Symbiodinium associations with diseased and healthy scleractinian corals

DOI 10.1007/s00338-008-0464-6 REPORT Symbiodinium associations with diseased and healthy scleractinian corals A. M. S. Correa Æ M. E. Brandt Æ T. B. Smith Æ D. J. Thornhill Æ A. C. Baker Received: 16 June 2008 / Accepted: 18 December 2008 Ó Springer-Verlag 2009 Abstract Despite recent advances in identifying the causative agents of disease in corals and understanding the impact of epizootics on reef communities, little is known regarding the interactions among diseases, corals, and their dinoflagellate endosymbionts (Symbiodinium spp.). Since the genotypes of both corals and their resident Symbiodinium contribute to colony-level phenotypes, such as thermotolerance, symbiont genotypes might also contribute to the resistance or susceptibility of coral colonies to disease. To explore this, Symbiodinium were identified using the internal transcribed spacer-2 region of ribosomal DNA from diseased and healthy tissues within individual coral colonies infected with black band disease (BB), dark spot syndrome (DSS), white plague disease (WP), or yellow blotch disease (YB) in the Florida Keys (USA) and the US Communicated by Biology Editor Dr. Ruth Gates A. M. S. Correa M. E. Brandt A. C. Baker Division of Marine Biology and Fisheries, Rosenstiel School of Marine and Atmospheric Science, University of Miami, 4600 Rickenbacker Cswy, Miami, FL 33149, USA A. M. S. Correa (&) A. C. Baker Department of Ecology, Evolution and Environmental Biology, Columbia University, MC 5557, 1200 Amsterdam Avenue, New York, NY 10027, USA e-mail: amr2007@columbia.edu T. B. Smith Center for Marine & Environmental Studies, University of the Virgin Islands, 2 John Brewers Bay, St. Thomas, Virgin Islands 00802, USA D. J. Thornhill Department of Biology, Bowdoin College, 6500 College Station, Brunswick, ME 04011, USA Virgin Islands. Most of the diseased colonies sampled contained B1, B5a, or C1 (depending on host species), while apparently healthy colonies of the same coral species frequently hosted these types and/or additional symbiont diversity. No potentially parasitic Symbiodinium types, uniquely associated with diseased coral tissue, were detected. Within most individual colonies, the same dominant Symbiodinium type was detected in diseased and visually healthy tissues. These data indicate that specific Symbiodinium types are not correlated with the infected tissues of diseased colonies and that DSS and WP onset do not trigger symbiont shuffling within infected tissues. However, few diseased colonies contained clade D symbionts suggesting a negative correlation between hosting Symbiodinium clade D and disease incidence in scleractinian corals. Understanding the influence of Symbiodinium diversity on colony phenotypes may play a critical role in predicting disease resistance and susceptibility in scleractinian corals. Keywords Coral Dark spot syndrome Disease Internal transcribed spacer-2 (ITS-2) White plague disease Introduction Disease outbreaks in the Caribbean have led to the mortality of scleractinian corals over large reef tracts (Porter et al. 2001; Miller et al. 2006), resulting in changes to coral community structure (Bruckner and Bruckner 1997, 2006; Aronson and Precht 2001) and the loss of structural reef habitats (Hughes 1994). Most coral disease research in the last two decades has therefore focused on the etiology and ecosystem-level impacts of diseases (Richardson 1998; Sutherland et al. 2004; Bruno et al. 2007). However,

another critical prerequisite for predicting and managing epizootics is to identify genetic differences that underlie disease resistance or susceptibility between individuals and populations (Lesser et al. 2007). To identify these phenotypes in coral algal symbioses, it may be necessary to explore functional differences not only between coral genotypes, but also between different types of Symbiodinium, the dinoflagellate algal endosymbionts living within coral tissues. Although relatively little is known regarding the interactions among diseases, corals, and Symbiodinium spp. (but see Ulstrup et al. 2007; Danovaro et al. 2008; Roff et al. 2008a, b; Stat et al. 2008), functional differences between algal symbionts are known to influence the distribution of reef coral colonies (e.g., Iglesias-Prieto et al. 2004) and coral responses to environmental stress (Rowan et al. 1997; Glynn et al. 2001; Baker 2004; Berkelmans and van Oppen 2006; Jones et al. 2008). Experiments have shown a genetic basis for some physiological differences between symbionts (e.g., Warner et al. 2006), and these differences have been found to contribute to coral colony phenotypes such as thermotolerance. Symbiodinium contains eight subgeneric clades (A H, e.g., Pochon et al. 2006) and numerous types within these clades (reviewed by Baker 2003). The internal transcribed spacer-2 (ITS-2) region of the ribosomal DNA has been used most widely to differentiate symbiont types in the Caribbean (e.g., LaJeunesse 2002). While some ITS-2 types have been shown to exhibit ecological patterns such as those described above (e.g., Frade et al. 2008), functional differences still need to be determined for much of the extensive genetic diversity within Symbiodinium (van Oppen and Gates 2006). Exploring the relationships between Symbiodinium and coral disease incidence may reveal the functional significance of symbiont genotypes. For example, Toller et al. (2001) sampled Symbiodinium from 12 colonies of Montastraea annularis (sensu lato) displaying disease signs consistent with yellow blotch disease (YB). They reported atypical clade A symbionts within YB lesions on 11 out of 12 of the colonies (based on the small-subunit 18S rdna), whereas adjacent, healthy tissue from the same colonies contained clade C symbionts which are commonly detected in healthy colonies of these species at the sampled depths. These atypical symbionts were suggested as potentially parasitic Symbiodinium that flourished at the expense of their host. A logical extension of Toller et al. (2001) work is to characterize these atypical Symbiodinium genotypes at a higher resolution (e.g., using the ITS-2), and ultimately to test their functional significance as disease-inducing or parasitic types. It is also possible that the onset of disease within coral tissues impacts symbiont community composition. In most individual coral colonies, typically 1 or 2 Symbiodinium variants are numerically dominant (Knowlton and Rohwer 2003; Baird et al. 2007). However, high-sensitivity techniques, such as real-time PCR, indicate that many corals likely contain additional cryptic Symbiodinium diversity present at low abundances (Mieog et al. 2007). Environmental changes may induce shifts within Symbiodinium communities, providing opportunities for previously rare types to become dominant within individual colonies (Baker 2003). Elevated sea surface temperatures, excess ultraviolet radiation, altered irradiance, and the presence of pollutants are the environmental factors that affect coral algal symbioses (Glynn 1996; Brown et al. 2000; Douglas 2003; Jones 2004; Bruno et al. 2007). The onset of disease in coral tissues, however, might also change the in hospite endosymbiont environment (e.g., decreased penetration of photosynthetically active radiation, alteration of the host chemical environment by bacterial communities, or decrease in nutrient [nitrogen] transfer) to the extent that shifts occur in the dominant Symbiodinium type within infected tissues. Hypothetically, the diseases most likely to induce shifts in symbiont communities are those that: (1) persist in the same area of coral tissue for long periods and (2) affect environmental parameters known to influence Symbiodinium associations with healthy colonies. Finally, certain Symbiodinium species could impart a measure of disease resistance or susceptibility to their host colonies. The Coral Probiotic Hypothesis (Reshef et al. 2006; Rosenberg et al. 2007) suggests that diverse populations of microorganisms live in symbiosis with corals and that the relative abundances of these organisms are modified as environmental conditions change, allowing the coral holobiont (coral and symbionts) to adapt to new conditions. Much of the research focus in terms of this hypothesis has been on populations of bacteria (e.g., Pantos et al. 2003; Pantos and Bythell 2006; Ritchie 2006), which are suggested to be the most diverse and abundant probiotic organisms within corals (Rohwer et al. 2002). However, Symbiodinium are also symbiotic microbes that can be used to test predictions of the Coral Probiotic Hypothesis. The Symbiodinium communities associated with diseased scleractinian coral tissues, and their relationship to coral epizootics, are largely uncharacterized. To provide a more complete view of disease resistance and susceptibility at the colony level, this study aimed to: (1) determine the diversity of dominant symbionts associated with diseased scleractinian corals in the Florida Keys (USA) and the U.S. Virgin Islands (USVI); (2) identify any Symbiodinium types uniquely associated with diseased coral tissue; (3) determine whether diseased and visually healthy tissues within the same coral colony contain different dominant symbionts; and (4) determine whether particular Symbiodinium taxa are correlated with host colony disease resistance or susceptibility.

Materials and methods Study sites and collections Samples were collected from 71 diseased scleractinian coral colonies (comprising 12 coral species and 4 diseases, Table 1) at four sites (Cheeca Patch, Coral Gardens, French Reef, and Little Grecian) in the Florida Keys (USA) and seven sites in the US Virgin Islands (USVI) (Fig. 1) using SCUBA during April and September 2006 (Florida Keys) and April 2007 (USVI). Representative infections identified as dark spot syndrome (DSS), white plague disease (WP), yellow blotch disease (YB), and black band disease (BB) are shown in Fig. 2; species presented similar signs in both sampled geographic locations. DSS was identified as discolored dark areas of tissue on colonies Table 1 Disease, host species, location, collection depth, and genetic identity (ITS-2 type) of Symbiodinium samples collected from diseased coral colonies in the Florida Keys, USA (FK) and St. Thomas and St. John, US Virgin Islands (USVI) Disease Host Location Depth (m) Symbiodinium ITS-2 type BB Colpophyllia natans FK 3 B1 C. natans USVI 23 C3L Diploria strigosa (2, 2) FK, USVI 4, 1 8 B1 Montastraea cavernosa (2) FK 3 C3 DSS Agaricia sp. FK 6 C3a Colpophyllia natans FK 3 B1 Dichocoenia stokesii (1, 1) FK, USVI 7, 6 B1 Diploria labyrinthiformis FK 4 B1 Meandrina meandrites (3) USVI 11 12 B1 Siderastrea siderea (17, 1) FK, USVI 2 10, 4 B5a S. siderea (1, 1) FK, USVI 5, 10 B5a, C3 S. siderea (8) USVI 2 10 C1 S. siderea USVI 3 C1, D1a S. siderea FK 4 D: D1a, H: A3 a S. siderea (2) USVI 3 7 D1a Siderastrea radians (2) USVI 4 5 C46 WP Colpophyllia natans USVI 20 B1 C. natans USVI 20 D: B1, H: C1 Diploria strigosa USVI 11 B1 Montastraea annularis USVI 4 D: C3, H: C7, D1a Montastraea faveolata (5) FK 2 7 B1 M. faveolata USVI 20 D1a Montastraea franksi USVI 23 D: B1, C1 a, H: B1 M. franksi USVI 23 D: B1, C7 a, H: C7 a M. franksi USVI 23 C7 a M. franksi USVI 23 D: C7 a, H: C7 a, D1a M. franksi USVI 20 D: C7 a, D1a, H: C7 a Montastraea sp. USVI 23 C7 YB Montastraea annularis USVI 8 B1, C1, D1a M. annularis USVI 4 D: C1, D1a, H: D1a Montastraea faveolata FK 4 B1 M. faveolata FK 8 B10 M. faveolata USVI 18 C7 BB Black band disease, DSS dark spot syndrome, WP white plague disease, YB yellow blotch disease. Numerals in parentheses next to host names indicate the number of colonies independently sampled from each location. For colonies that contained different symbionts in their diseased and visually healthy tissues, D: and H: designate the ITS-2 types identified from each respective part of the colony. All other colonies contained the listed ITS-2 types in both their diseased and visually healthy tissues a Coral Symbiodinium combinations that have not been reported previously (based on LaJeunesse 2002; Santos and LaJeunesse 2006; and the compiled data in Baker and Romanski 2007)

South Florida, U.S.A. CG CP CR TA LG TR FR 20 km 90 0 W N U.S. Virgin Islands 20 0 N Black Point Range Cay Flat Cay St. Thomas St. John Fish Bay Seahorse Cottage Shoal Buck Island South Capella 5 km 0 500 1,000 km Fig. 1 Coral collection sites in the Florida Keys, FL (USA) and St. Thomas and St. John, US Virgin Islands (USVI). CG = Coral Gardens; CP = Cheeca Patch; CR = Conch Reef; FR = French Reef; LG = Little Grecian; TA = Tavernier Rocks; TR = Triangles. Only diseased coral colonies were sampled at Cheeca Patch, Coral Gardens, and French Reef (Florida Keys) and at Seahorse Cottage Shoal and South Capella (USVI). Only apparently healthy coral colonies were sampled at Conch Reef, Tavernier Rocks, and Triangles (Florida Keys) (Borger 2005). WP signs were identified as a sharp line of rapid tissue loss initiating at the base or margin of a colony (Bythell et al. 2004). Although disease progression rates were not measured in the USVI, tissue loss occurred at an average rate of 0.7 cm 2 /day in Florida, which is consistent with rates originally described for white plague type I by Dustan (1977). BB was identified as a line of thick black material delineating recently denuded skeleton from live tissue (Antonius 1973), and YB as yellowed areas of tissue occurring as blotches or lines on a colony (Cervino et al. 2001). Two polyps were biopsied from the top of each diseased coral colony: one from actively diseased tissue and one from healthy tissue 2 cm away from the diseased tissue margin (Fig. 3). Polyp biopsies were performed using a 60 ml syringe (without a needle) held flush with the rim of a corallite. The plunger of the syringe was pulled out, sucking the polyp tissue into the syringe. The syringe inlet was then placed into a 13 mm plastic Swinney filter holder (Pall, PA, USA) containing a glass fiber A/D filter (3.1lm pore size, Pall, PA, USA) and the tissue was expelled onto the filter by depressing the syringe plunger. The syringe was flushed with seawater and debris removed from the inlet by hand prior to each biopsy. After each dive, filters were removed from their filter holders using tweezers, placed into 1.5 ml microcentrifuge tubes and preserved with 95% EtOH. This syringe technique does not remove skeletal material and minimizes damage to the coral colony (full recovery typically occurs within 2 to 4 weeks), making it particularly useful for experiments that require repetitive micro-sampling, or that involve rare or endangered corals. Coral individuals were photographed before and after biopsies were taken (Fig. 3, see also Kemp et al. (2008) for a similar sampling method). A total of 48 apparently healthy Siderastrea siderea and Montastraea faveolata colonies were sampled at four sites (Conch Reef, Little Grecian, Tavernier Rocks and Triangles) in the Florida Keys (USA during June 2001, May 2003 (M. faveolata only) or March 2006, and from five sites (Black Point, Buck Island, Fish Bay, Flat Cay, and Range Cay) in St. Thomas and St. John, USVI (S. siderea only) during April 2007 (Fig. 1). In this study, samples from apparently healthy colonies collected in 2006 (M. faveolata, N = 6; S. siderea, N = 4) represent part of

Fig. 2 Representative images of a black band disease (BB), b dark spot syndrome (DSS), c white plague disease (WP), and d yellow blotch disease (YB) identified during this study. In Fig. 2b, two corallites are white; polyps were sampled from these corallites prior to photographing D DSS 2 cmh healthy before after Fig. 3 Generalized depiction of sampling strategy for diseased coral colonies. Schematic shows a colony infected with dark spot syndrome, with healthy and dark spot-infected tissue ( DSS ) labeled accordingly. Each diseased sample ( D ) was taken from a polyp centrally located within an infected tissue patch or band. After sample D was collected, the nearest periphery of the infected tissue patch or band was located and a healthy polyp (sample H ) was collected 2 cm directly outward from the diseased periphery. Before and after photo inset demonstrates the high efficacy of the syringe sampling method in terms of removing tissue with minimal skeletal damage. Color differences between photos are due to differences in ambient lighting, not the visual appearance of the colony Thornhill et al. s (2006b) ongoing monitoring of individual colonies at Little Grecian (Fig. 1); these samples were collected and extracted as described in that reference. All other samples were collected from the tops of colonies using a hammer and core with a 1.3 cm diameter and were preserved in 95% EtOH. It should be noted that in the Florida Keys, healthy colonies were available at all diseased reefs sampled; diseased and healthy colonies were sampled from different reefs in the Florida Keys (except for Little Grecian) due to sampling permit constraints. Molecular analyses Samples were extracted using a modified organic protocol (Baker et al. 1997). The Symbiodinium ITS-2 region was amplified from each sample using the primers ITSintfor 2 and ITS 2 clamp (LaJeunesse and Trench 2000) under the following conditions: an initial denaturing step of 948C for 3 min followed by 35 cycles of 1 min at 948C, 1 min at 588C, and 1 min at 748C, followed by a single cycle of 7 min at 748C. Products were electrophoresed on 1.2% agarose gels to check for amplification success. Strong amplifications were maintained at -208C. Samples amplifying weakly during the initial PCR underwent an extra processing step to increase the number of target amplicons for downstream applications. This extra step entailed excising the target fragment band from the 1.2% agarose gel and beating it with autoclaved ground glass blast media (Grainger, FL, USA) for 2 min in 700 ll of ddh 2 0; this material was then stored at 208C overnight. The resultant liquid was then used as template in a 2nd PCR reaction the following day and amplification success was again checked on a 1.2% agarose gel. Ultimately, amplified products (resulting from either 1 or 2 PCR reactions) from all

samples were electrophoresed on denaturing gradient gels (DGGE, 35 75%) using a CBS Scientific system (Del Mar, CA, USA). Prominent band characteristics of unique profiles (as described by LaJeunesse 2002) were excised, bead beaten, and re-amplified using the ITSintfor 2 and ITS 2 clamp primers (without the GC clamp) under the conditions described above (constituting either a 2nd or 3rd PCR reaction). Sequencing was performed using Big Dye Terminator v. 3.1 cycle sequencing kit and an Applied Biosystems 3730xl DNA Analyzer (Foster City, CA, USA). Sequences were assembled and edited using the Vector NTI TM Advance 10 software (Invitrogen, Carlsbad, CA, USA) and were then identified using BLAST searches of GenBank to determine the most similar known Symbiodinium type cf. LaJeunesse (2001). As a check of GenBank identifications, sequences were grouped according to clade and aligned with a database of known Symbiodinium diversity for that clade using Clustal2 (Thompson et al. 1997). Pairwise alignment parameters included a gap opening of 10.00 and a gap extension of 0.10. Multiple alignment parameters included a gap opening of 15.00 and a gap extension of 0.30 (Hall 2001). Clade-level alignments were corrected manually and then trimmed to the length of the shortest sequence; final alignments (including gaps) for clades A D were 156, 241, 334, and 292 base pairs long, respectively. TCS version 1.21 (Clement et al. 2000) was then used to group identical sequences within each clade-level alignment with 95% certainty. Gaps were treated as a fifth character state. Sequences within the same TCS-defined haplotype were all scored as the same Symbiodinium type; each haplotype took the name of the lowest alphanumeric Symbiodinium type cf. LaJeunesse (2001) that it contained. GenBank and TCS-derived identifications of sequenced samples were further checked by comparing the DGGE gel banding pattern produced by a sequenced sample with the published DGGE banding profile of its putative Symbiodinium ITS-2 match based on GenBank and TCS. The Symbiodinium ITS-2 types within non-sequenced samples were subsequently identified by matching the DGGE gel banding pattern of each nonsequenced sample to the banding pattern of a sequenced sample on the same gel. Statistical analyses For each disease, the null hypothesis (that the same dominant symbiont is present in infected and visually healthy tissues of each diseased colony) was tested. To undertake these comparisons, each sampled colony was designated as having the same or different dominant symbionts in its infected versus healthy tissues. Samples containing multiple types were scored as same only if each type was present in both infected and healthy tissues. The number of actual same versus different colonies was then compared to a theoretical null distribution for the same sample size, in which all colonies were designated as same. Chi-square tests were used to examine BB, DSS, WP, and YB symbiont distributions; the null hypothesis was rejected for a given disease if the actual number of sampled same versus different colonies differed significantly from the null distribution. In each sampled region for M. faveolata (Florida Keys only) and for S. siderea (Florida Keys and USVI), Shannon s diversity index was calculated for the total Symbiodinium community sampled from: (1) diseased and visually healthy tissues of diseased S. siderea colonies and (2) apparently healthy colonies. Shannon s t-test (Hutcheson 1970) was then used to test the differences in the total diversity of Symbiodinium communities in diseased versus apparently healthy colonies. Diversity statistics were calculated using the Biodiversity Calculator (Danoff-Burg and Xu 2005). These analyses were conducted separately for each region to avoid confounding measures of total Symbiodinium diversity with biogeographic changes in the distribution of symbiont diversity within a given coral host species. A likelihood ratio chi-squared test was used to test the differences in the presence of Symbiodinium ITS-2 type D1a within (1) diseased and visually healthy tissues of diseased S. siderea colonies versus (2) apparently healthy colonies of S. siderea (both regions combined). Results The most frequently observed disease-host combination in the Florida Keys and USVI in shallow depths (\10 m) was DSS on S. siderea (N = 31, 54% of diseased colonies observed). In the USVI at C20 m, WP signs were most frequently observed in the genus Montastraea (N = 7, 70% of diseased colonies observed). A summary of the diseases detected and the coral species from which they were sampled is presented in Table 1. All Symbiodinium identified in this study were previously known ITS-2 types (Fig. 4, Tables 1, 2). The most commonly detected ITS-2 types within diseased coral tissues were B1, B5a, and C1. S. siderea in the Florida Keys predominantly hosted clade B symbionts, while conspecifics in the USVI most often contained members of Symbiodinium clade C (Tables 1, 2). Three previously unreported host-symbiont combinations were detected (7 of 119 colonies, based on LaJeunesse 2002; Santos and LaJeunesse 2006; and the compiled data in Baker and Romanski 2007): Montastraea franksi with C1 or C7 and S. siderea with A3. In general, individual colonies contained the same dominant symbiont types within diseased versus healthy

A3 B1 A3 B1 B5a B10 C1 C3 B5 B5a INT{ B10 C1 C3 C3a C3L C7 C46 D1a C3L C3a tissues; this included all individual colonies with BB (N = 8), 98% of colonies with DSS (N = 42) and 80% of colonies with YB (N = 5, Table 1). Colonies with these diseases did not differ significantly from the null distribution, in which all colonies had the same dominant symbiont within its diseased and visually healthy tissues. However, 38% of colonies with WP (N = 16) contained different dominant symbionts within diseased and visually healthy tissues (v 2 = 7.385, P \ 0.01); the null hypothesis was therefore rejected for WP-infected colonies. In the Florida Keys, the Shannon diversity index (H 0 ) of the total Symbiodinium community collected from diseased S. siderea colonies (H 0 = 0.5671) was significantly lower than that detected in apparently healthy conspecifics (H 0 = 1.3654, P \ 0.01 for two-tailed Shannon s t-test, Fig. 5a). In this region, 11% of DSS-infected S. siderea (N = 19) hosted multiple clades/types, while 39% of apparently healthy conspecifics from the same region (N = 13) harbored multiple clades/types. Meanwhile in the USVI, the total Symbiodinium community collected from diseased S. siderea colonies (H 0 = 1.0776) was also significantly lower than that from apparently healthy conspecifics (H 0 = 1.4912, p \ 0.05 for two-tailed Shannon s t-test). In this region, 15% of DSS-infected S. siderea (N = 13) hosted multiple clades/types, while 64% of healthy conspecifics from the region (N = 14) harbored multiple clades/types (Tables 1, 2). Overall, apparently healthy C7 C46 D1a HET D1 Fig. 4 Representative PCR-DGGE ITS-2 fingerprints (profiles) of Symbiodinium types observed in diseased and healthy coral colonies from the Florida Keys (USA) and St. Thomas and St. John, US Virgin Islands. Alphanumeric designation for each symbiont type is given above the corresponding fingerprint profile: uppercase letters indicate clade, numbers represent ITS-2 type, and lowercase letters (if present) denote a rdna paralog (intragenomic variation that is a diagnostic part of the fingerprint). Arrows indicate diagnostic bands excised from the gel and sequenced. Many other bands were also excised and sequenced, however, the majority of these were either: (1) intragenomic variation that is not a diagnostic part of a fingerprint (denoted INT ); (2) heteroduplexes (mismatched DNA strands formed during the re-annealing stage of PCR, denoted HET ); or (3) host DNA sequences (not present in this figure) { S. siderea colonies (N = 27) contained significantly more Symbiodinium ITS-2 type D1a than diseased conspecifics (N = 32) (D1a in apparently healthy colonies: N = 10, 37% vs. diseased colonies: N = 4, 13%; v 2 = 4.872, P \ 0.05, Fig. 5b). In M. faveolata colonies from the Florida Keys, the total Symbiodinium community diversity of diseased (WP- or YB-infected, H 0 = 0.4101) colonies was not significantly different from that of apparently healthy colonies (H 0 = 0.7393). However, 10% of healthy conspecifics (N = 21) hosted multiple clades/types, while diseased M. faveolata colonies (N = 7) did not harbor multiple clades/types. Furthermore, Symbiodinium ITS-2 type D1a was detected from apparently healthy M. faveolata colonies (colonies with D1a: N = 6 of 21, 29%), but not from diseased conspecifics (colonies with D1a: N = 0 of 7, 0%). Discussion DSS and WP did not induce symbiont community change Within most individual colonies sampled, the same Symbiodinium type(s) were detected from diseased and visually healthy tissues (Table 1); BB-, DSS-, and YB-infected colonies did not differ significantly from the null distribution in which diseased and visually healthy tissues contained the same symbionts. The likelihood of committing a Type II statistical error (failure to reject the null hypothesis when an alternative hypothesis is actually true) based on these data is low for DSS infections, due to a robust sample size. The null hypothesis was rejected only for WP-infected colonies; however, it is unlikely that WP infection itself induced the differences observed in these colonies. Most of the colonies containing different symbionts in WP-infected and visually healthy tissues were members of the Montastraea species complex. This species complex commonly hosts multiple symbiont types distributed heterogeneously over the colony surface (Rowan and Knowlton 1995; Rowan et al. 1997; Kemp et al. 2008). In such host species, disease-induced changes to symbiont communities would be indicated by: (1) differences in the symbiont composition within infected and visually healthy tissue; and (2) repeated identification of the same symbiont type from the infected tissues of colonies, regardless of the symbiont diversity detected from adjacent, visually healthy tissues. Repeated identification of the same symbiont type from infected tissues is important because it indicates that the detected differences in symbiont distribution within colonies are directional (i.e., the disease constitutively changes infected tissues such that a particular symbiont type is

Table 2 Host species, location, number of colonies (N), collection depth and genetic identity (ITS-2 type) of Symbiodinium samples collected from apparently healthy Montastraea faveolata and Siderastrea siderea colonies in the Florida Keys, FL (FK, USA) and St. Thomas and St. John, US Virgin Islands (USVI, USA) a Coral Symbiodinium combinations that have not been reported previously (based on LaJeunesse 2002; Santos and LaJeunesse 2006; and the compiled data in Baker and Romanski 2007) Host Location N Depth (m) Symbiodinium ITS-2 type Montastraea faveolata FK 15 2 7 B1 1 3 B1, D1a 1 5 B10, D1a 4 3 D1a Total M. faveolata FK 21 2 7 B1, B10, D1a Siderastrea siderea FK 1 6 A3 a, B1 1 3 B1 5 3 4 B5a 2 5, 10 B5a, C3 1 4 B5a, C3, D1a 1 10 B5a, D1a 1 10 C3 1 3 D1a Total S. siderea FK 13 3 10 A3 a, B1, B5a, C3, D1a S. siderea USVI 1 3 B1, C1, C3 3 2 5 B1, D1a 3 5 9 B5a, C3 1 3 C1, D1a 3 5 8 C3 1 7 C3, D1a 2 4 6 D1a Total S. siderea USVI 14 2 9 B1, B5a, C1, C3, D1a (a) 100 ** * (b) 100 * Percent of colonies sampled 80 60 40 20 Percent of colonies sampled 80 60 40 20 Diseased Healthy N = 19 N = 13 Siderastrea siderea, FK Diseased Healthy N = 13 N = 14 S. siderea, USVI WP & YB N = 7 Montastraea faveolata, FK = clade A = clade B = clade C = clade D Healthy N = 21 Diseased Healthy Diseased Healthy N = 32 N = 27 N = 7 N = 21 S. siderea, FK & USVI M. faveolata, FK = no ITS-2 type D1a = ITS-2 type D1a detected detected Fig. 5 a b Symbiodinium diversity in diseased and apparently healthy colonies from the Florida Keys, FL (USA) and St. Thomas and St. John, US Virgin Islands. In Fig. 5a, * and ** denote significant differences (P \ 0.05 and P \ 0.01, respectively, for twotailed Shannon s t-test) in the total Symbiodinium community repeatedly favored within infected tissues), and that symbiont differences between infected and visually healthy tissues are not random (e.g., due to heterogeneous distribution of symbionts prior to disease infection). Five different symbionts were detected in the WP-infected tissues sampled in this study; it is unlikely that WP infection diversity within diseased versus apparently healthy corals. In Fig. 5b, * denotes significant differences (P \ 0.05 for likelihood ratio chisquared test) in the number of diseased colonies containing Symbiodinium ITS-2 type D1a versus apparently healthy conspecifics. FK = Florida Keys; USVI = US Virgin Islands bestowed a competitive advantage on all of these different types (Table 1). Since the same symbiont type was not repeatedly identified from infected tissues, the most parsimonious explanation for the detected differences within the sampled WP-infected Montastraea colonies is that they contained heterogeneous distributions of symbionts prior to

disease infection, and that the differences were not induced by WP infection itself. Taken together, these data indicate that DSS and WP do not induce symbiont community shifts in the tissues they infect. Kirk et al. (2005) also found no difference in Symbiodinium associated with diseased and healthy tissue in sea fans exhibiting aspergillosis, a disease caused by the fungus Aspergillus sydowii. In this study, only one colony infected with BB or YB (N = 13) contained different symbionts within its diseased and visually healthy tissues; however, additional infected colonies must be sampled to conclusively support the null hypothesis for these diseases. In particular, additional YB-infected colonies should be examined, since: (1) differences have been detected elsewhere in the Caribbean (Toller et al. 2001) and (2) YB progresses relatively slowly through host tissue at rates of approximately 5 15 cm year -1 (Cervino et al. 2001; Bruckner and Bruckner 2006) and therefore might alter the environment experienced by Symbiodinium over time scales that would allow for symbiont community shifts. Symbiodinium diversity in diseased and healthy colonies Symbiont diversity within diseased corals was similar to that of apparently healthy conspecifics in that all samples contained known Symbiodinium ITS-2 types; no novel types were detected uniquely from diseased coral tissue. Three of the coral symbiont combinations detected during this study have not been published previously (denoted with a in Tables 1, 2), but in contrast to Toller et al. s (2001) report of an atypical symbiont restricted to YB lesions on Montastraea spp. colonies, these were not specifically associated with a disease infection. Instead, these coral symbiont combinations likely reflect that the diversity of coral algal associations has been underestimated due to a lack of sufficient sampling with adequately sensitive genetic techniques (Apprill and Gates 2007 but see Thornhill et al. 2007; Baird et al. 2007; Baker and Romanski 2007). Only six studies have examined Caribbean Symbiodinium diversity in scleractinian corals using the ITS-2 marker (LaJeunesse 2002; Diekmann et al. 2003; Banaszak et al. 2006; Thornhill et al. 2006a, b; Frade et al. 2008) and of these, only three sampled C5 individuals per coral host species. As the total number of colonies sampled per host species increases, additional novel coral symbiont combinations are likely to be identified. Overall, diseased S. siderea colonies contained significantly lower Symbiodinium community diversity, and fewer harbored multiple clades/types, as compared to apparently healthy S. siderea colonies from the same region and depth range (Fig. 5a, Tables 1, 2). Similarly, fewer Symbiodinium clades were detected overall from diseased versus apparently healthy M. faveolata colonies (Florida Keys only), although the overall symbiont diversity was not significantly different between these groups. Although apparently healthy M. faveolata and S. siderea colonies from the Florida Keys were collected in 2001, 2003 (M. faveolata only), and 2006, while diseased colonies were sampled from this region in 2006, differences in the total symbiont diversity within diseased versus healthy colonies of these species are unlikely to be an artifact of sampling. Thornhill et al. (2006b; pers. comm.) have monitored the symbiont diversity within individual colonies of M. faveolata and S. siderea in the Florida Keys since 1998 and 2002, respectively. They showed that colonies of these species generally did not change their symbiont types from 2000/2002 through 2007, although some fluctuations in the relative abundance of different Symbiodinium types were observed. Since the dominant symbionts present in these colonies during 2001 are known to be representative of what the colonies contained in 2006, it can reasonably be assumed that samples from other apparently healthy colonies of M. faveolata and S. siderea taken from these study areas in 2001 and 2003 are also representative of the dominant symbionts present in 2006. Furthermore, healthy and diseased USVI S. siderea colonies were both sampled during April 2007 and, in this region, healthy colonies also contained a higher diversity than diseased colonies. Why do diseased S. siderea colonies contain lower symbiont diversity compared to healthy conspecifics? One possible explanation is that symbiont shifts are triggered by disease onset and occur pervasively throughout both the infected and visually healthy tissues of diseased colonies, resulting in lower overall symbiont diversity relative to that occurring in apparently healthy conspecifics. However, it seems rather unlikely that only some tissue areas show disease signs if all tissue of an infected colony is affected by disease onset. It is more parsimonious to assume that disease-related symbiont shifts would be restricted to colony tissue areas showing disease signs. Based on the within-colony comparisons of infected and visually healthy tissue reported here, DSS does not induce symbiont community shifts that are restricted to the infected tissues of diseased colonies. Therefore taken together, it is unlikely that Symbiodinium diversity within diseased colonies was reduced after infection with DSS. The most parsimonious explanation for the observed pattern is that a connection exists between the Symbiodinium diversity that S. siderea colonies host and their relative disease susceptibility. Symbiodinium D1a as a possible disease-resistant type In this study, a negative correlation was observed between hosting Symbiodinium type D1a and disease incidence in

S. siderea (Fig. 5b). Furthermore, type D1a was identified from only 10 out of the 60 diseased colonies that could have potentially hosted it (Table 1), based on previous detections of clade D from conspecifics (Colpophyllia natans, Diploria labyrinthiformis, M. annularis, Montastraea cavernosa, M. faveolata, M. franksi, Siderastrea radians, and S. siderea are all capable of hosting clade D; based on Santos and LaJeunesse 2006, the compiled data in Baker and Romanski 2007, and the apparently healthy colonies in this study). For example, although nearly 30% of the apparently healthy M. faveolata colonies sampled contained type D1a, this type was not detected within diseased conspecifics (Fig. 5b). These observations led to the hypothesis that Symbiodinium D1a plays a role in disease resistance. Symbiodinium ITS-2 type D1a could directly confer a measure of disease resistance to its coral host cf. the Coral Probiotic Hypothesis (Reshef et al. 2006). It has been suggested that coral-associated bacteria could provide corals with disease resistance by niche exclusion of pathogenic bacteria from the coral surface and tissues, and by producing antibiotics (Reshef et al. 2006). Symbiodinium could potentially act in either of these capacities; they are commonly found on coral surfaces and in mucus, and it has recently been documented that some Symbiodinium are capable of producing terpenes, a class of hydrocarbons that frequently exhibit anti-microbial or anti-inflammatory properties (Newberger et al. 2006). First steps in determining whether Symbiodinium ITS-2 type D1a directly confers disease resistance to coral colonies include: (1) investigating whether Symbiodinium type D1a is capable of producing terpenes; and (2) testing predictions of the Coral Probiotic Hypothesis (Reshef et al. 2006, p. 2072) in terms of Symbiodinium. It is also possible that ITS-2 type D1a confers disease resistance to corals indirectly. Lesser et al. (2007) suggest that many coral diseases result from opportunistic, nonspecific bacteria that exploit the compromised health of coral individuals that have been exposed to environmental stressors, such as elevated temperature. Some coral species have been observed to switch from thermally sensitive symbionts to symbionts in clade D following thermal bleaching events (Jones et al. 2008). Furthermore, evidence indicates that coral colonies hosting Symbiodinium clade D are less likely to bleach in response to temperature stress (Baker et al. 2004; Rowan 2004; Berkelmans and van Oppen 2006). A positive correlation has also been demonstrated between bleaching and disease in coral colonies (Muller et al. 2007). Taken together, these findings may indicate that corals hosting type D1a are less susceptible a priori to opportunistic diseases because they are less likely to suffer weakened health status and/or bleach as a result of environmental stressors such as elevated temperature. This negative correlation between hosting Symbiodinium ITS-2 type D1a and disease incidence should be confirmed for a diversity of scleractinian corals (e.g., M. faveolata) and diseases, and further explored through monitoring health status and symbiont community structure within individual colonies over time as well as with manipulative experiments that directly test the susceptibility of different holobionts to disease. It is crucial that subsequent studies track the dominant Symbiodinium within coral colonies and relate this to the bleaching, disease, and survival trajectories of holobionts. Although some clade D symbionts may function as a safety parachute, enabling colonies to survive short-term thermal stress events (Berkelmans and van Oppen 2006; Jones et al. 2008), there may be tradeoffs to hosting clade D that can affect the long-term fitness of its coral host. For example, juveniles of two Acropora species hosting clade D grew two to three times slower than juvenile conspecifics hosting clade C (Little et al. 2004, but see Smith et al. 2008). Thus, further observation and experimentation are necessary to determine the environmental conditions and timescales under which symbionts in clade D lead to a net cost or benefit to corals inclusive fitness. This study represents the first comprehensive survey of Symbiodinium communities associated with multiple species of diseased scleractinian corals. Although unusual or novel symbiont types were not found in association with diseased tissues, three novel coral Symbiodinium combinations were identified, supporting the notion that diversity in coral Symbiodinium associations has generally been underestimated due to small within-species sample sizes. The data presented here indicate that DSS- and WPinfections do not result from or induce shifts in the dominant Symbiodinium type within colonies. Colonies hosting Symbiodinium ITS-2 type D1a, however, may be indirectly resistant to opportunistic pathogens because their health status is less likely to be compromised by environmental stress. The frequent detection of Symbiodinium types B1, B5a, and C1 in diseased colonies implies that coral individuals hosting these types may be relatively susceptible to environmental stressors and therefore prone to infection by opportunistic disease agents. If the percentage of coral colonies hosting ITS-2 type D1a increases on reefs, this may lead to a decrease in the incidence of scleractinian coral diseases in the Caribbean (as a result of better overall colony health). Further study of Symbiodinium distributions within diseased and healthy coral colonies should increase our understanding of scleractinian coral disease resistance and susceptibility. Acknowledgments This work was conducted in the Florida Keys under National Marine Sanctuary permits FKNMS-2006-013-A1 (to AMSC) and FKNMS-2001-030 (to ACB), and under Florida Fish and Wildlife Conservation Commission permit 01S-620 (to ACB). Collections in the US Virgin Islands were made under Division of Fish

and Wildlife Collection/Export Permit No. STT017-07 (to TBS). We thank Joanne Delaney and Brian Keller for FKNMS permit support and Bill Valley and the Florida Keys National Marine Sanctuary for boat use. We thank William K. Fitt, Gregory W. Schmidt, and Dustin W. Kemp for their contributions to sample collection and molecular analysis. We are grateful to Mark D. Fitchett for his advice on statistical analysis. We thank Craig Starger for sequencing work and Rob DeSalle and George Amato for sequencing support through the Sackler Institute for Comparative Genomics at the American Museum of Natural History. We are grateful to Mary Alice Coffroth, Peter Glynn and three anonymous reviewers, whose comments improved previous versions of this manuscript. 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