone library sequences).
Specifically, Sweet et al.  found an increase in Cyanobacteria and Actinobacteria associated with all DSS-affected samples and that the Cyanobacteria were dominated by a species of Oscillatoria. The genus Oscillatoria (reference sequence is Pseudocillatoria coralii) is represented by two OTUs out of the 4,978 OTUs shared by all of our samples and was detected in half of our DSS samples (DRTOSSD08, VIISSSD06, VIISSSD09, and VIISSSD10) but none of our healthy (i.e., normally pigmented) samples (Fig. S3). In addition, there were two other cyanobacterial OTUs (53258 – Phormidium and 53730 – Leptolyngbya) that had higher relative intensities in a subset of the DSS samples (VIISSSD06, VIISSSD07, VIISSSD09, VIISSSD10) but were present in both healthy and DSS samples (Fig. S4). All four of these cyanobacterial genera have been associated with black band disease (BBD) , ,  and so clearly have some capacity to negatively impact corals in certain conditions. Unlike BBD, there was no indication of increased sulfate-reducing or sulfur-oxidizing bacterial populations in our DSS samples.
Only two OTUs were unique to DSS and present in more than 50% of (>4) DSS samples in our study: 53656 representing Chroococcidiopsis, a genus of Cyanobacteria in the class Oscillatoriophycideae and OTU 7560 representing an unclassified Vibrio. Sweet et al.  also found two Vibrio species that were only present in DSS-tissues, but concluded that, like V. carchariae , they were probably not directly involved as pathogens. Further, they also detected Oscillatoria sp. in lower abundance in one of their healthy samples, leading them to conclude that this cyanobacterium may contribute to the dark pigmentation of DSS in St. intersepta but is unlikely to be a pathogenic cause . Our S. siderea data corroborate these conclusions (Figs. S2, S3, S4).
Another aspect of the St. intersepta study was that different size DSS lesions were sampled and a difference was found in the bacterial communities between small (1–2 cm) and larger (5–10 cm, >10 cm) spots, suggesting a community shift as the lesion aged . Unfortunately, all of our DSS areas would be categorized as small, although many were collected from the edge of a much larger older lesion (i.e., central dead area with algal recruitment; Fig. S1). This sort of purpling along the edge of an older lesion (e.g., one caused by black band disease) has been described as Dark Spot Syndrome Type II .
Previous histological work on S. siderea colonies from the Bahamas, Little Cayman, Florida Keys, and Puerto Rico described unidentified endolithic fungal cells associated with dark spot lesions , . Sweet et al.'s  recent discovery of a fungal ribotype consistently associated with DSS in St. intersepta that was genetically similar to a fungal plant pathogen (Rhytisma acerinum) that causes ‘tar spot’ disease adds some molecular evidence that DSS may be a fungal disease. Unfortunately, we were unable to amplify fungal 18S rRNA from our S. siderea samples due to the ITS primers being overwhelmed by coral 18S rRNA. Further, due to the collection and processing methods we followed, it was not possible to examine any of the samples by histology to look for physical signs of fungal infection. Given the uncertainty of whether DSS is a disease or a non-specific stress response to any physical, chemical, or microbiological insult , , as well as being unclear if these signs represent different conditions in different coral species , it is imperative that future studies combine histology with microbiology to link potential causal agents to a specific pathology at the cellular level . Future studies in the vein of Closek et al.  may shed more light on what the coral host is doing during these occurrences, providing greater insight into coral immune responses.
S. siderea maintains a geographically-conserved bacterial community with considerable diversity (>600 genera). There is notable overlap between the bacterial community composition of S. siderea and its sister species S. stellata at high levels of taxonomy.
Our data do not support the hypothesis that DSS is a bacterial disease. There was neither a single dominant bacterial group observed in all DSS samples to indicate a primary pathogen, nor was there a community shift toward a specific or predictable secondarily opportunistic consortium. Future work should focus on determining if this discoloration indicates the same type of lesion across different coral species via histological analyses. Metagenomics also could be used to address the presence of a particular fungal ribotype.
Photos showing examples of dark spot lesions sampled in this study. A: DRTOSSD08, B: DRTOSSD10, C: VIISSSD08, D: VIISSSD06, E: VIISSSD07, F: VIISSSD10. Samples with the prefix DRTO are from Dry Tortugas National Park. Samples with the prefix VIIS are from the Virgin Islands National Park.
Vibrionaceae OTUs present in at least two samples are listed with bar lengths representing the relative OTU intensity. Genus and species names are given for each OTU; when the OTU was unclassified at the genus level, the family name was used.
OTU intensity values are displayed in place of samples on a non-metric multidimensional scaling plot (based on a Bray-Curtis similarity matrix) of post-scale normalized data for (A) Oscillatoria_52941 and (B) Oscillatoria_53657 (reference GenBank entry is Pseudoscillatoria coralii). The intensity values ranged from 12 to>25,000 and are shown in exponential notation; e.g., 3E3 = 3103 = 3,000.
Cyanobacterial OTUs present in at least two samples are listed with bar lengths representing the relative OTU intensity. Genus and species names are given for each OTU; when the OTU was unclassified at the genus level, the family name was used.
Field assistance by T. McDole, B.J. Reynolds, and V.H. Garrison is gratefully acknowledged. Any use of trade, firm, or product names is for descriptive purposes only and does not imply endorsement by the U.S. Government.
Conceived and designed the experiments: CAK. Performed the experiments: CAK YMP LMT. Analyzed the data: CAK YMP LMT TZD MAG GLA. Contributed reagents/materials/analysis tools: CAK TZD GLA. Wrote the paper: CAK YMP MAG.
- 1. Harvell CD, Kim K, Burkholder JM, Colwell RR, Epstein PR, et al. (1999) Emerging marine diseases–climate links and anthropogenic factors. Science 285: 1505–1510.
- 2. Pandolfi JM, Bradbury RH, Sala E, Hughes TP, Bjorndal KA, et al. (2003) Global trajectories of the long-term decline of coral reef ecosystems. Science 301: 955–958.
- 3. Cróquer A, Weil E (2009) Changes in Caribbean coral disease prevalence after the 2005 bleaching event. Dis Aquat Org 87: 33–43.
- 4. Hughes TP (1994) Catastrophes, phase-shifts and large-scale degradation of a Caribbean coral reef. Science 265: 1547–1551.
- 5. Rogers C (2009) Coral bleaching and disease should not be underestimated as causes of Caribbean coral reef decline. Proc R Soc B 276: 197–198.
- 6. Richardson LL, Aronson RB (2002) Infectious diseases of reef corals. Proc 9th Int Coral Reef Symp 1: 1225–1230.
- 7. Turnbaugh PJ, Ley RE, Hamady M, Fraser-Liggett C, Knight R, et al. (2007) The human microbiome project. Nature 449: 804–810.
- 8. Veron JEN (2000) Corals of the world. Townsville, Australia: Australian Institute of Marine Science. 1382 p.
- 9. Gil-Agudelo DL, Fonseca DP, Weil E, Garzón-Ferreira J, Smith GW (2007) Bacterial communities associated with the mucopolysaccharide layers of three coral species affected and unaffected with dark spots disease. Can J Microbiol 53: 465–471.
- 10. Sekar R, Kaczmarsky LT, Richardson LL (2008) Microbial community composition of black band disease on the coral host Siderastrea siderea from three regions of the wider Caribbean. Mar Ecol Prog Ser 362: 85–98.
- 11. Sekar R, Mills DK, Remily ER, Voss JD, Richardson LL (2006) Microbial communities in the surface mucopolysaccharide layer and the black band microbial mat of black band-diseased Siderastrea siderea. Appl Environ Microbiol 72: 5963–5973.
- 12. Cárdenas A, Rodriguez-R LM, Pizarro V, Cadavid LF, Arévalo-Ferro C (2012) Shifts in bacterial communities of two caribbean reef-building coral species affected by white plague disease. ISME J 6: 502–512.
- 13. Solano OD, Suarez GN, Moreno-Forero SK (1993) Blanqueamiento coralino de 1990 en el Parque Nacional Natural Corales del Rosario (Caribe, colombiano). Boletín de Investigaciones Marines y Costeras-INVEMAR 22: 97–111.
- 14. Borger JL (2005) Dark spot syndrome: a scleractinian coral disease or a general stress response? Coral Reefs 24: 139–144.
- 15. Garzón-Ferreira J, Gil-Agudelo DL (1998) Another unknown Caribbean coral phenomenon? Reef Encounter 24: 10.
- 16. Goreau TJ, Cervino J, Goreau M, Hayes R, Richardson L, et al. (1998) Rapid spread of Caribbean coral reef diseases. Rev Biol Trop 46: 157–171.
- 17. Borger JL (2003) Three scleractinian coral diseases in Dominica, West Indies: distribution, infection patterns and contribution to coral tissue mortality. Rev Biol Trop 51: 25–38.
- 18. Rogers CS (2010) Words matter: recommendations for clarifying coral disease nomenclature and terminology. Dis Aquat Org 91: 167–175.
- 19. Gil-Agudelo DL, Garzón-Ferreira J (2001) Spatial and seasonal variation of dark spots disease in coral communities of the Santa Marta area (Columbian Caribbean). Bull Mar Sci 69: 619–629.
- 20. Cervino J, Goreau TJ, Nagelkerken I, Smith GW, Hayes R (2001) Yellow band and dark spot syndromes in Caribbean corals: distribution, rate of spread, cytology, and effects on abundance and division rate of zooxanthellae. Hydrobiologia 460: 53–63.
- 21. Borger JL, Steiner SCC (2005) The spatial and temporal dynamics of coral diseases in Dominica, West Indies. Bull Mar Sci 77: 137–154.
- 22. Porter JW, Torres C, Sutherland KP, Meyers MK, Callahan MK, et al. (2011) Prevalence, severity, lethality, and recovery of dark spots syndrome among three Floridian reef-building corals. J Exp Mar Biol Ecol 408: 79–87.
- 23. Weil E, Rogers CS (2011) Coral reef diseases in the Atlantic-Caribbean. In: Dubinsky Z, Stambler N, editors. Coral Reefs: An Ecosystem in Transition. New York: Springer.
- 24. Weil E (2004) Coral reef diseases in the wider Caribbean. In: Rosenberg E, Loya Y, editors. Coral Health and Disease. Berlin: Springer. pp. 35–68.
- 25. Work TM, Richardson LL, Reynolds TL, Willis BL (2008) Biomedical and veterinary science can increase our understanding of coral disease. J Exp Mar Biol Ecol 362: 63–70.
- 26. Gil-Agudelo DL, Smith GW, Garzón-Ferreira J, Weil E, Petersen D (2004) Dark spots disease and yellow band disease, two poorly known coral diseases with high incidence in Caribbean reefs. In: Rosenberg E, Loya Y, editors. Coral Health and Disease. Berlin: Springer. pp. 337–350.
- 27. Gochfeld DJ, Olson JB, Slattery M (2006) Colony versus population variation in susceptibility and resistance to dark spot syndrome in the Caribbean coral Siderastrea siderea. Dis Aquat Org 69: 53–65.
- 28. Correa AMS, Brandt ME, Smith TB, Thornhill DJ, Baker AC (2009) Symbiodinium associations with diseased and healthy scleractinian corals. Coral Reefs 28: 437–448.
- 29. Voss JD, Richardson LL (2006) Coral diseases near Lee Stocking Island, Bahamas: patterns and potential drivers. Dis Aquat Org 69: 33–40.
- 30. Pantos O, Cooney RP, Le Tissier MDA, Barer MR, O'Donnell AG, et al. (2003) The bacterial ecology of a plague-like disease affecting the Caribbean coral Montastrea annularis. Environ Microbiol 5: 370–382.
- 31. Kellogg CA, Piceno YM, Tom LM, DeSantis TZ, Gray MA, et al. (2013) Comparing bacterial community composition between healthy and white plague-like disease states in Orbicella annularis using PhyloChip G3 microarrays. PLOS ONE 8: e79801.
- 32. Sunagawa S, Woodley CM, Medina M (2010) Threatened corals provide underexplored microbial habitats. PLOS ONE 5: e9554.
- 33. Kellogg CA, Piceno YM, Tom LM, DeSantis TZ, Zawada DG, et al. (2012) PhyloChip microarray comparison of sampling methods used for coral microbial ecology. J Microbiol Meth 88: 103–109.
- 34. Galkiewicz JP, Kellogg CA (2008) Cross-kingdom amplification using Bacteria-specific primers: complications for studies of coral microbial ecology. Appl Environ Microbiol 74: 7828–7831.
- 35. Hazen TC, Dubinsky EA, DeSantis TZ, Andersen GL, Piceno YM, et al. (2010) Deep-sea oil plume enriches indigenous oil-degrading bacteria. Science 330: 204–208.
- 36. McDonald D, Price MN, Goodrich J, Nawrock EP, DeSantis TZ, et al. (2012) An improved Greengenes taxonomy with explicit ranks for ecological and evolutionary analyses of bacteria and archaea. ISME J 6: 610–618.
- 37. Edgar R, Domrachev M, Lash AE (2002) Gene Expression Omnibus: NCBI gene expression and hybridization array data repository. Nucl Acids Res 30: 207–210.
- 38. Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, et al. (2010) QIIME allows analysis of high-throughput community sequencing data. Nat Methods 7: 335–336.
- 39. Clarke KR (1993) Non-parametric multivariate analyses of changes in community structure. Aus J Ecol 18: 117–143.
- 40. Sweet M, Burn D, Croquer A, Leary P (2013) Characterization of the bacterial and fungal communities associated with different lesion sizes of dark spot syndrome occurring in the coral Stephanocoenia intersepta. PLOS ONE 8: e62580.
- 41. Hsiang T, Tian X (2007) Sporulation and identity of tar spot in maple in Canada. Acta Silv Lign Hung Special Edition 71–74.
- 42. Kramarsky-Winter E, Arotsker L, Rasoulouniriana D, Siboni N, Loya Y, et al. (2013) The possible role of cyanobacterial filaments in coral black band disease pathology. Microb Ecol 1–9.
- 43. Miller AW, Richardson LL (2011) A meta-analysis of 16S rRNA gene clone libraries from the polymicrobial black band disease of corals. FEMS Microbiol Ecol 75: 231–241.
- 44. Rasoulouniriana D, Siboni N, Ben-Dov E, Kramarsky-Winter E, Loya Y, et al. (2009) Pseudoscillatoria coralii gen. nov., sp. nov., a cyanobacterium associated with coral black band disease (BBD). Dis Aquat Org 87: 91–96.
- 45. Barneah O, Ben-Dov E, Kramarsky-Winter E, Kushmaro A (2007) Characterization of black band disease in Red Sea stony corals. Environ Microbiol 9: 1995–2006.
- 46. Brandt ME, McManus JW (2009) Disease incidence is related to bleaching extent in reef-building corals. Ecol 90: 2859–2867.
- 47. Gray MA, Stone RP, McLaughlin MR, Kellogg CA (2011) Microbial consortia of gorgonian corals from the Aleutian islands. FEMS Microbiol Ecol 76: 109–120.
- 48. Probst AJ, Lum PY, John B, Dubinsky EA, Piceno YM, et al. (2014) Microarray of 16S rRNA gene probes for quantifying population differences across microbiome samples. In: He Z, editor. Microarrays: Current Technology, Innovations and Applications: Caister Academic Press. pp. 99–120.
- 49. Lins-de-Barros MM, Cardoso AM, Silveira CB, Lima JL, Clementino MM, et al. (2013) Microbial community compositional shifts in bleached colonies of the Brazilian reef-building coral Siderastrea stellata. Microb Ecol 65: 205–213.
- 50. Lins-de-Barros MM, Vieira RP, Cardoso AM, Monteiro VA, Turque AS, et al. (2010) Archaea, bacteria, and algal plastids associated with the reef-buliding corals Siderastrea stellata and Mussismilia hispida from Búzios, South Atlantic Ocean, Brazil. Microb Ecol 59: 523–532.
- 51. Frias-Lopez J, Bonheyo GT, Jin Q, Fouke BW (2003) Cyanobacteria associated with coral black band disease in Caribbean and Indo-Pacific reefs. Appl Environ Microbiol 69: 2409–2413.
- 52. Gantar M, Sekar R, Richardson LL (2009) Cyanotoxins from black band disease of corals and from other coral reef environments. Microb Ecol 58: 856–864.
- 53. Renegar DA, Blackwelder PL, Miller JD, Gochfeld DJ, Moulding AL (2008) Ultrastructural and histological analysis of Dark Spot Syndrome in Siderastrea siderea and Agaricia agaricites. Proceedings of the 11th International Coral Reef Symposium 185–189.
- 54. Galloway SB, Work TM, Bochsler VS, Harley RA, Kramarsky-Winter E, et al. (2007) Coral Disease and Health Workshop: Coral Histopathology II. NOAA Technical Memorandum NOS NCCO 56 and NOAA Technical Memorandum CRCP 4. Silver Spring, MD: National Oceanic and Atmospheric Administration. 84 p.
- 55. Mydlarz LD, Holthouse SF, Peters EC, Harvell CD (2008) Cellular responses in sea fan corals: granular amoebocytes react to pathogen and climate stressors. PLOS ONE 3: e1811.
- 56. Pollock FJ, Morris PJ, Willis BL, Bourne DG (2011) The urgent need for robust coral disease diagnostics. PLOS Pathogens 7: e1002183.
- 57. Work T, Meteyer C (2014) To understand coral disease, look at coral cells. EcoHealth 1–9.
- 58. Closek CJ, Sunagawa S, DeSalvo MK, Piceno YM, DeSantis TZ, et al. (2014) Coral transcriptome and bacterial community profiles reveal distinct Yellow Band Disease states in Orbicella faveolata. ISME J In Press.