Introduction

Coral reefs provide goods and services that are valuable to many millions of people throughout tropical coastal areas (Moberg and Folke 1999). Reef-building corals (Order Scleractinia) are central to coral reefs, providing much of the productivity and calcification required to build these ecosystems (Muller-Parker and D’Elia 1997). These organisms form mutualistic symbioses with dinoflagellates Symbiodinium spp., which provide corals with abundant photosynthetic energy, enabling them to lay down copious quantities of calcium carbonate and thrive in nutrient-poor environments (Muller-Parker and D’Elia 1997).

Unfortunately, coral reefs are facing a number of serious threats and are severely in decline (Hughes et al. 2003; Hoegh-Guldberg et al. 2007). These threats are arising from declining coastal water quality and over-fishing, as well as warming and acidification of the world’s oceans as a result of rising atmospheric carbon dioxide and other greenhouse gases. Rapid changes in water temperature, for example, have caused sudden breakdown of the mutualistic endosymbiosis of corals and dinoflagellates (mass coral bleaching; Hoegh-Guldberg 1999). Mass bleaching events have had a serious impact on coral reefs throughout the world since 1979 when they were first reported in the literature. Our current understanding of the temperature tolerance of corals suggests that projected sea temperatures will soon approach and exceed the known thermal thresholds to reef-building corals, putting in doubt the future of coral-dominated reef systems (Hoegh-Guldberg 1999; Hoegh-Guldberg et al. 2007).

Due to the potentially devastating impacts of climate change on reef-building corals and coral reefs in general, considerable attention has been given to the physiology and ecology of coral–dinoflagellate symbioses, particularly on the factors and mechanisms that cause its maintenance or breakdown (see Lesser 1997; Ralph et al. 2001). Other studies have focused on the diversity, phylogeny (e.g., Carlos et al. 1999; Lajeunesse 2001; Santos et al. 2002; Rodriguez-Lanetty 2003; Coffroth and Santos 2005), biogeography, community ecology (e.g., Rodriguez-Lanetty et al. 2001; LaJeunesse et al. 2003; Sampayo et al. 2007) and the acquisition of Symbiodinium (e.g., Lewis and Coffroth 2004; Pasternak et al. 2006; Gomez-Cabrera et al. 2008), which forms an important base for understanding the dynamics of coral–dinoflagellates endosymbiosis. By contrast, however, studies exploring the biology and ecology of Symbiodinium in other reef microhabitats are rare.

Studies on the diversity of Symbiodinium have revealed that it is a very diverse group with at least nine distinct genetic clades, A-I (Baker 2003; Coffroth and Santos 2005; Stat et al. 2006; Pochon and Gates 2010). Clades A, B, C and D are the predominant symbionts of scleractinians, while clade E is found in sea anemones, and clades F, G and H are common in foraminifera (Baker 2003; Coffroth and Santos 2005; Stat et al. 2006). Clades C and D can also inhabit in foraminifera, while clades F and G sometimes can be found, although rarely, in scleractinians (Rodriguez-Lanetty et al. 2002; Pochon et al. 2004, 2007; Pochon and Pawlowski 2006). Clade I was recently discovered, and it establishes symbiosis with foraminifera (Pochon and Gates 2010). Clade C has a very wide range of hosts, which include a marine ciliate (Lobban et al. 2002) in addition to scleractinian and foraminiferan hosts (Pochon et al. 2001, 2004, 2007; Pochon and Pawlowski 2006).

Within each Symbiodinium clade, there is even more diversity grouped in subclade types (Baker 2003; Coffroth and Santos 2005; Stat et al. 2006). Clade C is the most diverse Symbiodinium lineage in the Pacific with more than 100 subclade types (LaJeunesse et al. 2003; Pochon et al. 2004; Sampayo et al. 2007). Some of these subclade types can be identified with several molecular markers including large subunit ribosomal DNA (LSUrDNA) and internal transcriber spacers (ITS) (LaJeunesse et al. 2003; Sampayo et al. 2009). Comprehensive reviews of Symbiodinium diversity can be found in LaJeunesse (2001), Baker (2003), LaJeunesse (2005), Coffroth and Santos (2005), and Stat et al. (2006).

Symbiodinium spend part of their life cycle as free-living Gymnodinium-like dinoflagellates and in some cases re-infect corals each generation (Gomez-Cabrera et al.2008; Adams et al.2009). Despite the fairly extensive information on Symbiodinium as coral symbionts, there are a number of key questions surrounding these organisms. For example, our understanding of the importance of other habitats that they use, their population dynamics through space and time, and how they are taken back into host corals cells remain incomplete. This said, there are a growing number of studies indicating that Symbiodinium are present in the seawater column (Gou et al. 2003; Coffroth et al. 2006; Manning and Gates 2008; Pochon et al. 2010), interstitial water of sands (Carlos et al. 1999; Hirose et al. 2008; Pochon et al. 2010), rocky reefs and seagrasses (Coffroth et al. 2006) and on benthic macroalgae (Porto et al. 2008). Moreover, it has been demonstrated that both pelagic and benthic Symbiodinium can establish symbiosis with corals (Lewis and Coffroth 2004; Coffroth et al. 2006; Adams et al.2009), although some genotypes are apparently unable to establish symbiosis (Coffroth et al. 2006; Pochon et al. 2010).

Macroalgae are the most abundant benthic component of many coral reefs (Wilkinson 2004; Diaz-Pulido 2008). Benthic macroalgae release organic substances (Khailov and Burlakova 1969; Wada et al. 2007), which may promote the establishment of microbial communities on their surfaces (Armstrong et al. 2000; Longford et al. 2007). Also, many epiphytic dinoflagellates show a distinct preference for macroalgal hosts which may release growth-stimulating algal compounds or provide large surface areas for attachment (Morton and Faust 1997; Parson and Preskitt 2007). Consequently, there is a strong possibility that macroalgae may serve as an important reservoir for Symbiodinium within coral reef habitats.

Porto et al. (2008) found Symbiodinium associated to the benthic macroalgae Halimeda spp., Lobophora variegata, Amphiroa spp., Caulerpa spp. and Dictyota spp. in Caribbean coral reefs. However, the potential role of macroalgae as a source of Symbiodinium to infect reef-building corals still remains an important unknown. This study explores the presence, identity and potential relationship of macroalgal-associated Symbiodinium to those associated with reef-building corals on the southern Great Barrier Reef. Specifically, this study evaluates whether the Symbiodinium subclades found in macroalgal microhabitats are the same as those found in reef-building corals.

Materials and methods

Sample collection

Seawater samples were collected from macroalgal microhabitats and sediments in coral reefs from the Heron and Keppel Islands on the southern Great Barrier Reef, Australia (Fig. 1). Four types of samples were taken:

Fig. 1
figure 1

Location of study sites: a southern Great Barrier Reef, b Heron Island and Keppel Islands. Specific collection sites are also shown around c Keppel Islands and d Heron Island. (Images modified from Google Earth©)

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  1. (1)

    Complete sections of macroalgal thalli and cyanobacterial mats were collected with associated seawater, taking care to avoid pieces of the substrate to which the macroalgae were attached.

  2. (2)

    Crustose coralline algae (CCA) and tiny algal turfs were collected with associated seawater and substrates, which were broken off from the calcareous matrix.

  3. (3)

    Sediments were collected with associated seawater.

  4. (4)

    Seawater from the interstitial space of sediments and from the surface of algal turfs or CCAs was collected with 50 ml syringes.

Field collection was made with 0.5-L, 1-L and 2-L plastic bags, depending on the amount of material, and some samples needed more than one 2-L bag. A total of 33 macroalgal samples (including one cyanobacterial sample and four CCA samples) and four sediment samples were collected (Table 1). Two sediment samples were collected near sand-dwelling algal turfs (one from 2 m deep in Heron Island and the another from 12 m deep in Keppel Islands), while two others were collected away from macroalgae.

Table 1 Results of molecular analyses (PCRs, cloning and sequencing) for macroalgal and sediment samples
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Each sample was vigorously shaken after collection and filtered through a 200-μm-pore-size mesh. Samples were then filtered again through a 0.5-μm Millipore GFC using a vacuum pump (Capex 8C). The differential pressure in the filter was about 720 mbar. To preserve the potential Symbiodinium DNA content, each filter paper was immersed in 20% Dimethylsulfoxide (DMSO) within a dark flask (covered with aluminium foil) and transported to the Centre for Marine Studies at The University of Queensland, where the samples were stored in a freezer (−20°C) until the DNA extraction.

DNA extraction

In order to remove the DMSO, which is undesirable for the following procedures, each filter paper was cut in smaller pieces and rinsed in DNA-Buffer [50 mM EDTA (pH 8.0), 0.4 M NaCl]. DNA was then isolated from filter papers with the Phenol–Chloroform method, following the protocol of Vidal et al. (2002). This protocol was originally designed to improve the DNA extraction from CCA but also enhanced DNA extractions from any alga. This protocol is desirable for samples with low or indeterminate amounts of DNA. The steps involved in this protocol that were intended for cleaning up and grinding CCAs were skipped, given the different material. Consequently, the isolation of the DNA was started by transferring pieces of each filter paper to a 2-ml eppendorf tube containing 700 μl of extraction buffer (4 M Urea, 250 mM Tris–HCl (pH 8.0), 250 mM NaCl, 50 mM EDTA (pH 8.0), 5% 2-Mercaptoethanol, 2% Sodium dodecyl sulphate) and 15 μl of Proteinase K (20 mg ml−1). The final dried pellet was re-suspended in 50 μl of TE buffer (10 mM Tris–HCL (pH 8.0), 1 mM EDTA). The quality and concentration of the extracted DNA was analyzed through gel electrophoresis and NanoDrop spectrophotometry (Thermo Scientific, Wilmington, USA); samples with less than 15 ng μl−1 of DNA were concentrated through vacuum centrifuging at 37°C for 20 min.

PCRs, cloning and sequencing

The variable domains D1 and D2 of 28S large subunit ribosomal DNA (28S-LSUrDNA) were used given that they provide moderate resolution of Symbiodinium to the subclade level (e.g., LaJeunesse et al. 2003; Sampayo et al. 2009). D1 and D2 28S-LSUrDNA of potential Symbiodinium were amplified using the Toha PCR primer set (see Rodriguez-Lanetty et al. 2001): forward (Toha F): 5′-CCT CAG TAA TGG GGA ATG AAC A-3′ and reverse (Toha R): 5′-CCT TGG TCC GTG TTT CAA GA-3′. All PCR contained >15 ng of template DNA, 1× PCR buffer, 2.5 mM MgCl2, 0.2 mM of each primer, 200 mM dNTP and 0.1 mM Taq polymerase platinum, and filter-sterilized water for a total volume of 20 μl. The PCR conditions involved an initial denature period of 2 min at 94°C, followed by 30 cycles of 15 s at 94°C, 15 s at 60°C, 60 s at 72°C and a final extension period of 5 min at 72°C. After the PCR, the samples were held at 4°C. The PCR products were purified with the QIAquick PCR purification kit, QIAGEN.

The different 28S-LSUrDNA fragments contained at each sample were separated and cloned in TOP TEN cells (Invitrogen, AU) by using the pGEM-T Vector System, following the manufacturer’s protocol. For ligation, 32 ng of PCR products (3:1 insert:vector molar ratio) was used. Four clones per library were PCR-amplified at their 28S-LSUrDNA, purified with QUIAquick and sent to the Australian Genomic Research Facility for sequencing. Unfortunately, because of time limitations, it was not possible to sequence more clones per library.

Data analysis

Amplified and sequenced DNA was compared to Symbiodinium sequences using the Basic Local Alignment Search Tool (BLAST; Altschul et al. 1990) and the GenBank database. Those Symbiodinium sequences that showed the highest similarity scores with the resulting sequences and E-values <e−10 were downloaded. 28S-LSUrDNA sequences of the Symbiodinium subclade types C1, C1b, C3, C15, C17, C21 and C27, described and reported by LaJeunesse et al. (2003), were also downloaded. Additionally, at least two 28S-LSUrDNA sequences of the Symbiodinium clades A, B, C, D, E, F and H were also downloaded.

All resulting and GenBank sequences were aligned using ClustalX 2.0. Sequences that did not belong to Symbiodinium (according to the BLAST search) were not aligned. Phylogenetic inference analyses were performed with the aligned sequences to determine the phylogenetic position of Symbiodinium isolates obtained in this study (Table 2). Sequences that belonged to Symbiodinium, according to the BLAST analyses but had too much baseline noise, were excluded from the phylogenetic analyses. The phylogenetic analyses were then performed with the Maximum-Parsimony and Maximum-Likelihood methods, using the best fit model of evolution (TrN + G) according to jModeltest (Posada 2008). These analyses were run using the beta version of the PAUP 4.0 software (Sinauer Associates, Massachusetts, USA). Gaps were treated as a fifth character state, starting trees were obtained via stepwise addition, sequence addition was simple, and it used the tree-bisection-reconnection (TBR) branch-swapping algorithm to find the best tree(s). If more than one tree was found, then the 50% Majority-Rule Consensus Tree was calculated. Five hundred bootstrap replicates were performed for each analysis.

Table 2 List of Symbiodinium 28S-LSUrDNA sequences used in this study for phylogenetic tree reconstruction
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Results

The 28S-LSUrDNA sequences from a total of 25 samples were amplified. Of these, 24 were cloned while the remaining sediment sample was directly sequenced and identified as a non-symbiotic dinoflagellate. Among 96 clones that were sequenced (four clones per library), 21 were identified as belonging to Symbiodinium (~22% of clones sequenced) and 16 as non-symbiotic dinoflagellates. The remaining 59 clones were either of poor-quality or had sequences similar to platyhelminthes or non-symbiotic ciliates (e.g., Paramecium). Symbiodinium were associated with samples from the green macroalgae Chlorodesmis fastigiata, Halimeda opuntia and H. discoidea, the red macroalgae Hypnea spinella and H. pannosa, Laurencia intricata and Asparagopsis taxiformis, the brown macroalgae Padina sp. and Lobophora variegata, and algal turfs (see Fig. 2, Tables 1 and 2).

Fig. 2
figure 2

Representative examples of macroalgae that were shown to harbor Symbiodinium spp. a and b Algal turfs from Heron Island, c Asparagopsis taxiformis from Keppel Islands, d Hypnea pannosa from Heron Island, e Lobophora variegata from Keppel Islands, f Padina sp. from Heron Island, g Chlorodesmis fastigiata from Heron Island, h Halimeda opuntia from Heron Island

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Non-symbiotic dinoflagellates (e.g., Gyrodinium, Karlodinium and Prorocentrum) were associated with samples from Porolithon onkodes (CCA), cyanobacterial mats, sediments and also Halimeda discoidea, Hypnea pannosa, Laurencia intricata, Asparagopsis taxiformis, Lobophora variegata and algal turfs. All Symbiodinium found in macroalgal microhabitats belonged to the clade C and were similar to strains of Symbiodinium that establish symbiosis with corals, according to the BLAST search (score >920; E-value = 0.0; similarity >98%) (Table 3). Only Dc11 from Lobophora variegata (score = 720) showed a score <920 in the BLAST search.

Table 3 List of Symbiodinium isolated from macroalgal microhabitats and their subclades (according to the phylogenetic analysis)
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The phylogenetic analyses revealed that all Symbiodinium isolated from macroalgal microhabitats belonged to the clade C (Fig. 3). Interestingly, however, they are spread between at least two subclades. Strain Dc58, isolated from Hypnea pannosa, grouped with Symbiodinium C15 (a strain that normally associates with the coral Porites; LaJeunesse et al. 2003); strains Dc25, Dc54, Dc60, Dc70, Dc72 grouped with Symbiodinium C3 (a strain associating with Acropora, Favia and a large number of other coral genera; LaJeunesse et al. 2003). Dc30 grouped with Symbiodinium C17 (a strain that associates with Montipora; LaJeunesse et al. 2003) but with a very low bootstrap support (<50%). Strains Dc1, Dc2, Dc3, Dc4, Dc6, Dc8, Dc15, Dc21, Dc27, Dc68, Dc82, Dc88 did not clearly fall within any subclade. Sequences Dc36 and Dc11 were not included in the phylogenetic analyses because they had too much baseline noise. Phylogenetic positions of macroalgal-associated Symbiodinium did not vary much among trees generated by Maximum-Parsimony and Maximum-Likelihood. The tree inferred by Maximum-Parsimony (data not shown) was a consensus (50% majority rule) of ten trees. Subclades C3 and C15 were supported with bootstrap values of 96 and 88%, respectively.

Fig. 3
figure 3

Phylogenetic tree inferred by Maximum-Likelihood (ML) from the 28S-LSUrDNA of 19 macroalgal-associated Symbiodinium and 31 Symbiodinium reference sequences, downloaded from the GenBank. Clades, subclades and ML bootstrap values >50% are indicated at their respective branches. Names of strains indicate their host/environmental origin. Colors next to macroalgal-associated strains indicate their macroalgal phyla (green macroalgae, red macroalgae or brown macroalgae)

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Discussion

The present study has found evidence that macroalgae may serve as potential microhabitats for Symbiodinium spp. when they are outside their cnidarian hosts. Symbiodinium were found associated with macroalgae from a wide variety of depths and sites (Heron-reef flat, Heron-4th point, Keppel-Middle Island, Keppel-Halfway Island) on the southern Great Barrier Reef. Symbiodinium were associated with a wide variety of macroalgal forms that included foliose (Padina sp., Lobophora variegata), turfing filamentous (mixed algal turfs), non-turfing filamentous (Chlorodesmis fastigiata), calcareous articulated (Halimeda spp.) and corticated macrophytes (Laurencia intricata, Hypnea spp., Asparagopsis taxiformis). Symbiodinium also associated with a range of macroalgal phyla including Chlorophyta (green algae: Chlorodesmis fastigiata, Halimeda spp.), Ochrophyta (Phaeophyceae, brown algae: Lobophora variegata, Padina sp.) and Rhodophyta (red algae: Asparagopsis taxiformis, Hypnea spp., Laurencia intricata).

While the present study did not find any Symbiodinium associated with Cyanobacteria, or the red macroalgae Porolithon onkodes, Plocamium sp. and Peyssonnelia sp., the finding of other dinoflagellates that were associated with Porolithon onkodes and Cyanobacteria suggest that it would be too early to rule out these macroalgae as potential microhabitats for Symbiodinium. Moreover, the presence of Symbiodinium associated to Plocamium sp. and Peyssonnelia sp. was evaluated on few samples per species (see Table 1), thereby limiting the chance of finding Symbiodinium.

Symbiodinium was not found to be associated with sediments on the southern Great Barrier Reef. The presence of Symbiodinium in reef sediments, however, has been demonstrated by previous studies in the Caribbean Sea (Porto et al. 2008) and Pacific Ocean (Carlos et al. 1999; Hirose et al. 2008; Adams et al. 2009; Pochon et al. 2010). Previous studies have emphasized the limitations of directly extracting DNA from sediments, especially the action of sediment-associated substances interfering with the DNA extraction process (Steffan et al. 1988; Lovell and Piceno 1994; Gray and Herwig 1996). For that reason, studies reporting sediment-associated Symbiodinium have generally used direct isolation and culture of potential candidates before molecular identification (e.g., Carlos et al. 1999; Hirose et al. 2008). Given that the present study did not use direct isolation and culture, it would be premature to rule out the presence of Symbiodinium in sediments on the southern Great Barrier Reef.

Previous studies of free-living Symbiodinium have shown contrasting effectiveness at recovering Symbiodinium sequences from clone libraries. Littman et al. (2008) were 1% efficient (four Symbiodinium among 319 clones) using 18S rDNA, while Manning and Gates (2008) were >90% efficient using a hypervariable region of 23S (see Table 2 of Manning and Gates 2008). A rapid comparison of this study with these studies suggests that the recovery of Symbiodinium from clone libraries was moderately efficient here (22%; 21 Symbiodinium among 96 clones). The very low efficiency found by Littman et al. (2008) may be explained by the fact that their genetic analyses were employed on sediments samples where molecular analyses have limitations as explained above.

Most Pacific corals harbor Symbiodinium clade C, although most Caribbean corals, on the other hand, are associated with Symbiodinium clades B, with significant numbers of Caribbean corals also having clades A and C (Baker et al. 1997; LaJeunesse et al. 2003). The same biogeographic distribution was also found in Symbiodinium from the water column by Manning and Gates (2008). Interestingly, the present study only found Symbiodinium clade C to be associated with macroalgae in the southern Great Barrier Reef, while Porto et al. (2008) found a large range of Symbiodinium clades (A, B and C) associated to benthic macroalgae in Caribbean coral reefs. Nevertheless, it is likely that the diversity of macroalgal-associated Symbiodinium was underestimated in this study as it explored as many macroalgal microhabitats as possible but the number of sequenced clones per library was low. It is possible that more Symbiodinium clades/subclades appeared if more clones per library were sequenced.

Subclades Dc25, Dc54, Dc60, Dc70 and Dc72 from the present study grouped with Symbiodinium C3 (96% bootstrap support; Figs. 3, Table 3). This is interesting given that C3 is a generalist subclade which establishes symbioses with many coral species in Pacific and Caribbean coral reefs (LaJeunesse et al. 2003; Table 3) as well as some foraminifera (Pochon et al. 2007). The occurrence of this strain in several samples illustrates the potential importance of macroalgae as a source of Symbiodinium for a huge number of coral species. Strain Dc58 grouped with Symbiodinium C15 (88% bootstrap support), which associates with thermally tolerant corals (Porites spp. and Montipora digitata; LaJeunesse et al. 2003) and foraminifera (Pochon et al. 2004). It is believed that thermal tolerance of those hosts is given by Symbiodinium C15. Red macroalgae were the only macroalgal phylum that grouped with symbiotic strains of Symbiodinium (i.e., C3 and C15), while green and brown macroalgae did not clearly associate to any subclade. Given the fact that this pattern may change if more clones per library were sequenced, it will require verification in follow-up studies.

It is possible that some macroalgal-associated Symbiodinium found in the present study were in hospite, associated to soritid foraminifera or macroscopic ciliates. The latter have been reported to harbor Symbiodinium clade C and inhabit macroalgal microhabitats (Pochon et al. 2004, 2007; Lobban et al. 2002, Pochon and Pawlowski 2006). It is also possible that some Symbiodinium came from symbiotic larvae; however, the collection of samples occurred in August, which is 2 months before the coral spawning in the Great Barrier Reef (Willis et al. 1985; Babcock et al. 1986). Symbiodinium C15 and C3 have been previously isolated from both corals and foraminifera (Pochon et al. 2004, 2007; Pochon and Pawlowski 2006); thus, it is uncertain whether the macroalgal-associated Symbiodinium C15 and C3 found here were free-living or foraminifera-associated (although careful stereomicroscopic examination of the filter papers with the samples did not find soritid foraminifera). Nevertheless, the presence of Symbiodinium C3 and C15 in Pacific macroalgal microhabitats may also suggest, with limitations, a potential close link between coral zooxanthellae and macroalgal-associated Symbiodinium populations/communities.

It is interesting to consider why Symbiodinium may associate with macroalgae. Porto et al. (2008) suggest that Symbiodinium associate to benthic macroalgae because their intricate branching networks with high surface-to-volume ratios provide substrate, light attenuation and refuge for Symbiodinium. On the other hand, studies on other epiphytic dinoflagellates suggest that they associate with some benthic macroalgae primarily because of the large amount of surface area upon which to attach (Parson and Preskitt 2007) and possibly because they release organic substances that stimulate the growth and provide other resources for epiphytic dinoflagellates (Morton and Faust 1997). Given that the water column associated with coral reefs tends to be nutrient poor, organic substances along with inorganic nutrients released by benthic macroalgae (Khailov and Burlakova 1969; Wada et al. 2007) may play a critical role in the survival of Symbiodinium outside their coral hosts. This idea requires further exploration to establish the relative importance of these macroalgal organic and inorganic compounds for the survival of Symbiodinium.

In conclusion, the present study has shown that Symbiodinium are spread among several macroalgal taxa and functional groups on the southern Great Barrier Reef. Some of these Symbiodinium may be in hospite within foraminifera or ciliates; however, the presence of Symbiodinium C3 and C15 in macroalgal microhabitats may also suggest, with limitations, a potential close link between coral zooxanthellae and macroalgal-associated Symbiodinium communities, and a continuum between symbiotic and environmental Symbiodinium populations. This study has implications for the role of other reef organisms and habitats as potential reservoirs for the symbionts that inhabit reef-building corals. A more complete description of these potential reservoirs is important if we are to continue to improve our understanding of the all-important mutualistic symbiosis between corals and dinoflagellates.