Ultrastructural, Immunohistochemical, and Biochemical Variation Amongst Symbiodinium Isolates by Maria Mazzillo Mays A dissertation submitted to the Graduate Faculty of Auburn University in partial fulfillment of the requirements for the Degree of Doctor of Philosophy Auburn, Alabama August 3, 2013 Keywords: Symbiodinium; symbiosis; coral reef; Aiptasia pallida; symbiosome; cell-cell recogntion Copyright 2013 by Maria Mazzillo Mays Approved by Stephen C. Kempf, Chair, Associate Professor, Department of Biological Sciences Eduardus C. Duin, Associate Professor, Department of Chemistry & Bio-Chemistry Anthony G. Moss, Associate Professor, Department of Biological Sciences Scott R. Santos, Associate Professor, Department of Biological Sciences Christine A. Sundermann, Professor, Department of Biological Sciences ii Abstract Symbiodinium spp. reside intracellularly in a complex symbiosome (host and symbiont- derived) in a specific host-symbiont associations. Symbiodinium is a diverse genus with variation equivalent to that of other dinoflagellate orders. First, this investigation examines antigenic variation in the algal mucilage secretions at the host-symbiont interface. Cultured Symbiodinium from a variety of clades were labeled with one or the other of two antibodies that bind to components of the symbiont mucilage (PC3 antibody, developed using a clade B alga cultured from Aiptasia pallida; BF10 antibody, developed using a clade F alga cultured from Briareum sp.). PC3 antigen was found in cultured Symbiodinium from clades A and B, but not clades C, D, E and F. The correlation between labeling and clade may account for some of the specificity between host and symbiont in the field. Within clades A and B there was variation in the amount of label present. BF10 antigen was more specific and only found in cultures of the same cp23S- rDNA strain the antibody was created against. Since the mucilage forms the host-symbiont interface, variation in its molecular composition is likely to be the source of signals involved in recognition and specificity. Next, I examined variation in the ultrastructural morphology amongst 5 Symbiodinium isolates (2 from clade A, and 1 from clade B, 1 from clade C, and a free-living isolate from clade E) to create a baseline standard for future ultrastructural work and to determine if there is any ultrastructural basis for taxonomic description of species within this genus. Ultrastructural examinations were performed using High Pressure Rapid Freezing. All 5 isolates studied exhibited the standard dinokaryon nucleus, a peripheral multi-lobed chloroplast iii with a single stalked pyrenoid surrounded by a starch cap, a dinoflagellate-type mitochondria with tubular cristae, and various other cellular inclusions. Additional small electron dense inclusions were typically positioned near the cell periphery and may be related to mucocysts sometimes seen in dinoflagellates. Some isolates (CCMP421 and Y109) had a few cells that exhibited large electron dense structures containing calcium oxalate crystals. Cell size amongst different isolates was significantly different and consistent with previous studies on the same or similar isolates. Examination of the ultrastructure of 5 isolates of Symbiodinium revealed that there is very little variation in morphological characters. Therefore, it is unlikely that isolates or species of Symbiodinium can be usefully differentiated on the basis of morphology alone. Finally, in order to continue examining Symbiodinium isolates for variation that may impact the association with a specific host, I investigated the variation of storage products (lipid droplets and starch granules) within 5 isolates. Amongst the 5 isolates examined, the two isolates from clade A contained the least amount of both starch and lipid. However, another small isolate from clade B contained some of the largest amounts of both starch and lipid. A clade F isolate also contained many storage products while the only free-living isolate examined, a clade E isolate, contained more lipid and starch than the clade A symbionts but less than the clade B and F isolates. This variation in storage products amongst isolates grown under the same culture regime reveals physiological differences that may relate to an individual isolate's suitability as a symbiotic partner under certain environmental conditions such as those arising from global climate change. Further biochemical examination of excreted exudate for the presence of carbohydrate and protein components found that in all 5 isolates, more carbohydrate compounds were secreted into the culture medium than protein. Examination of the composition of PC3 antigen, a component of the exudate of some Symbiodinium isolates, found that the antigen is iv composed, at least in part, of carbohydrate structures. The presence of carbohydrate components in the exudate compounds that are associated with the cell surface is consistent with the hypothesis that initiation and perhaps maintenance of the host-symbiont relationship is mediated by a glycan/lectin interaction. v Acknowledgments I would like to thank Dr. Stephen C. Kempf for acting as my major professor and the chair of my dissertation committee and for teaching me much about the art of electron microscopy. I would also like to thank the members of my committee, Dr. Eduardus Duin, Dr. Anthony Moss, Dr. Scott Santos, and Dr. Christine Sundermann for their guidance throughout my graduate career at Auburn. Dr. Michael Miller (AU AMIL), Dr. Roger Bridgman (AU Hybridoma Lab), and Dr. John Dennis (AU Vet) were also vital to my graduate research at Auburn University. Thanks also go to Dr. William Fitt and Dr. Beth Richardson at the University of Georgia for providing use of equipment at UGA. I would also like to thank Ivey Ellis, Shanna Hanes, Katy Prince, Limin Gong, Nate Kirk, Kevin Kocot, and Spyridon Mourtzinis for their assistance and advice in this research. My husband, David, also deserves a huge amount of gratitude for his support during my graduate career. My incredibly large family has also been very supportive in my pursuit of this degree. I would like to thank my grandfather, Vincent Mazzillo for teaching me to be tenacious; my grandmother, Jean Barr for teaching me that I can do anything I put my mind to; my grandfather, Donald Barr for teaching me to be ?always good?. Jean Mazzillo, Vince Mazzillo, Drew Mazzillo, Meredith Teat Mazzillo, Amanda Mazzillo, Coker Mays, Julia Mays, Beth Mays Woodrow, Seth Woodrow, Ryleigh Woodrow, Reagan Woodrow, Mia Smith, Donald Smith, Seralyn Smith, Katherine Smith, Robert Dotzey, Keith Jacobson, and Noelle Easterday have also been a constant source of strength, support and love. vi Table of Contents Abstract ................................................................................................................................... ii Acknowledgments .................................................................................................................... v List of Tables ......................................................................................................................... vii List of Figures ....................................................................................................................... viii List of Abbreviations................................................................................................................ x I. A review of Symbiodinium symbioses with an emphasis on initiation and maintenance of the Cnidaria-Symbiodinium association ...................................................................................... 1 II. Immunohistochemical variation amongst Symbiodinium isolates .................................... 35 III. Ultrastructural variation amongst Symbiodinium isolates ................................................ 57 III. Ultrastructural and biochemical variation amongst Symbiodinium isolates demonstrates variation in storage molecules .......................................................................................... 84 References ..........................................................................................................................105 vii List of Tables Chapter II. Table 1 ................................................................................................................ 42 Chapter II. Table 2 ................................................................................................................ 47 Chapter III. Table 1 ............................................................................................................... 60 Chapter IV. Table 1 ............................................................................................................... 90 Chapter IV. Table 2 ............................................................................................................... 99 viii List of Figures Chapter I. Figure 1 .................................................................................................................. 5 Chapter I. Figure 2 .................................................................................................................. 9 Chapter II. Figure 1 ............................................................................................................... 45 Chapter II. Figure 2 ............................................................................................................... 48 Chapter II. Figure 3 ............................................................................................................... 50 Chapter III. Figure 1 .............................................................................................................. 65 Chapter III. Figure 2 .............................................................................................................. 66 Chapter III. Figure 3 .............................................................................................................. 68 Chapter III. Figure 4 .............................................................................................................. 69 Chapter III. Figure 5 .............................................................................................................. 70 Chapter III. Figure 6 .............................................................................................................. 72 Chapter III. Figure 7 .............................................................................................................. 74 Chapter III. Figure 8 .............................................................................................................. 75 Chapter III. Figure 9 .............................................................................................................. 77 Chapter III. Figure 10 ............................................................................................................ 78 Chapter III. Figure 11 ............................................................................................................ 80 Chapter III. Figure 12 ............................................................................................................ 82 Chapter IV. Figure 1 ............................................................................................................. 94 ix Chapter IV. Figure 2 ............................................................................................................. 96 Chapter IV. Figure 3 ............................................................................................................. 98 Chapter IV. Figure 4 ............................................................................................................101 x List of Abbreviations 2XCTAB cetyltrimethylammonium bromide 5.8S a segment of ribosomal RNA ANOVA Analysis of Variance BF10 antibody created against the mucilage from a clade F Symbiodinium cp23S chloroplast 23S rRNA gene ELISA enzyme linked immunosorbant assay ITS1 internal transcribed spacer 1 ITS2 internal transcribed spacer 2 MFSW millepore filtered sea water np-H2O nanopure filtered water nr28S nuclear ribosomal large subunit nr18S nuclear ribosomal small subunit PBS phosphate buffered saline PC3 antibody created against the mucilage from a clade B Symbiodinium RFLP restriction fragment length polymorphism 1 I. A REVIEW OF SYMBIODINIUM SYMBIOSES WITH AN EMPHASIS ON INITIATION AND MAINTENANCE OF THE CNIDARIAN-SYMBIODINIUM ASSOCIATION. ABSTRACT Endosymbioses are relationships where a symbiont lives inside a host organism. These can be situations where the symbiont lives within a host cavity like the gut, between the cells of host tissues or within the cells themselves. These associations can be vital to the lives of both partners who are joined in symbiosis. Symbiodinium sp. is a common endosymbiont of various invertebrate species that are best known for their associations with cnidarians where they reside in the cytoplasm of host endodermal cells. These dinoflagellates form a diverse genus that was originally thought to be a single pandemic species. The Cnidarian-Symbiodinium partnership is vital not only to the host cnidarian, but also to the health of the entire coral reef ecosystem. The physical and nutritional basis for the coral reef ecosystem relies on the photosynthetically fixed carbon that is translocated from Symbiodinium to its cnidarian host. This association is diverse with a huge genetic variation in symbiont (similar to other dinoflagellate orders), and many potential host organisms. Despite the diversity in members, the association between Symbiodinium and its cnidarian host is usually specific with an association between one host species and one or a few symbiont types. The specificity in this relationship is conferred in a number of different ways. First, some hosts transfer their symbionts directly to their eggs or larvae (vertical transmission). If the larvae are aposymbiotic (have no symbionts) after leaving the parent and obtain the symbionts from the environment (horizontal transmission), specificity can be conferred a number of ways. During infection the host may feed only on material that is likely to contain the preferred symbiont isolate leading to a specific association. Just after 2 infection, specificity may be conferred by a cell-cell recognition process that determines acceptance or rejection of the symbiont. Specificity in the association may be achieved later via a process in which the symbiont better suited to the host environment outcompetes other symbiont types. Over the last 50 years there has been an increase in coral bleaching events. This massive loss of symbionts from host tissues often leads to the death of the host and thus the loss of reef habitat. While the exact cellular mechanisms behind the breakdown of the symbiosis during bleaching are not fully understood, it is clear that both the hosts and symbionts show varying degrees of susceptibility to thermal stress (the major cause of most bleaching events). Once both 1) the process of infection and 2) the mechanisms resulting in bleaching are understood, better management decisions can hopefully be made to ameliorate the devastation of the reef environment that is likely to occur as sea temperatures rise due to global warming. ENDOSYMBIOSIS Symbioses are relationships between at least two different species that can range from parasitism to mutualism (Lewin, 1982). A parasitism is characterized by one member of the symbiosis benefiting (the parasite) while the other is harmed (the host). A mutualism is a relationship in which both members benefit (Lewin, 1982). While these definitions seem straightforward, recent research into different symbioses tends to suggest that many associations are mutualistic under certain conditions and parasitic if the conditions (environmental, health of one member, etc.) change (Lewin, 1982). An endosymbiosis is a specialized symbiotic relationship in which one member resides within or between the cells of the other. Endosymbioses have specialized terminology, the smaller member of the endosymbiosis residing inside the host (either intracellulary or in 3 intercellular space) is called the symbiont. The larger member is the host (Lewin, 1982). In cases where the symbiont resides intracellularly, it is generally encased in at least one layer of host membrane. This membrane is called the symbiosome membrane (Roth and Stacey, 1989). A good example of an endosymbiotic relationship where one member resides intercellularly is between humans and our symbiotic gut bacteria. In most cases this is a mutualism, the symbiont provides the host with certain molecules such as vitamins while the host provides nutrients and an optimal environment for growth (Backhed et al., 2005). A well- studied example of intracellular endosymbiosis is between bacteria in the genus Rhizobia and legumes. Legumes are known as nitrogen-fixing plants; however the endosymbiont Rhizobia actually fixes the nitrogen. Rhizobia live in special anaerobic compartments (an enzyme required for nitrogen fixation, nitrogenase, is oxygen sensitive) in the root nodules of legumes providing the plant with fixed nitrogen. The Rhizobia in turn obtain carbon compounds from the host and live in a protected environment (Young and Johnston, 1989; Sturz, 1997; Oke and Long, 1999). Since the endosymbiont is residing within the host, there is the potential for an immune reaction against the symbiont. For example, the human gut hosts mutualistic bacteria, but can be plagued by disease-causing bacteria such as Mycobacterium paratuberculosis, Listeria monocytogenes, and certain strains of Escherichia coli; these pathogens can be controlled by the host maintaining a mutualism with symbiotic bacteria that bolster the immune system (Damaskos and Kolios, 2008). In endosymbioses there is usually a high degree of specificity between host and symbiont (Sturz, 1997). In the root nodules of legumes, some species of Rhizobium thrive in certain legumes and not in others (Sturz, 1997). Niche partitioning also maintains some endosymbionts in the root nodules and others only in the foliage or stem (Sturz, 1997). 4 Here I am interested in the endosymbiosis between dinoflagellates of the genus Symbiodinium and cnidarian hosts. This association forms the foundation, both physically and nutritionally of the coral reef ecosystem (Richmond and Wolanski, 2011). This symbiosis is an area of current interest due to its recognized importance in providing for extreme biodiversity and the fact that the effects of global climate change have put this association, and therefore the many species living in coral reef ecosystems, at risk (Rodrigues and Grottoli, 2007). SYMBIODINIUM SYMBIOSIS: THE MEMBERS Dinoflagellates of the genus Symbiodinium occur in association with many invertebrate species in the marine environment. In the symbiosis between Symbiodinium and cnidarians, the symbionts provide the host with photosynthetically fixed carbon (e.g., Davy and Cook, 2001; Goodson et al., 2001; Muller-Parker and Davy, 2001). Various studies have concluded that the photosyntheate can be transferred as a variety of different compounds, most commonly glycerol (Muscatine, 1967) or glucose and succinate/fumarate (Whitehead and Douglas, 2003). In return, the host waste products (N, CO2, etc) and UV protection (Lesser, 1989; Muscatine, 1990) are of benefit to the symbiont, as is the protected intracellular environment where the symbionts are maintained. SYMBIODINIUM SP. LIFECYCLE, MORPHOLOGY, AND TAXONOMY ?Zooxanthellae? was a blanket name historically used to refer to dinoflagellates and some diatoms (Brandt, 1881). In this symbiosis it has most often described a group of single-celled photosynthetic symbiotic dinoflagellates of the genus Symbiodinium. Initially, these gymnodinoid symbionts were described morphologically as a single species, Symbiodinium 5 microadriaticum Freudenthal (1962). This species was initially assumed to be pandemic based on ultrastructural studies (Taylor, 1968; Taylor, 1974). The initial described life cycle was based on cultured Symbiodinium, and is presumed to be similar to what is seen in hospite and in free-living dinoflagellates in the genus Symbiodinium. The dominant stage is the vegetative phase (Figure 1A) characterized by a single, thin-walled Figure 1. Symbiodinium microadriaticum life cycle taken from Freudenthal, 1962. A. Vegetative cell. B. Vegetative cell dividing. C. Vegetative cyst. D. Zoospore. E. Motile zoospore. F. Non- motile aplanospore. G. Vegetative cyst division to form two subspecies. H. Gametogenesis? I. Release of gametes? 6 autotrophic cell with a visible chloroplast and an unobtrusive nucleus. Over time, the cell loses some chloroplast volume and the accumulation body grows. Eventually, if this cell continues to grow without dividing it will lose chloroplast volume and die (Freudenthal, 1962). However, the vegetative phase generally undergoes cell division (Figure 1B), producing two daughter cells each with half the parent cytoplasm and organelles, with the exception of the accumulation body which is transferred whole to one of the daughter cells. The vegetative cell may also form a cyst with a thickened cell wall (Figure 1C) that can become a motile zoospore (Figure 1D and 1E) or non-motile aplanospore (Figure 1F). While no sexual reproduction has been observed, it is hypothesized that the vegetative cyst may also produce gametes (Freudenthal, 1962). The motile zoospore (gymnodinoid cell) seen in culture (Figure 1D and 1E) has few food reserves and chloroplasts (compared to the vegetative cyst), but does contain longitudinal and transverse flagella for movement (Freudenthal, 1962). The motile zoospore becomes the dominant life cycle stage at dawn, reaching a maximum at midday (Yacobovitch et al., 2004). At the end of this diel ecdysis cycle, they become sessile and settle. These zoospores were not observed reproducing sexually (Freudenthal, 1962). The vegetative cyst may also divide to form 2 daughter vegetative cells or autospores (Figure 1G). Upon emerging from the cyst stage, the daughters are normal independent vegetative cells (Freudenthal, 1962). While life cycle studies have not identified gametogenesis or gametes (Freudenthal, 1962; Schoenberg and Trench, 1980b), information gathered using molecular techniques has indicated that Symbiodinium is haploid in the vegetative life stage, both in culture and in the host (Santos and Coffroth, 2003). At many loci, Symbiodinium had only one allele indicating haploidy in the group. Additionally, the data differed from what would be expected under Hardy-Weinberg equilibrium, further indicating that Symbiodinium are haploid (Santos and 7 Coffroth, 2003). While it appears that during most of its life cycle Symbiodinium is haploid, there is evidence for sexual reproduction of some kind. Allozyme (isoenzyme), RAPD, and DNA fingerprinting analyses show high levels of variation in both cultured and field sampled populations (Schoenberg and Trench, 1980a; Goulet and Coffroth, 1997) that suggests sexual recombination (Baillie et al., 1998; Baillie et al., 2000a; LaJeunesse, 2001). Further evidence examining isoenzyme similarity and phylogenetic similarity found that the two are not correlated, providing evidence for recombination of alleles by sexual reproduction of some kind (Tibayrenc et al., 1991; Baillie et al., 2000b; LaJeunesse, 2001). Taxonomically, the species is difficult to place. It possesses a gymnodinoid form, which may place it in the genus Gymnodinium, if both morphology and life cycle are considered, the species does not fit in any described dinoflagellate genus. As stated above, novel genus Symbiodinium was created by Freudenthal (1962) and assumed a pandemic type species Symbiodinium microadriaticum Freudenthal (Taylor, 1974). However, symbionts from different hosts differed morphologically (Trench, 1979) and in specificity with their host (Kinzie et al., 1977; Kinzie and Chee, 1979). In 1980, further work by Schoenberg and Trench examined isoenzymes, soluble protein patterns, morphology, specificity, and infectivity to examine variation amongst Symbiodinium (1980a; 1980b; 1980c). The symbionts used in these studies were isolated from a variety of hosts including Anthozoans (including Actinaria, Alcyonaria, Hexactiniida, Scleractinia, and Zoanthidea), Scyphozoans, and one molluscan sample (Bivalvia). The isoenzyme data arranged the 17 isolates into 12 ?strains?. Soluble protein patterns mostly agreed with isoenzyme ?strain? identity, though soluble protein suggested greater difference between ?strains? (Schoenberg and Trench, 1980a). 8 Morphological studies using light microscopy revealed that different ?strains? had different average diameters (Schoenberg and Trench, 1980a). Aposymbiotic Aiptasia tagetes were reinfected with the ?strains? studied here and diameter of each specific Symbiodinium ?strain? was maintained, suggesting size is a function of Symbiodinium ?strain? and not of host (Schoenberg and Trench, 1980b). Different outer membranes were seen in situ using the electron microscope including a layer of membranous vesicles associated with the plasma membrane, the inner pellicle membrane, the pellicle (later referred to as the cell wall), and an outer pellicle membrane (Schoenberg and Trench, 1980b). ?Strains? used to successfully infect aposymbiotic Aiptasia tagetes that were similar in isoenzyme and soluble protein analysis to the original A. tagetes symbiont formed dense stable relationships more quickly (Schoenberg and Trench, 1980c) suggesting specificity in the host- symbiont relationship. The hypothesis was put forth that specificity was dependent on recognition during infection (Schoenberg and Trench, 1980c). Chromosome number and volume of symbionts from different hosts also found marked differences between ?strains? and were not due to differences in ploidy (Blank and Trench, 1985). These results support the idea that there are multiple genetically distinct species, within the genus Symbiodinium. Molecular investigations of the Symbiodinium small ribosomal subunit RNA from symbiotic, non-cultured algae identified variation in restriction enzyme polymorphisms. From 16 cnidarian hosts, 6 different ssRNAs were sequenced (Rowan and Powers, 1991b). These preliminary studies also showed that some hosts contained only 1 type while other hosts showed a mixture of symbiont types (Rowan and Powers, 1991b). Again using small ribosomal subunit RNA, high levels of diversity were found within the previously supposed pandemic group called 9 Symbiodinium that were comparable to the diversity among different orders of non-symbiotic dinoflagellates (Rowan and Powers, 1992; LaJeunesse, 2001; Coffroth and Santos, 2005). Subsequent investigations (McNally et al., 1994; LaJeunesse, 2001) into Symbiodinium diversity revealed similar high levels of difference. Examination of a variety of different ?strains? (from Schoenberg and Trench, 1980a-c and Blank and Trench, 1985) showed that these symbiotic dinoflagellates grouped with the genus Gymnodinium (McNally et al., 1994). Figure 2. Phylogenetic relationships of zooxanthellae from different host species taken from Rowan and Powers (1991a). Genetic distances were found using ssRNA sequences, groups A, B and C correspond to the RFLPs (group C includes RFLPs D1 and D2). 10 Interestingly, some strains from distant locations appear closely related (e.g. Indo-Pacific strain from the bivalve Corculum cardissa and Caribbean strain from the zoanthid Zoanthus sociatus), while other geographically close ?strains? appear genetically distant (e.g. Pacific strain from the bivalve C. cardissa and Hawaiian strain from the anemone Aiptasia pulchella) (McNally et al., 1994). Another study using the internal transcribed spacer (ITS) regions of 47 Symbiodinium cultures (some novel cultures and some previously described by Freudenthal, 1962; Trench and Blank, 1987; Trench and Thinh, 1995) identified 17 distinct ?types? or ?strains? (LaJeunesse, 2001). Some of the ?strains? cultured from distant locations appear related, indicating a global distribution of these ?strains? of Symbiodinium (LaJeunesse, 2001). To distinguish ?strains?, it was necessary to develop a reliable method. Initially, ?strains? were placed in 3 clades A, B, and C (Figure 2; Rowan and Powers, 1991a) identified by differences in restriction fragment length polymorphism (RFLP) patterns and small ribosomal subunit RNA sequences (Rowan and Powers, 1991a; Carlos et al., 1999). Subsequent work has revealed 9 recognized clades with multiple ?strains? in each clade (Rowan and Powers, 1991b; Carlos et al., 1999; LaJeunesse and Trench, 2000; LaJeunesse, 2001; Pochon et al., 2001; Pochon et al., 2004; Stat et al., 2006; Pochon and Gates, 2010). Chloroplast and mitochondrial genomes have also been probed to elucidate differences between clades. The first Symbiodinium phylogeny to use an organellar genome used chloroplast large subunit (23S)-rDNA sequences (Santos et al., 2002). In sequences from 35 dinoflagellate cultures (from a variety of hosts and locations) the topology of the cp 23S-rDNA Domain V phylogeny was not statistically different from nuclear rDNA topologies and supported the major Symbiodinium clades (Santos et al., 2002). Domain V of the cp23S-rDNA evolves 9-30 times faster than V1-V4 regions of nuclear small subunit (18S) rDNA, 1-7 times faster than D1-D3 11 regions of nuclear large subunit (28S) rDNA, and 0.27-2.25 times faster than the ITS-rDNA (Santos et al., 2002) and could be an extremely useful sequence in exploring differences between closely related Symbiodinium (Santos et al., 2002). Use of the mitochondrial genome of Symbiodinium and the cytochrome oxidase subunit I (cox1) (Takabayashi et al., 2004) supported the division between clade A and clades B, C, D, E, and F. Relationships within clades B, C, D, E, and F however, were not as well resolved using cox1 as they are when using nrDNA or cp23S-rDNA (Takabayashi et al., 2004). Relationships between the major clades are congruent with those determined from nrDNA and cp23S-rDNA; the relationships within the major clades are robust therefore, any further studies are likely to find variation within these clades, but the major clade divisions should be upheld (Takabayashi et al., 2004). Using the chloroplast genome for future studies may allow for fine genetic differences to be understood since the chloroplast genome has higher variability than the nuclear or mitochondrial genomes (Barbrook et al., 2005). The chloroplast psbA gene produced similar trees to the n28S-rDNA. Additionally, the psbA minicircle and either a longer fragment or other minicircles may provide sufficient variation to determine relationships within phylotypes (Barbrook et al., 2005). The diversity in what was once considered a single, pandemic species is still being examined using a variety of DNA sequences (Santos and Coffroth, 2003; Santos and Coffroth, 2005; Barbrooik et al., 2006; Hunter et al., 2007; Thornhill et al., 2007; Pochon and Gates, 2010). 12 HOSTS While most hosts of Symbiodinium sp. are cnidarians, there are also hosts in other invertebrate groups. Various radiolarians (Phylum Radiolaria), foraminiferans (Phylum Foraminifera), sponges (Phylum Porifera), nudibranchs (Phylum Mollusca, Class Gastropoda), clams (Phylum Mollusca, Class Bivalvia), and a few Acoelomorph species (Phylum Acoelomorpha) are the major examples of such associations (Kempf, 1984; Hill, 1996; Hoegh- Guldberg and Hinde, 1986a; Carlos et al., 2000; Gast and Caron 2001; Ruppert et al., 2004; Barneah et al., 2007a). PROTISTA. Foraminifera are amoeboid protists with pseudopods and a calcium carbonate shell (Hallock et al., 2003). They contribute to the primary production of the oceans where they are able to increase in number due to the carbon fixation of their algal symbionts. Since these are single-celled organisms, the symbionts reside intracellularly. Small subunit ribosomal DNA was used to determine foraminifera symbiont diversity which included dinoflagellates (including Symbiodinium sp.), prasinophytes, and prymnesiophytes (Gast and Caron, 1996; Pawlowski et al., 2001). Radiolarians are single-celled amoeboid protists that produce a mineral skeleton. They have a rapid turn-over rate and are found throughout the oceans (sometimes to such an extent that their skeletal remains cover vast portions of the ocean floor) (Gast and Caron, 2001). Radiolarians have a wide variety of siliceous skeleton size and structure. Like foraminiferans, they associate with a variety of symbionts including dinoflagellates, prasinophytes, and prymnesiophytes. Radiolarians have also been shown to include Symbiondinium sp. amongst their endosymbiotic assemblage (Gast and Caron, 2001). 13 PORIFERA. Sponges may maintain symbionts either inter- or intra-cellularly (Taylor et al., 2007). Like cnidarians, sponges use the photosynthetically fixed carbon compounds from the algae for metabolic processes (Carlos et al., 1999). A boring sponge, Anthosigmella varians, has been shown to have enhanced growth and boring ability that is correlated with higher symbiont concentrations. This indicates that the sponge uses the photosynthetically fixed carbon from Symbiodinium for energy (Hill, 1996). PLATYHELMINTHES. Acoelomorphs are small flatworms lacking an epithelium lined gut that resemble coelenterate planula larvae. A few acoelomorph symbiotic associations have been well-studied in the past including a prasinophyte, a diatom, and a dinoflagellate (Amphidinium) (Barneah et al., 2007a). The symbiosis of these basal bilaterians with Symbiodinium sp. has only begun to be examined (Barneah et al., 2007a; Ogunlana et al., 2005). Current research identifying Symbiodinium sp. as a symbiont of acoelomorphs shows a high degree of specificity with Waminoa sp. hosting a single strain of Symbiodinium (Barneah et al., 2007a). Further studies with Waminoa brickneri have shown maternal transmission of the algal endosymbionts at the oocyte stage (Barneah et al., 2007b). MOLLUSCA. Symbiotic nudibranch species possess symbionts within host-derived ?carrier? cells in or associated with the digestive gland (Kempf, 1984). Nudibranch symbioses range from simple temporary to complex long-term associations (Kempf, 1984; Hoegh-Guldberg and Hinde, 1986a). A simple, temporary symbiosis begins with feeding and lasts only a few days before symbionts are expelled in fecal matter (Kempf, 1991). In such associations, constant feeding on a symbiotic prey maintains a continuous presence of symbionts in the host tissues. A complex long-term relationship includes host morphology that supports the association (e.g. increased cerrata surface area or number, positioning symbionts near irradiance, and specialized 14 internal structure to maintain the symbionts) and allows the host to survive periods of starvation (Kempf, 1984; Hoegh-Guldberg and Hinde, 1986b). Nudibranchs derive nutrition from symbionts and survive longer in the light than dark while starved (Kempf, 1984; Hoegh- Guldberg and Hinde, 1986b). Nudibranch hosts may have large lipid deposits in the symbiont containing cells, suggesting nutrient transfer from Symbiodinium to the host (Kempf, 1984; Rudman, 1991). Many nudibranchs acquire symbionts when they feed and feed only on certain prey species (housing specific Symbiodinium strains), resulting in specific associations between the nudibranch and Symbiodinium (Rudman, 1991; Rudman and Berguist, 2007). In some cases, the nudibranch host does not feed on Symbiodinium containing prey so any specificity is due to the Symbiodinium (Kempf, 1984). Unlike symbioses between Symbiodinium and coelenterate hosts, where some hosts associate almost exclusively with one clade, most bivalve hosts associate with more than one symbiont, sometimes simultaneously. In a study examining 6 clam species (Tridacna gigas, T. squamosa, T. crocea, Hippopus hippopus, H. porcellanus, and Corculum cardissa), each associated with at least 4 Symbiodinium strains (Carlos et al., 2000). Clam symbionts vary in location dependent on host species and can be located in the haemal sinuses of the siphon (T. gigas; Carlos et al., 2000), the gonads, or the digestive gland (Strombus tricornis; Berner et al., 1986; Garcia-Ramos and Banaszak, 2007). However, unlike most other bivalve hosts, T. gigas also maintains algal symbionts extracellularly near the gape. The symbionts may also be defecated when the population becomes too dense for the host (Trench et al., 1981). CNIDARIA. The phylum cnidaria is radially symmetric and has an oral-aboral axis (Ruppert et al., 2004). The basic body plan in the phylum Cnidaria includes two body forms, polyp and medusa. In general, these forms are composed of a gastrovascular cavity opening to 15 the external environment via a mouth surrounded by tentacles (Ruppert et al., 2004). The polyp consists of a stalk attached to substrate at the pedal disc and an oral disc opposite. The oral disc is effectively the feeding apparatus and has tentacles projecting from the edge. These forms are generally sessile, benthic organisms and are the prominent and/or only life stage in the Anthozoa and the Staurozoa (Ruppert et al., 2004). The medusa, on the other hand, is shaped like a bell with the exumbrella (outer non-oral side of the umbrella disk) forming the aboral surface. The oral disc lies opposite the aboral surface in the subumbrella with tentacles surrounding it. Medusa are generally buoyant and have a thick, gelatinous connective tissue (mesoglea) separating the epidermis and gastrodermis. The medusa is the dominant form in the Hydrozoa, Scyphozoa, and Cubozoa (Ruppert et al., 2004). The body wall of cnidarians is composed of two tissue layers, an epidermis and gastrodermis with a layer of extracellular material, the mesoglea (and occasionally wandering cells sometimes with a tissue-like organization), between the two cellular layers (Ruppert et al., 2004). Both the epidermis and gastrodermis contain specialized cells (e.g., cnidocytes, muscle, nerve, glandular, and ciliated cells; Schmid et al., 1999; Ruppert et al., 2004). Cnidarians include a cell type specific to the phyla called a cnidocyte, a specialized cell the animal uses to sting and immobilize its prey (Ruppert et al., 2004). As such, cnidarians are general, filter- feeding carnivores that catch and disable prey using their cnidocytes. Cnidocytes discharge their nematocysts when stimulated by a mechanical or chemical stimulus that indicates a prey organism is nearby (Holstein and Tardent, 1984; Watson and Hessinger, 1989). SYMBIOTIC CNIDARIANS. The major classes with symbiotic members are Anthozoa, Scyphozoa, and Hydrozoa (Ruppert et al., 2004). The host in which Symbiodinium was initially described was Cassiopea xamachana, an upside-down jelly in the class Scyphozoa (Freudenthal, 16 1962; Fitt and Trench, 1983). The symbionts are phagocytosed by the gastrodermal cells similar to the manner food is endocytosed. Lysosomes do not fuse with a symbiont containing vacuole, possibly because the symbiont prevents this fusion (Fitt and Trench, 1983). Corals that house Symbiodinium generally live in warm, nutrient-poor waters and form reefs, anemones often live in similar habitats, but do not secrete a calcium carbonate skeleton (Davy and Cook, 2001). Corals receive much of the carbon used in respiration from their Symbiodinium assemblage (Verde and McCloskey, 1998). The symbiont fixes carbon and donates photosynthetically fixed carbon to the host, this carbon supports the metabolism and reproduction of the host and aids in building the calcium carbonate skeleton (Verde and McCloskey, 1998). Cnidarian hosts donate their waste products to Symbiodinium (such as CO2 and inorganic nutrients- Muscatine, 1990; Yellowlees et al., 2008). As a result of this nutritional exchange, symbiotic corals are especially good reef builders due to the carbon transferred from the symbionts (Lang et al., 1992). NUTRITIONAL ASPECTS OF THE SYMBIOSIS The cnidarian-Symbiodinium symbiosis is based on nutritional transfer of photosynthetically fixed carbon from symbiont to host. Some studies have shown that the transfer of photosynthetically fixed carbon can provide up to 100% of the hosts metabolic carbon needs in a healthy coral (Muscatine et al., 1981). Studies have examined symbioses to determine what carbon compound and how much photosynthetically fixed carbon is transferred. While there is variation between associations when using the same method, by examining many hosts the average amount of photosynthetic carbon and the most common molecules transferred in the cnidarian-Symbiodinium symbiosis can be estimated. 17 Within hosts in the genus Cassiopea, Symbiodinium provides 5-10% of net photosynthate to the host (C. andromedea; Hofmann and Kremer, 1981) providing approximately 169% of the hosts metabolic demand (C. xamachana; Verde and McCloskey, 1998). An early study using 14C found that 45-50% of photosynthetically fixed carbon was transferred to Anthopleura elegantissima while only 20 to 25% was transferred in Palythoa townsleyi (Trench, 1971a). Another method for determining the amount of photosynthetic carbon used by the host is the CZAR (contribution of translocated Symbiodinium carbon to animal daily respiratory requirements) defined by the equation CZAR=(Pz * T)/Ra where Pz is the gross amount of C fixed by the Symbiodinium, T is the percent that is translocated to the host, and Ra is the amount of C respired by the animal (Muscatine and McClosky, 1981). Two experiments using this methodology with Pocillopora damicornis and Fungia scutaria found that 63% and 70% of metabolic needs, respectively, were supplied by carbon from the symbiont (Muscatine and McClosky, 1981). The CZAR of Stylophora pistillata in light was 143%, while in shade it was only 58% (Muscatine et al., 1984). Transfer to Anemonia sulcata under normal conditions provided 110% of the needs of the host (Stambler and Dubinsky, 1987); providing sufficient energy for host respiration and some additional energy for growth or reproduction. Further studies on the transfer of photosynthetically fixed carbon using 13C found high levels of transfer. In Acropora pulchra transfer of >86% of the total photosynthetically fixed carbon to host tissues was seen over a 20 day period (Tanaka et al., 2006). This is in agreement with some earlier CZAR studies that suggested a supply of 100% of the host?s needs (Muscatine et al., 1984; Stambler and Dubinsky, 1987). The percent of photosynthetically fixed carbon translocated varies greatly amongst host species (the usual average reported is 25%-60%; Trench, 1993) making it almost impossible to assign an average translocation percentage across 18 species. Most researchers that attempt to collect and analyze this data report a range of efficiencies or focus on the diversity of translocation efficiency seen in the Symbiodinium/invertebrate symbioses (Davy and Cook, 2001; Grottoli et al., 2006; Tanaka et al., 2006; Loram et al., 2007; Gordan and Leggat, 2010). Comparing fed to starved anemones found the same rate of translocation of photosynthetically fixed carbon (16%) regardless of host nutrition status in Aiptasia pallida (Davy and Cook, 2001). Another variable is the type of symbiont isolate (clade/strain/species) within the host. Determining the impact of different Symbiodinium isolates on translocation rates in the giant sea anemone, Condylactis gigantea, found that both clade A and clade B symbionts released between 30-40% of their fixed carbon to the animal tissues; however, hosts with clade A symbionts showed higher incorporation into the lipid and amino acids fractions than hosts with clade B algae (Loram et al., 2007). In Acropora millepora, symbionts of type C1 transferred more carbon than hosts with symbionts of clade D (Cantin et al., 2009). Determining which carbon compound is translocated to the host involves determining what compounds the alga secretes containing photosynthetically fixed carbon. The symbionts of both corals and Tridacna crocea produced primarily glycerol in the presence of host tissue (Muscatine, 1967). While incubated with 14C in culture, glycerol again was the major product with alanine, glucose, fumaric acid, succinic acid, and glycolic acid released in smaller quantities (Trench, 1971b). These compounds were secreted in increased quantities in the presence of host homogenates (Trench, 1971c). In Anemonia viridis, 14C labeled glucose and succinate/fumarate given to the host produced the same signature as anemones incubated with NaH14CO2 in the light suggesting that glucose and succinate/fumarate are transferred from Symbiodinium to A. viridis (Whitehead and Douglas, 2003). 19 ULTRASTRUCTURAL ASPECTS OF THE SYMBIOSIS Early ultrastructural description of the endosymbiont from Anemonia sulcata (described as being ?akin? to Symbiodinium microadriaticum but likely another isolate from the genus Symbiodinium) described the thick cellulosic wall surrounding a single lobed chloroplast with a single stalked pyrenoid. Cells also contained a large vacuole termed an accumulation body because it appears to accumulate waste materials (Taylor, 1968). The nucleus is approximately spherical, large and the chromosomes have the typical dinoflagellate appearance (not undergoing decondensation even during resting stages; Dodge, 1966). Further ultrastructural studies have described Symbiodinium from several other hosts including Cassiopeia sp., Condylactis sp. (Kevin et al., 1969), Tridacnidae (Taylor, 1969), Ragactis lucida, Montipora verrucosa, Zoanthus sociatus (Trench and Blank, 1987), Aiptasia pallida, and Phyllactis flosculifera (Wakefield et al., 2000). The anthozoan hosts house their symbionts within symbiosome membranes in the gastrodermal cells (Wakefield and Kempf, 2001). The symbiosome membrane consists of the host membrane that covers the symbiont in situ derived from the original phagocytic vacuole from infection. Beneath this outer layer, there are many other membranous layers observed within the host that may result from a repeated ecdysis cycle within the host symbiosome membrane (Wakefield et al., 2000; Wakefield and Kempf, 2001). INITIATION OF THE CNIDARIAN-SYMBIODINIUM SYMBIOSIS The Symbiodinium-cnidarian symbiosis can be initiated in two ways. Vertical transmission of the symbiont can occur directly from the parent, with Symbiodinium included in the forming oocyte or larva by the parental polyp (Fautin, 2002). Horizontal transmission can occur once the larva is released from the parent and is either a planktotrophic larva or a settled 20 polyp. In horizontal transmission, larvae acquire symbionts from the environment, usually during feeding (Fautin, 2002). In experimentally manipulated Cassiopeia xamachana, during vertical transmission the host experiences higher levels of asexual reproduction (and therefore high host fitness) indicating the symbionts are mutualistic; whereas, during horizontal transmission the host experiences lower levels of asexual reproduction and thus lower fitness. This indicates that the horizontally transmited Symbiodinium strains infecting the host initially behave more like parasites (Sachs and Wilcox, 2006). In most species (including C. xamachana which uses vertical transmission in nature) one mode of transmission dominates or is the only possibility. In these species (ex. brooding corals that undergo vertical transmission; Magalon et al. 2007) there is little opportunity to determine how a change of transmission mode would affect the mutualistic or parasitic nature of the symbiosis. VERTICAL TRANSMISSION. Vertical transmission occurs in several ways. Maternal tissues can transfer symbionts to the egg before fertilization (Davy and Turner, 2003; Barneah et al., 2007b), the parent can brood larvae internally and transfer symbionts to the larvae before dispersal (Benayahu et al., 1988), or anything in between. While the exact process of the transfer may not be known in individual cases, in many of the brooding corals vertical transmission is assumed based on genetic identification of symbionts from a variety of individuals (Thornhill et al., 2006; Magalon et al., 2007). Regardless of the specific mechanism behind a particular vertical transmission event, the type of symbiont present in the offspring is the same as in the parent. Transmission in Anthopleura ballii occurs early in development when the ova become infected with Symbiodinium just prior to spawning. The ova are then fertilized and undergo 21 cleavage (Davy and Turner, 2003). Development continues normally with the symbionts sequestered to one side of the embryo (Davy and Turner, 2003). This species may have adopted this form of vertical transmission that occurs early in development because it generally thrives in areas where Symbiodinium are scarce (the temperate zone) (Davy and Turner, 2003). This is an excellent example of the early initiation of vertical transmission being intimately tied to an organism?s development. Other examples of vertical transfer via ova containing Symbiodinium sp. are found in Porites cylindrica, Montipora digitata, Pocillopora verrucosa (Hirose and Hidaka, 2006), and P. damicornis (Harii et al., 2002). In the case of the Pocillopora species, symbionts are sequestered in one part of the ovum, embryo, and the larva. The blastomeres containing Symbiodinium become endodermal tissues of the host (Hirose et al., 2000). However, in Porites cylindrica and Montipora digitata the symbionts are equally distributed throughout the cytoplasm of the oocyte and during early cleavage only becoming sequestered to the endoderm during gastrulation when blastomeres containing Symbiodinium drop into the blastocoel and develop into the gastrodermis (Hirose and Hidaka, 2006). In all these cases the symbionts must become part of the endodermal tissues at some point in development (Hirose and Hidaka, 2006). In Euphyllia glabrescens, Symbiodinium are distributed evenly throughout the oocyte, and later throughout the epidermis and gastrodermis (Huang et al., 2008). Following planulation, symbionts are redistributed to the gastrodermis and by metamorphosis the symbionts are found only in the gastrodermis (Huang et al., 2008). This indicates that Symbiodinium infection may interact with host development. HORIZONTAL TRANSMISSION. Horizontal transmission can happen a variety of different ways contingent on the host life stage when infection occurs. Horizontal infection is 22 considered open since offspring can potentially obtain any Symbiodinium type present in the water column. This is seen commonly in broadcast spawning scleractinian corals (van Oppen, 2004). There are a few different proposed mechanisms by which horizontal transmission can result in a specific host-symbiont association. These include 1) preferential feeding seen in the planulae of Aiptasia tagetes and Anthopleura elegantissima on material that includes conspecific components (such as symbionts and associated host membranous material expelled by other hosts; Riggs et al., 1988; Schwarz et al., 2002) 2) chemical attraction of the motile symbiont to the host (Kinzie, 1974; Pasternak et al., 2006), 3) phototaxis of the motile symbiont to the host (host pre-symbotic larvae of Fungia scutaria show an increased production of light from luminescent green fluorescent protein (GFP) which symbionts swarm toward; Hollingsworth et al., 2004; Hollingsworth et al., 2005) and 4) uptake of a number of Symbiodinium types followed by selection of only one or a few isolates (Dunn and Weis, 2009). Depending on the host- symbiont association, interactions between some or all of these mechanisms could result in a stable association between host and symbiont. In typical horizontal transmission Symbiodinium is phagocytosed into the endodermal cells during feeding. One study in Aiptasia tagetes fed the planula larvae one of four diets; crab, Artemia flakes, Symbiodinium, the unicellular alga Isochrysis, or a combination of these (Riggs et al., 1988). All the food items were eaten and throughout the study (35 days), the planulae retained symbionts that were ingested as part of their diet. While this study did not examine phagocytosis of symbionts or preference in food choice, it did indicate that planula could and did ingest symbionts and maintain them during the study. In Heteroxenia fuscencens, aposymbiotic planula larvae are released from the parent, settle rapidly (within 7-8 hours), and within about 3 days are symbiotic (Benayahu et al., 1989). 23 Symbiodinium are acquired during metamorphosis, along with the appearance of tentacles, and are initially near the mouth. This is consistent with the host becoming symbiotic via feeding (Benayahu et al., 1989). Fungia scutaria also releases aposymbiotic larva that acquire symbionts during feeding. When trying to reinfect these larvae with Symbiodinium from Fungia scutaria, Montipora verrucosa, Porites compressa, and Pocillopora damicornis, the homologous Symbiodinium strain (from the parent species) infected the aposymbiotic larvae more easily (Weis et al., 2001), indicating specificity between host and symbiont. Aposymbiotic Anthopleura elegantissima obtain symbionts while feeding as a planula larva. During feeding trials, planulae of Anthopleura elegantissima did not feed when presented with just Symbiodinium (Schwarz et al., 2002). However, if macerated animal tissue (of various species) was added, a feeding response was elicited and larvae ingested any available particulate matter. After ingestion, animal tissue is digested, but symbionts are endocytosed and remain undigested while held inside vesicles in the endodermal cells (Schwarz et al., 2002). Feeding on material that contains macerated animal tissue is not unique and has also been documented in Aiptasia tagetes (Riggs et al., 1988), and Fungia scutaria (Weis et al., 2001). One hypothesis is that the initiation of feeding in the presence of macerated host tissue could show a preference for symbiont containing material (Schwarz et al., 2002), supporting the theory of specificity early in uptake for food likely to contain a preferred symbiont isolate. This may be the case since, in nature, when symbiont populations within a host get too large the host will expel some material containing living Symbiodinium and host membrane remnants (Steele, 1977; Sachs and Wilcox, 2006). Another method for obtaining a specific symbiont-host relationship could rely on symbionts seeking out the appropriate host, possibly through chemical cues (Kinzie, 1974; 24 Pasternak et al., 2006). Aposymbiotic planulae of Pseudopterogorgia bipinnata observed during experimental infection had symbionts swimming either towards the mouth or near the polyp suggesting the host is somehow attracting symbionts (Kinzie, 1974). In a study with motile Symbiodinium isolated from Cassiopeia xamachana, taxis assays to various host compounds showed attraction to compounds containing nitrogen (ammonium, nitrate, urea, and amino acids), while no response was observed to phosphate, sulfate, sugars, glycerol, or vitamins (Fitt, 1985). High concentrations of ammonia didn?t attract Symbiodinium, but more dilute concentrations led to some attraction. This suggests that symbionts are attracted to concentrations of nitrogen-containing compounds excreted by hosts under natural conditions (Fitt, 1985). There is no data to suggest that attraction to nitrogen compounds may convey specificity on the system (e.g., a host attracts only the alga of its preferred association) and it seems much more likely that this is a general response to nitrogen-containing compounds which all hosts excrete as waste. In the soft coral Heteroxenia fuscescens, Symbiodinium swimming became slower and directed to the source of host attractants (in this case, attractants were chemical substances associated with the juvenile polyp, but not with the adult; Pasternak et al., 2004; Pasternak et al., 2006). Further studies investigated the potential for Symbiodinium phototaxis. Initially, researchers noted an upregulation of green fluorescent protein (GFP) in pre-symbiotic larvae of the mushroom coral (Fungia scutaria) that coincided with larval ability to accumulate Symbiodinium (3-5 days after fertilization) (Hollingsworth et al., 2004; Hollingsworth et al., 2005). Further investigation revealed Symbiodinium swarmed toward the light source under green light, but under UV and blue light the symbiont swam away from the light source (Hollingsworth et al., 2005). The host secretion of more GFP in pre-symbiotic hosts combined 25 with increased phototaxis of the symbiont towards green light suggests that Symbiodinium may swim toward these pre-symbiotic planulae larvae. Chemotaxis and phototaxis to host attractants (chemical and light) suggest the symbiont is actively seeking a host. However, chemotaxis and phototaxis probably are not responsible for specificity as many types of Symbiodinium would likely be attracted to green light and nitrogenous compounds. Further research into the next steps in infection that result in a specific association are necessary. SPECIFICITY Since vertical transmission results in the offspring receiving the same symbiont isolate as the parent, these associations should be highly specific. In horizontal transmission, the host obtains symbionts from the environment. The variety of symbionts present in the water column may allow for variation during the uptake process and in the association. The two transmission methods could confer different levels of variability or specificity. The current evidence is far from conclusive, but seems to suggest high levels of specificity (in most cases) regardless of transmission type (Benayahu et al., 1988; Davy and Turner, 2003; van Oppen, 2004; Barneah et al., 2007b). Examinations of Acroporid corals in Indonesia and the central Great Barrier Reef found that diversity of Symbiodinium in Montipora species (vertical transmitters) and Acropora species (horizontal transmitters) was not significantly different. Both genera had symbionts mostly from one clade (about ? of hosts within each genus hosted symbionts from one clade). Though the two coral genera hosted some symbionts from the same clade (C), these were relatively divergent from each other (5% sequence divergence) (van Oppen, 2004) suggesting that mode of uptake (either horizontal or vertical) does not affect specificity of the association (van Oppen, 2004). 26 Investigations on Red Sea soft coral symbionts found that hosts practicing vertical transmission associated exclusively with Clade A, while horizontal transmitting hosts contained Clade C (Barneah et al., 2004). Therefore, in the Red Sea, hosts with horizontal uptake developed specificity for Clade C symbionts; while Clade A symbionts co-evolved with vertical transmitting hosts (Barneah et al., 2004). Due to high genetic variability amongst Clade C symbionts it is likely that in the Red Sea some Clade C symbionts have a specific physiological adaptation to their host environment, resulting in highly specific associations (Barneah et al., 2004). Any contradictions between these two studies could indicate that specificity related to transmission type varies between different hosts or geographic regions. This early evidence for specificity between host and symbiont may call into question the adaptive bleaching hypothesis. The adaptive bleaching hypothesis states that bleaching may allow hosts to change Symbiodinium strains to a strain more suitable to the extant environmental conditions (and less susceptible to future bleaching). In 2006, Goulet examined 442 coral species from 43 studies to determine whether the hosts could associate with multiple symbiont types. Presumably, the ability to change symbiont strain is linked to the ability for a host to associate with multiple strains, either at different depths, locations or microclimates within the host (Goulet, 2006). Using these criteria, only 23% of coral species associate with multiple clades and could change symbiont suggesting that most corals will not change symbionts under environmental stress and only hosts that already maintain symbionts not susceptible to high temperature bleaching would survive (Goulet, 2006). A response to the Goulet review (2006) re-evaluated the data and found that 64% of the corals used in the study (198 out of 307 species; excluding species noted only to the genus level) were sampled 5 times or less. Scleractinian species sampled at least 10 times did associate with 27 multiple clades; while scleractinians sampled fewer times did not (Baker and Romanski, 2007). The octocorals were more specific to one clade than scleractinians; almost all of the scleractinian families associated with multiple clades (only the Rhizangiidae associated with only 1 clade) (Baker and Romanski, 2007). Many of the scleractinian corals examined are actually a dynamic community with the ability to associate with multiple symbionts (Baker and Romanski, 2007). Thus in many of the species Goulet (2006) considered, she may have missed the potential for association with multiple clades due to insufficient sampling. However, the original data set (Goulet, 2006) found that in 87.5% of cases diversity in symbiont hosting was seen with less than 5 samples (Goulet, 2007). It is still unclear whether this data supports a lack of flexibility (Goulet, 2006; Goulet, 2007; Goulet et al., 2008) or the ability for many scleractinians to change symbionts (Baker and Romanski, 2007). Both scleractinian corals and octocorals are generally horizontal transmitters (Schwarz et al., 1999; Koike et al., 2004) and horizontal transmission dominates (about 85% of species) across all cnidarians (Koike et al., 2004). The variation in association with multiple strains between the octocorals (most associating with 1 Symbiodinium isolate) and scleractinians (many species associating with multiple isolates) is inconsistent with the conclusion that mode of transmission affects the specificity of the relationship. While there appears to be a difference in specificity between octocorals and scleractinians however; there is no difference in transmission mode. MECHANISMS OF SPECIFICITY. Once it was established that there was some level of specificity in these symbioses, investigators wanted to know how the specific relationships are established. Initial studies in the Symbiodinium-cnidarian association investigated what stable associations could be established. Investigations of Symbiodinium infection in A. pallida found 28 that infection with clade B symbionts was more successful (faster and with denser populations) than with clades C and A. This suggested a mechanism for recognition of the symbiont after phagocytosis or different rates of division among clades within the host (Belda-Baillie et al., 2002). For Cassiopeia xamachana, the scyphistomae took up Symbiodinium freshly isolated from several host species; however, Symbiodinium originally extracted from C. xamachana established a denser, stable population twice as fast (14 vs. 27 days) as those from other hosts (Colley and Trench, 1983). Infection in similar associations reveals some possibilities for the mechanisms involved in specificity between Symbiodinium and the cnidarian hosts. Symbiotic chlorellae showed decreased rates of infection in aposymbiotic Paramecium bursaria when the surface of chlorella was coated with antibodies or lectins. This suggests molecules on the surface of chlorellae are involved in recognition during the onset of symbiosis in this system (Reisser et al., 1982). In the Hydra viridis association with chlorellae-like symbionts, reinfection was inhibited with lectin-treated symbionts (Meints and Pardy, 1980). These studies may suggest possible mechanisms for recognition in the Symbiodinium-cnidarian association. Examination of the glycoproteins on Symbiodinium during the initiation of symbiosis with Aiptasia pallida showed that infection decreased when symbionts were treated with lectins altering the surface coat (Lin et al., 2000). Investigation of the macromolecules associated with the cell wall of Symbiodinium found that the cell wall contained cellulose and proteins (Markell et al., 1992). These proteins are hypothesized to be glycoproteins because other microalgae have a high content of glycoprotein associated with the cell wall (Miller et al., 1974; Catt et al., 1976). Subsequent investigation of the macromolecules exuded into culture media identified the amino acid and sugar composition 29 as components of large molecular weight glycoconjugates (Markell and Trench, 1993). Glycoconjugates were released in varying amounts by different Symbiodinium isolates, but all essential amino acids along with neutral sugars (glucose, galactose, fucose, ribose, and mannose) were present in the exudate (Markell and Trench, 1993). A more recent study examined glycoproteins in initiation of this association by using either enzymes to presumably destroy recognized glycoprotein moieties or lectins to mask carbohydrate moieties. These treatments were found to hinder the establishment of the symbiosis (Lin et al., 2000). In addition to studies examining the surface coat, several studies have focused on the molecules in the phagosomes and how they affect retention or digestion of the symbionts. One of these is Rab7, part of the Rab family which is a group of GTPase regulators of vesicular trafficking (Zerial and McBride, 2001). Aiptasia pulchella Rab protein (ApRab7) is homologous to other Rab7 proteins found in the human, mouse, and worm genomes (Chen et al., 2003). Live photosynthetically active symbionts were included in phagosomes without ApRab7, while heat- killed or photosynthetically-impaired symbionts were housed in phagosomes with ApRab7 (Chen et al., 2003). Further investigation showed the Rab5 homolog (ApRab5) was present in early endocytic and phagocytic compartments that promote symbiont survival (Chen et al., 2004). ApRab5 was mostly excluded from phagosomes containing dead algae (Chen et al., 2004). This could suggest that live Symbiodinium retain ApRab5 in the symbiosome membrane so they may persist in the host cell and exclude ApRab7 for the same purpose (Chen et al., 2003; Chen et al., 2004). ApRab11 showed similar patterns to ApRab7 and was included in phagosomes of heat-killed Symbiodinium, photosynthetically inhibited Symbiodinium, or latex beads and actively excluded from phagosomes that contained live Symbiodinium (Chen et al., 30 2005). These proteins could act as one mechanism in the Symbiodinium-cnidarian symbiosis to establish and maintain the association. If multiple symbionts are phagocytosed in horizontal transmitting hosts or after a bleaching event, some of the Symbiodinium isolates will be maintained while others are digested or egested. This process is referred to as ?the winnowing? (Nyholm and McFall-Ngai, 2004). In Fungia scutaria, both homologous and heterologous symbionts are phagocytosed into host cells, but shortly thereafter homologous algae were found to localize between the oral and aboral ends while heterologous algae did not localize in any specific place (Rodriguez-Lanetty et al., 2004; Rodriguez-Lanetty et al., 2006). In Acropora tenuis and A. millepora, juveniles become initially colonized by both heterologous and homologous algae, but by 3.5 years of age the polyps were hosting homologous symbionts (Abrego et al., 2009b). Further investigation in A. tenuis found that ITS-1 type C1 symbionts initially dominated infections, but after 96 h, type D symbionts (the homologous type) began to dominate, suggesting that the winnowing process begins shortly after infection (Bay et al., 2011). On the cellular level, Fungia scutaria exhibited an increase in caspase activation and apoptosis when infected with a heterologous symbiont, suggesting that apoptosis may be involved in the post-phagocytic winnowing (Dunn and Weis, 2009). With the high levels of specificity in the cnidarian-Symbiodinium symbiosis, there are many possible mechanisms that may play a part in establishing the relationship between host and symbiont. Specificity may be due in part to the origin of symbionts (either from parent or from environment; Benayahu et al., 1988; Davy and Turner, 2003; van Oppen, 2004; Barneah et al., 2007b). Recognition may also occur at the symbiosome membrane because of glycoprotein coats secreted by the alga (Weis et al., 2001). Other studies suggest the symbiont may be responsible for maintaining the association by halting host digestion (Chen et al., 2003; Chen et al., 2004; 31 Chen et al., 2005) or that a post-phagocytic winnowing process may establish the stable relationship (Dunn and Weis, 2009). An alternative mechanism, the ?Test Tube Hypothesis?, suggests algae are ingested unselectively to the digestive vacuole and then subjected to a selection process similar to the selection process that would occur in the media in a test tube (Huss et al., 1994). The algae preadapted to live in the host environment will be maintained and reproduce faster and more effectively than other symbionts (Huss et al., 1994). At this point, there is evidence for each of these mechanisms and each or combinations of 2 or more may be important in many Symbiodinium symbioses. BREAKDOWN OF THE SYMBIOSIS Symbiodinium symbioses can break down in a process which leaves the invertebrate host devoid of symbionts. The breakdown of the Symbiodinium-cnidarian symbiosis resulting in the loss of Symbiodinium from the host or loss of pigments from the symbionts is called bleaching. Thus far a number of causative agents have been implicated in bleaching, among these are changes in temperature, ultraviolet radiation, disease, and pollution (Jokiel and Coles, 1977; Glynn, 1991; Hoegh-Guldberg, 1999; Hughes et al., 2003; Hoegh-Guldberg et al., 2007). Due to recent rises in global temperature, the bleaching agents that are best understood and most often seen are those of temperature and ultraviolet radiation (either alone or in concert). Over the last 50 years the frequency and scale of bleaching events has increased rapidly (Hughes et al., 2003). Using this information to create predictive models indicates that reefs in some areas (Micronesia and Western Polynesia in particular) could be extremely vulnerable to catastrophic collapse over the next 30-50 years (Donner et al., 2005). Increases of just 1?C predict an increase in bleaching 32 occurrence involving 50% to 82% of reefs while a more dramatic temperature increase of 2?C would increase the occurrence of bleaching to 97% of all reefs (Berkelmans et al., 2004). The precise cellular causes of bleaching and which partner in the relationship initiates the event are beginning to be understood. The process is a response to stress by the association (Perez et al., 2001; Perez and Weis, 2006; Weis, 2008). The translocation of energetic nutrients from Symbiodinium to the host has been shown to decrease in bleaching situations (high irradiance and temperature; Porter et al., 1989). Furthermore, study at the cellular level has suggested a variety of mechanisms including host cell detachment (Gates et al., 1992), apoptosis, and autophagy of host cells (Hanes and Kempf, 2013). These cellular responses appear to be related to the increase in reactive oxygen species (ROS) produced by damage to the photosynthetic membranes of the symbiont (Tchernov et al., 2004) and to mitochondrial membranes in both the host and symbiont (Weis, 2008) during thermal bleaching. In a healthy, non-stressed host, these reactive oxygen species would be managed by enzymes such as catalase and superoxide dismutase; however, these ameliorative mechanisms are not sufficient during bleaching episodes (Richier et al., 2005; Lesser, 2006; Merle et al., 2007; Weis, 2008). One reactive oxygen species, nitric oxide, has been well studied and is produced as a response to stress in the model anemone, Aiptasia pallida (Perez and Weis, 2006). Nitric oxide may act not just as a cytotoxin but also as a signaling molecule in host innate immunity (Fang, 2004; Pacher et al., 2007). While there is still much to be learned about the cellular processes behind bleaching, it is clear that thermally induced bleaching is a threat to coral reefs. While the cellular bleaching process is not completely understood, there has been research investigating variation in susceptibility to bleaching. Examination of host species reveals that heavier bodied corals have better survival rates than those with more delicate 33 branching structure (Loya et al., 2001; McClanahan, 2004). Additionally, studies examining the thermal tolerance of Symbiodinium in culture found that there is variation among isolates in regard to their response to elevated temperatures (Warner et al., 1996; Iglesias-Prieto and Trench, 1997; Rowan et al., 1997; Kinzie et al., 2001; Perez et al., 2001; Bhagooli and Hidaka, 2004). Often these studies found that clade D Symbiodinium were more tolerant to high temperatures (Baker et al., 2004; Chen et al., 2003; Rowan, 2004) while clade A symbionts were often identified as being more susceptible to damage from high temperatures (Diaz-Almeyda et al., 2011; Steinke et al., 2011). Likewise, in the field clade D dominated sites that routinely received elevated temperatures, indicating that clade D symbionts are better adapted to high temperature (Fabricius et al., 2004). Clade D also dominated reefs that had just undergone a bleaching event indicating either that clade D symbionts did not bleach from their hosts or that clade D symbionts readily re-colonized bleached hosts (Glynn et al., 2001; Jones et al., 2008; Oliver and Palumbi, 2011). Even though thermal stress acts on the holobiont (symbiont, host, and any associated bacteria), it is important to understand how both the host and symbiont impact the thermal tolerance and therefore the bleaching susceptibility of the coral as a whole. There is some evidence that non-fatal bleaching episodes may be an adaptive mechanism to change symbionts to a more robust association (Buddemeier and Fautin, 1993; Baker et al., 2004; Stat and Gates, 2011). A study of 3 genera (Acropora, Pocillopora, and Porites) on the Great Barrier Reef that encompassed the 1998 and 2002 bleaching events found that both Pocillopora and Acropora showed an increase in tolerance compared with what was expected in the 2002 bleaching event. This increase in tolerance may be in part due to selective mortality (of more susceptible individuals) during the 1998 bleaching but also may indicate some capacity for acclimatization and hope for coral reefs (Maynard et al., 2008). Information on the 34 susceptibility and tolerance of hosts and symbionts could be used to practice better reef management (transplanting more robust corals to reefs) which could in turn protect the coral reef ecosystem from collapse. Understanding both the bleaching process and the potential stability of a variety of host/symbiont associations should improve future reef management and appropriate responses to bleaching events. 35 II. IMMUNOHISTOCHEMICAL VARIATION AMONGST SYMBIODINIUM ISOLATES. ABSTRACT Symbiodinium (Freudenthal) reside intracellularly in a complex symbiosome (host and symbiont- derived) within cnidarian hosts in a specific host-symbiont association. Symbiodinium is a diverse genus with variation equivalent to other dinoflagellate orders. In this paper, our investigation into specificity examines antigenic variation in the algal mucilage secretions at the host-symbiont interface. Cultured Symbiodinium from a variety of clades were labeled with one of two antibodies to symbiont mucilage (PC3, developed using a clade B alga cultured from Aiptasia pallida; BF10, developed using a clade F alga cultured from Briareum sp.). The labeling was visualized with a fluorescent marker and examined with epifluorescence and confocal microscopy. PC3 antigen was found in cultured Symbiodinium from clades A and B, but not clades C, D, E and F. The correlation between labeling and clade may account for some of the specificity between host and symbiont in the field. Within clades A and B there was variation in the amount of label present. BF10 antigen was more specific and only found in cultures of the same cp23S-rDNA strain the antibody was created against. These results indicate that the mucilage secretions do vary both qualitatively and quantitatively amongst Symbiodinium strains. Since the mucilage forms the host-symbiont interface, variation in its molecular composition is likely to be the source of signals involved in recognition and specificity. INTRODUCTION Coral reef ecosystems are among the most diverse habitats in the world. Reefs provide benefits that include: ecotourism, fish hatchery and/or nursery for many commercial fisheries, species habitat, and coastal protection (Richmond and Wolanski, 2011). These diverse 36 ecosystems are built upon the relationship between Symbiodinium sp. and their host cnidarians. Hermatypic corals are the structural foundation of the reef in large part due to the photosynthetically fixed carbon compounds donated by Symbiodinium (Muscatine and Porter, 1977; Hallock, 2001). Unfortunately, the coral reef ecosystems are threatened world-wide by symbiont loss (commonly referred to as coral bleaching) that occurs due to increased sea surface temperature in addition to other stress-related factors resulting from human activities (Jokiel and Coles, 1977; Hoegh-Guldberg, 1999). Symbiodinium associate with a variety of hosts as endosymbionts. In symbioses with cnidarians, the symbionts reside intracellularly (Colley and Trench, 1983). The cnidarian/Symbiodinium symbiotic association can be initiated in one of two ways: either by vertical transmission (obtaining symbionts from the parent) or horizontal transmission (obtaining symbionts from the surrounding environment). Symbiodinium are acquired via horizontal transmission in 85% of invertebrate hosts (Schwarz et al., 2002). Association between Symbiodinium and the host cnidarian is based on nutrient exchange. The symbiont produces photosynthate that is transferred to the host and can potentially provide enough carbon based nutrients to meet a host?s energetic demands (Falkowski et al., 1984; Muscatine et al., 1984; Muscatine, 1990). The symbiont benefits from a protected environment and host waste products. These include nitrogen compounds such as ammonium and nitrate that are typically limiting on reefs (Muscatine and Porter, 1977; Muller-Parker and D'Elia, 1997), as well as CO2 (Pearse, 1970). The fact that this symbiosis exists in hermatypic corals allows for the energetic demands of higher calcification rates necessary for reef building (Pearse and Muscatine, 1971; Barnes and Chalker, 1990), as well as better conservation of nitrogen 37 compounds within the reef environment (Lewis and Smith, 1971; Burris, 1983; Ambariyanto and Hoegh-Goldberg, 1996). Due to their nearly identical morphology, symbiotic algae in corals were originally thought to be the single species Symbiodinium microadriaticum (Freudenthal, 1962); however, molecular methods have revealed that the genus actually is a complex of (at least) 9 clades (Rowan and Powers, 1991b; Carlos et al., 1999; LaJeunesse and Trench, 2000; LaJeunesse, 2001; Pochon et al., 2001; Pochon et al., 2004; Pochon and Gates, 2010). The variation within this one genus is equivalent to the diversity amongst other dinoflagellate orders (Rowan and Powers, 1992). Variation in thermal tolerance of Symbiodinium isolates is based on observational studies of hosts in different habitats (Oliver and Palumbi, 2011) or before and after bleaching (Glynn et al., 2001; Baker et al., 2004; Jones et al., 2008; LaJeunesse et al., 2010). Studies manipulating either host habitat in transplantation experiments or temperature regimes of cultured symbionts also indicate that certain symbiont isolates survive more effectively than other strains in thermally stressful environments and are therefore thermally tolerant (Iglesias- Prieto and Trench, 1997; Rowan, 2004; Berkelmans and van Oppen, 2006). In particular, strains A1, A2, D, and F2 are accepted as being thermally tolerant (Rowan, 2004; Baker et al., 2004; Robison and Warner, 2006; Steinke et al., 2011). Previous studies have revealed that the wide variation in both host and symbiont species generates potential for specificity or plasticity within the associations. Previous studies have examined a variety of associations and found varying levels of specificity between partners. Many studies of cnidarians have found host species typically paired with the same symbiont strain (Schoenberg and Trench 1980a, b, c; Rowan and Powers 1991a, b; Goulet and Coffroth, 2003a, b; Santos et al., 2003; Goulet and Coffroth, 2004; Baker et al., 1997). This specificity 38 exists both in host species that display vertical as well as horizontal transmission (van Oppen, 2004). Host species that can associate with more than one symbiont strain tend to show variation in their associated strains between colonies in different habitats (Rowan and Knowlton, 1995; Rowan et al. 1997) or within colonies at different depths or even parts of a given colony exposed to different light regimes (van Oppen et al., 2001; Ulstrup and van Oppen, 2003). Host species that in the field can associate with several symbiont strains may have varied mortality during a bleaching event; hosts containing more tolerant symbionts show reduced mortality compared to hosts with thermally susceptible symbionts (Jones et al., 2008; LaJeunesse et al., 2010). Following a bleaching event, hosts with multiple strains may change to a more thermally tolerant, dominant, symbiont strain (Baker et al., 2004; Berkelmans and van Oppen, 2006). Thus, the association between host and symbiont is often specific, but in some cases also allows for some variation depending on environmental conditions. Within the cells of the host, the symbiont resides within a host derived (Wakefield and Kempf, 2001) vacuole membrane with several layers of additional membranous material called the symbiosome (Roth, 1989). In the association between Symbiodinium and cnidarians, the symbiosome is composed of both host and symbiont components (Wakefield and Kempf, 2001). The symbiont component consists of multiple layers of a ?membranous material? that Wakefield and Kempf (2001) have suggested is composed of shed thecal plates from a reduced ecdysis cycle that occurs in situ. The host component consists of the phagosome membrane that resulted from the host cell endocytosing the symbiont (Colley and Trench, 1983; Fitt and Trench, 1983; Schwarz et al., 1999; Wakefield and Kempf, 2001; Schwarz et al., 2002). This membrane is sometimes referred to as the outer symbiosome membrane (Wakefield and Kempf, 2001). 39 The symbiont secreted layers of presumed shed thecal plates are intermingled with exudates produced by the symbiont (Wakefield and Kempf, 2001). Work by Markell et al. (1992) and Markell and Trench (1993) on cultured symbionts suggest that these exudates are largely composed of glycoproteins. Symbiont-derived glycoproteins that are water soluble in culture are secreted through the cell wall and are thought to form the symbiont component that is in direct contact with the host via the phagosome membrane (Wakefield and Kempf, 2001). The region of contact between host and symbiont exudate is the likely location of any recognition molecules involved in both initial uptake of the symbiont by the host and maintenance of the symbiosis. Lin et al. (2000) used lectins to mask the cell surface and proteolytic enzymes to remove cell surface glycoproteins and saw inhibition of symbiont uptake by the host in both types of treatment. This suggested that the affected glycoconjugates are used for cell-cell recognition in the initiation of this symbiosis (Lin et al., 2000). The process of recognition probably contains multiple steps. Initially, there is selection for living symbionts with an intact outer layer while dead symbionts or those whose outer layer is compromised are discarded (Fitt and Trench, 1983; Lin et al., 2000). However, there is also evidence that in nature the initial uptake of symbionts is dominated by ?highly infectious? symbiont strains (Abrego et al., 2009a) that may later be replaced by that hosts?s preferred symbiont strain. The mucilaginous outer layer is secreted in situ by symbionts already taken up by the host tissue (Wakefield and Kempf, 2001). Wakefield and Kempf?s observation suggests that this layer may be involved in later stages of the recognition process and that secretion of mucilage may be a factor in preventing the host from digesting or expelling the symbiont. Cell-cell recognition processes have been identified in a number of other organisms, from the mammalian immune system (Takeda et al., 2003) to predatory dinoflagellates (Jeong et al., 40 1997; Martel, 2006; Breckels et al., 2011). These processes have a few things in common: a molecule on each cell involved in the recognition and a mechanism for these molecules to come into contact with each other. These molecules can then bind and produce a cascade of events resulting in a variety of different outcomes. In predatory dinoflagellates, recognition results in the dinoflagellate either consuming its preferred prey or leaving the non-prey organism alone (Jeong et al., 1997; Martel, 2006; Breckels et al., 2011). In cnidarian/Symbiodinium associations, recognition (or lack thereof) results either in the symbiont remaining in the host gastrodermal cell or potential digestion and/or expulsion of the non-preferred partner. This study is directed toward an examination of variation in the composition of surface bound exudates of a number of strains of Symbiodinium. Such variation could be involved in recognition specificity and host/symbiont ?communication? in cnidarian host-Symbiodinium symbioses. MATERIALS AND METHODS ALGAL CULTURE. Symbiodinium isolates, representing a range of host species, geographic locations and genetic diversity (Table 1), were cultured in F/2 medium (without silica to prevent the growth of diatoms; Guillard and Ryther, 1962) using a 12/12 hr light/dark regimen at Auburn University. Light intensity was maintained at ~180 ?mol photons /m2/s using Sylvania Cool White 20W fluorescent bulbs. Samples were collected from each culture prior to the lights coming on to ensure cells were non-motile (Yacobovitch et al., 2004). DEVELOPMENT OF MONOCLONAL ANTIBODIES BINDING TO MUCILAGE FROM SYMBIODINIUM CULTURE. Two antibodies that bind components of Symbiodinium mucilage were used in this study. The first, PC3, was previously developed by Wakefield (2001) against a component of the mucilage from Symbiodinium ITS2 ?type? A4 of Aiptasia pallida 41 from the Florida Keys (Santos et al., 2002; Santos, personal communication). The second, BF10, was generated (see below) for this study using a Briareum sp. clade F (chloroplast large subunit (cp23S)-ribosomal DNA genotype F178) alga originally collected from the Florida Keys. Aliquots were divided into separate 15 ml glass centrifuge tubes and either left (1) untreated or subjected to (2) fixation in 4% paraformaldehyde containing 0.2 M Millonig's phosphate buffer for 30 minutes followed by rinsing in 0.2 M Millonig's phosphate buffer (3X) and resuspension in 15-16 megaohm Nanopure water (np-H2O), (3) boiling for 1 minute in Millepore filtered sea water (MFSW), (4) three rounds of freezing and thawing by transferring the sample to a 1.5mL microcentrifuge tube, dropping the tube in liquid nitrogen, then re-warming the tube, or (5) resuspending the sample in 50% ethanol and homogenizing with a Virtis Handishear Homogenizer for 2 minutes followed by centrifugation and resuspension in np-H2O. All 5 types of samples were combined and used as the antigen to produce monoclonal antibodies. Antibodies were produced and screened using standard monoclonal techniques as described by Wakefield (2001). In brief, this symbiosome slurry was injected into BALB\C mice followed by a second injection boost after 3 weeks. Blood serum was tested for reaction to the symbiosome slurry antigens using an enzyme-linked immunosorbant assay (ELISA); the mouse exhibiting highest titer of antibody was sacrificed and splenocytes were isolated. Hybridoma cells were created using a standard polyethylene glycol fusion technique with the splenocytes and AG- 8.653 myeloma cells 4:1. Antibody producing clones were screened by ELISA. Two antibody producing cell lines, BA6 and BF10, strongly labeled the mucilage of Symbiodinium cp23S- rDNA genotype F178. Hybridoma cells secreting these antibodies were cloned by limiting dilution and then recloned to assure that only this line of hybridoma cells was isolated. Antibody to be used in labeling was collected as spent culture medium, from the final cloned cell line, 42 aliquoted in 0.5mL amounts into 1.5 mL Eppendorf tubes, frozen, lyophilized, and frozen for future use. Table 1. Symbiodinium cultures examined for presence of antigens PC3 and BF10. Host Organism Culture ID Collection Location cp23S?rDNA designation Zoanthus sociatus Zs Jamaica A188 Unknown Host Y109 Japan A191 Cassiopea sp. CassKB8 Hawaii A194 Cassiopea xamachana Cx Jamaica A194 Cassiopea xamachana FLCass Florida A194 Pseudoplexaura porosa 719 Panama A194 Cassiopea sp. CassMJ300 Hawaii A198 Tridacna gigas T Hawaii A198 Cassiopea sp. CassEli Hawaii A198 Plexaura kuna 708 Panama Undetermined (A) Montastrea faveolata Mf11.4 Florida Undetermined (A) Aiptasia pallida FLAp4 Florida A193 Aiptasia pallida FLAp3 Florida A193 Aiptasia pallida FLAp210AB Florida B184 Plexaura kuna 706 Panama B184 Aiptasia pulchella Ap10 Japan B184 Porites evermanni Pe Hawaii B184 Zoanthus pacificus Zp Hawaii B184 Plexaura kuna 13 Florida B184 Plexaura kuna 702 Panama B211 Briareum asbestinum 579 Florida B223 Aiptasia pulchella Ap Hawaii B224 Plexaura flexuosa PurPFlex Florida B224 Montastrea faveolata Mf11.5b.1 Florida Undetermined (B) Unknown anemone Ap31 Japan C180 Montastrea faveolata Mf8.5Tb.2 Florida F178a Acropora sp. A002 Japan D206 Acropora sp. A001 Japan D206 Unknown CCMP421 New Zealand E202 Montipora verrucosa Mv Hawaii F178 Briareum sp. PtBr Florida F178 a This culture was classified as a clade C symbiont using the 18S RFLP (Santos, personal communication, 2012). Here, this strain was retested using the cp23S-rDNA sequence information and was reclassified as clade F178. Clade F and clade C are closely related and using the 18S RFLP they appear the same. 43 FIXATION AND IMMUNOLABELING FOR EPIFLUORESCENCE MICROSCOPY. Cultured Symbiodinium isolates (Table 1) were individually centrifuged and the culture medium removed. Pellets were resuspended in 4% paraformaldehyde in 0.2M Millonig's Phosphate Buffer and allowed to fix for 1 hour at room temperature. Samples were then rinsed once in 20mM PBS containing 0.1% sodium azide and 0.1% Triton X100 (PBS+) and then dehydrated and rehydrated through an ethanol series (np-H2O, 30% ethanol, 50% ethanol, 70% ethanol, 50% ethanol, 30% ethanol, np-H2O) for 5 minutes in each solution. Following rehydration, samples were blocked in 5% heat-inactivated goat serum in PBS+ (blocking medium) for 10 minutes at room temperature. Blocking medium was removed by centrifugation and samples were incubated with spent medium containing primary antibody (either PC3 or BF10) diluted 1:1 in blocking medium for 1 hour at room temperature (a third sample of each strain was incubated with only blocking medium for this hour as a negative control). Samples were then washed in PBS+ 3 times for 5 minutes each, followed by incubation in secondary antibody (Alexa fluor 488 Goat anti-mouse IgM (? chain) - Invitrogen, A-21042) diluted 1:300 in blocking medium for 30 minutes at room temperature in darkness (to prevent quenching). Samples were then rinsed 3 times in PBS+ for 5 minutes each and resuspended in a small amount of np-H2O as follows. Ten ?l of labeled sample was added to 7 ?l of suspended standard fluorescent beads (InSpeck Microscope Image Intensity Calibration Kit in Green, 2.5 ?m, I-7219) and plated on slides using a Shandon Cytospin 2 to ensure that beads and cells would lie flat on the surface of the slides. EPIFLUORESCENCE MICROSCOPY. Glycerol mounting media was used to mount cover slips on the slides which were then kept in pop up slide holders so that slides would remain covered before being viewed on the microscope. This would minimize any bleaching between taking the first micrograph and the last micrograph of a given run. A control slide was imaged at 44 the beginning of each run and then also imaged again at the end of the run to determine whether any quenching occurred during the period the microscope was being used. Slides were viewed on Nikon Eclipse E800 with a Qimaging Retiga camera under light of wavelength 488 nm (to visualize labeling) and 594 nm (to visualize algal autofluorescence). Micrographs were taken of at least 30 cells with adjacent standard fluorescent beads using NIS Elements Software (Nikon Corporation, 2006). All micrographs were taken using the same settings on the microscope and with all the same software settings. FLUORESCENCE AND STATISTICAL ANALYSIS. The fluorescence intensities of the algal mucilage layer and of the beads were measured using ImageJ software to calculate the highest brightness value from the histogram. The 0%, 0.3% and 1% InSpeck beads were used to create a standard curve. This standard curve could then be used to estimate the percent of fluorescence measured in the mucilage layer of the algal cell. Fluorescence differences among strains were compared using a one-way ANOVA (SAS Version 9.2) to ascertain if any significant differences in labeling existed amongst labeled Symbiodinium strains. Post-hoc multiple comparison tests were run with a Tukey-Kramer adjustment to examine which strains exhibited significant differences in fluorescence (SAS Version 9.2). EXTRACTION, AMPLIFICATION AND SEQUENCING OF Cp23S rDNA. Based on the labeling results (see below), there was a question about the cp23S-rDNA identity of Symbiodinium culture Mf8.5Tb.2. DNA extraction and sequencing were performed on this particular isolate as well as several others (to serve as controls) after the methods of Santos et al. (2002). PCR success was confirmed by electrophoresis in a 1% agarose gel and visualized by ethidium bromide staining and ultraviolet (UV) light. Amplicons were sent to Genewiz, Inc. for sequencing. 45 Figure 1. PC3 antibody labeling (Panels A-D) and BF10 antibody labeling (Panels E and F) of Symbiodinium strains. A. FLAp4 originally cultured from Aiptasia pallida collected from Long Key, FL. The cp23-rDNA genotype of this strain is A193. This strain contains the antigen PC3 antibody was created against and showed labeling. B. 702 originally cultured from Plexaura kuna collected in Panama. The cp23S-rDNA genotype is B211. This strain showed faint labeling. C. Ap31 originally cultured from an unknown anemone collected from Sesoko Jima, Okinawa, Japan. The cp23-rDNA genotype of this strain is C180. This strain did not show labeling. D. A002 originally cultured from Acropora sp. collected in Okinawa. The cp23S-rDNA genotype is D206. This strain did not show labeling. E. PtBr originally cultured from Briareum sp. collected from Long Key, FL. The cp23S-rDNA genotype is F178. This is the strain that contains the antigen that the BF10 antibody was created against and showed labeling. F. Mf8.5Tb2 originally cultured from Monstastrea faveolata collected from Florida. The cp23S-rDNA genotype determined in this study is F178. This strain showed very faint and diffuse labeling. All scale bars are 10 ?m. Arrows indicate a ring of green labeling around the cell; arrowheads indicate standard florescent beads. All images were adjusted using Photoshop auto levels to better visualize the florescence here. These adjustments were made only for visualizing and not for florescence measurements. 46 RESULTS DEVELOPMENT OF MONOCLONAL ANTIBODIES BINDING TO MUCILAGE FROM SYMBIODINIUM CULTURE. Two monoclonal antibodies (BA6 and BF10) were developed that labeled mucilage secreted by cultured Symbiodinium cp23S-rDNA genotype F178. In early examinations, BF10 labeled some isolates (typically isolates closely related to the isolate the antibody was created against), but did not label other isolates and was chosen for further investigation. In contrast, BA6 labeled mucilage from every Symbiodinium isolate tested (31 different cultures representing 18 cp23S-rDNA genotypes). IMMUNOCYTOCHEMISTRY. Incubation with the PC3 antibody (Wakefield et al., 2000) exhibited labeling only in Symbiodinium from clades A and B (Figure 1). In clade A, 1of 5 cp23S-rDNA genotypes (i.e., A191) did not show labeling (Table 2). Likewise for clade B, 2 of 4 cp23S-rDNA genotypes (i.e., B223 and B224) did not exhibit labeling. No labeling with the PC3 antibody was observed in the tested isolates from clades C, D, E and F (Table 2). Incubation with the BF10 antibody exhibited labeling only in 2 isolates, one of cp23S-rDNA genotype F178 and one of unknown cp23S-rDNA genotype (culture ID Mf8.5Tb.2). Analysis of the cp23S-rDNA sequence of culture Mf8.5Tb.2 identified this as cp23S-rDNA genotype F178 (Figure 3). STATISTICAL ANALYSIS OF ANTIGEN LABELING AMONG SYMBIODINIUM CLADES AND STRAINS. There were significant differences between Symbiodinium isolates in labeling intensity with the PC3 and BF10 antibodies (p<0.0001, one-way ANOVA). Multiple comparison tests found that the isolates could be divided into significantly different categories (Figure 2). For example, non-labeling isolates were significantly different from all those that labeled. Amongst those that labeled with the PC3 antibody, there are several groups composed 47 Table 2: Labeling fluorescence presence of each Symbiodinium culture with each of 2 antibodies. Host Organism Culture ID Cp23S-rDNA Label PC3 Label BF10 Zoanthus sociatus Zs A188 + - Unknown Host Y109 A191 - - Aiptasia pallida FLAp4 A193 + - Aiptasia pallida FLAp3 A193 + - Cassiopea sp. CassKB8 A194 + - Cassiopea xamachana Cx A194 + - Cassiopea xamachana FLCass A194 + - Pseudoplexaura porosa 719 A194 + - Cassiopea sp. CassMJ300 A198 + - Tridacna gigas T A198 + - Cassiopea sp. CassEli A198 + - Plexaura kuna 708 Undetermined (A) + - Montastrea faveolata Mf11.4 Undetermined (A) - - Aiptasia pallida FLAp210AB B184 + - Plexaura kuna 706 B184 + - Zoanthus pacificus Zp B184 + - Aiptasia pulchella Ap10 B184 + - Porites evermanni Pe B184 + - Plexaura kuna 13 B184 + - Plexaura kuna 702 B211 + - Briareum asbestinum 579 B223 - - Aiptasia pulchella Ap B224 - - Plexaura flexuosa PurPFlex B224 - - Montastrea faveolata Mf11.5b.1 Undetermined (B) - - Unknown anemone Ap31 C180 - - Montastrea faveolata Mf8.5Tb.2 F178a - + Acropora sp. A002 D206 - - Acropora sp. A001 D206 - - Unknown CCMP421 E202 - - Montipora verrucosa Mv F178 - - Briareum sp. PtBr F178 - + a This culture was classified as a clade C symbiont using the 18S RFLP (Santos, personal communication, 2012). Here, this strain was retested using the cp23S-rDNA sequence information and was reclassified as clade F178. Clade F and clade C are closely related and using the 18S RFLP they appear the same. 48 of isolates that are not significantly different from one another, but that were different from other groups of isolates (indicated by lines connecting isolates; Figure 2). Analysis of labeling with BF10 showed that most isolates did not label, and did not exhibit significantly different levels of background fluorescence, from other non-labeling isolates. The two isolates that did label with this antibody were significantly different in their levels of fluorescence, both from each other and from all other isolates (indicated by symbols above the bar; Figure 2). Figure 2: Graph of Percent Fluorescence of PC3 antibody labeling (filled bars) and BF10 antibody labeling (empty bars). Error bars indicate the standard deviation for each sample. Statistical analyses were performed comparing the PC3 labeling amongst all Symbiodinium cultures and another set of analyses were performed comparing the BF10 labeling amongst all Symbiodinium cultures. Solid lines indicate significance when labeling with the PC3 antibody. Cultures grouped under the same black line are not significantly different from each other but are significantly different from all other cultures (P<0.005). Symbols are used to indicate significance when labeling with the BF10 antibody. All other open bars did not show labeling with the BF10 antibody and are not significantly different from each other or from background fluorescence. Bars labeled with a star and hexagon are significantly different from all other bars (P<0.005). All bars falling below the dotted line do not exhibit specific labeling but are representative of minor differences in background fluorescence. 49 DISCUSSION Variation in symbiont strain both genetically, as seen in many studies (Rowan and Powers, 1991b; Carlos et al., 1999; LaJeunesse and Trench, 2000; LaJeunesse, 2001; Pochon et al., 2001; Pochon et al., 2004; Pochon and Gates, 2010), and antigenically as shown here, indicate a basis for specificity between the host and symbiont. The nature of the host and symbiont relationship is quite specific in many cases (LaJeunesse, 2002; Rodriguez-Lanetty et al., 2004; Santos et al., 2003). However, host colonies that are large or shaped in such a way that different areas of the colony have different light regimes may have different symbiont populations in these different habitats (Rowan et al., 1997; LaJeunesse and Trench, 2000; Rodriguez-Lanetty et al., 2001; van Oppen et al., 2001; LaJeunesse et al., 2003; Ulstrup and van Oppen, 2003; Iglesias-Prieto et al., 2004). For instance, in A. tenuis colonies at Pioneer Bay, C1 was found on the shaded side of the colony while C2 was found on the side of the colony exposed to light (van Oppen et al., 2001). This is indicative of a relatively complex process occurring between the host and the symbiont. The process of initiation of the symbiosis between Symbiodinium and its cnidarian host contains multiple steps. First the host and symbiont must come into contact with each other. Thus, as the host is sessile in most cases of horizontal transmission, it is likely that the process of encounter is often initiated by the motile stage of the symbiont. Symbiodinium have been shown to exhibit chemotaxis towards aposymbiotic host organisms, fed symbiotic hosts, and brine shrimp nauplii. However, no chemotaxis toward starved host organisms has been documented suggesting that at least part of the taxis is directed toward ammonia or nitrate (Fitt, 1984). Further investigation found that the taxis is directed toward a variety of nitrogen compounds such as ammonium, nitrate, urea, and some amino acids, but there was no taxis toward 50 phosphate, sulfate, vitamins, trace metals, or sugars (Fitt, 1985). These studies indicate that Symbiodinium taxes toward either a host or a prey item for the host when there is nitrogenous compound present. Investigations into coral bleaching indicate that bleaching hosts release Figure 3. PC3 and BF10 antigen presence in Symbiodinium strains indicated on cp23S-rDNA phylogeny. Strains inside a solid box showed some level of PC3 labeling. Strains inside a dashed box showed labeling with BF10 antibody. Symbiodinium strains without a box were tested here and showed no labeling with either antibody. Strains indicated as free-living dinoflagellates were not tested in this study. This phylogeny is an edited version of one originally presented in Santos et al, 2002. The phylogeny was constructed by maximum-parsimony (MP) of the cp23S-rDNA Domain V sequences. Values at nodes indicate bootstrap support vales (percentages based on 1000 re-samplings). The most parsimonious tree was 436 steps in length with a consistency index of 0.77. The host name a sample was collected from is listed with the culture ID and location of collection in parentheses. 51 nitrogenous compounds (Perez and Weis, 2006) or show less activity of nitric oxide synthase (Trapido-Rosenthal et al., 2001). Thus, symbionts and hosts may have a mechanism for reinfection via symbiont chemotaxis to hosts stressed by episodic bleaching by virtue of higher production of nitrogen compounds. Once the host and symbiont are in close proximity the symbiont must be ingested by the host. There is some level of recognition that occurs at this point as not all potential symbionts will enter a host coelenteron (Trench et al., 1981). Subsequently, symbionts are phagocytosed into the gastrodermal cells without entering a digestive endocytic pathway. The process of phagocytic uptake has been directly observed in Chlorohydra (McNeil, 1981). Once the symbiont is within the host cells, it must survive any attempt to be digested by the host. Any phagocytosed dead Symbiodinium will quickly be digested and only live Symbiodinium will be maintained in the gastrodermal cells indicating that living cells have some characteristic involved in symbiont uptake (Fitt and Trench, 1983). Examinations of infection in Fungia scutaria show that multiple strains of living Symbiodinium can initially infect the host, but after a short time only the preferred type of symbiont is present in the larvae (Rodriguez-Lanetty et al., 2006). In some situations, it appears that this early uptake period has few barriers to prevent non-specific associations with non-preferred Symbiodinium strains and that more infectious or opportunistic strains, such as clade D strains, can advantageously colonize a host (Abrego et al., 2009a). At some point, either during these initial steps or shortly following them, cell-cell recognition processes establish a more specific relationship between a host and its preferred symbiont. While it is possible that the selection we see is not due to early cell-cell recognition but instead due to which Symbiodinium isolate is better adapted to grow in that host environment, the symbiont would presumably still secrete cell-cell recognition molecules to avoid digestion and 52 bypass any host immune response. Regardless of which type of system is occurring here, the molecules involved in infection and specificity must be important during these early steps in infection. A number of studies have determined which Symbiodinium strains associate with which host cnidarians and the conditions that support such associations. In most cases, infection begins with a limited number of strains entering the host (often only a subset of what is present in the water column) and after a relatively short time it appears that the preferred association will establish itself (Belda-Baillie et al., 2002; Berkelmans and van Oppen, 2006; Thornhill et al., 2006). The preferred associations fall into the full gamut of possibilities. They can be highly specific, where one host is found with only a single symbiont strain on a given reef (LaJeunesse, 2002, Goulet and Coffroth, 2003a, Goulet and Coffroth, 2004, Thornhill et al., 2006). Alternatively, the association my be somewhat flexible where one host has a dominant symbiont but it may be one of a few strains(LaJeunesse et al., 2003, Iglesias-Prieto et al., 2004). In mixed assemblages one host might have a dominant strain with one or more background strains that vary between host animals (Rowan and Knowlton, 1995, Rodriguez-Lanetty et al., 2001, Santos et al., 2003). Hosts may also associate with several strains that occupy specifically low-light, medium light, and high light regions of the host (Rowan and Knowlton, 1995; Secord, 1995; Rowan et al., 1997; LaJeunesse and Trench, 2000; Rodriguez-Lanetty et al., 2001; van Oppen et al., 2001; Diekmann et al., 2003; Goulet and Coffroth 2003a, b; LaJeunesse et al., 2003; Ulstrup and van Oppen, 2003; Goulet and Coffroth, 2004; Iglesias-Prieto et al., 2004). The diversity of associations suggest that processes that establish a particular symbiotic state may involve a combination of cell-cell recognition and differential survival of symbionts. In this scheme, as the 53 association develops, Symbiodinium that are better adapted to the local host microenvironment steadily displace those that are less well adapted. Work on symbioses involving Chlorella has implicated the algal surface layer in the cell- cell recognition and initiation of the symbiosis (Meints and Pardy, 1980; Reisser et al., 1982). In this study we have investigated the mucilage that comprises the outermost surface of the Symbiodinium cell to determine if molecular variation exists within this layer that could be used in a cell-cell recognition process during the initiation of the symbiosis. One of the antibodies, BA6, created against a clade F algal mucilage layer positively labeled all cultures that were tested. Previous work has described similar components among strains in the outer mucilage layer of macromolecules (Markell et al., 1992). While this antigen is not a good candidate for identifying differences amongst Symbiodinium strains that may be useful in recognition, it does suggest that the mucilage in all strains has some components in common. These common elements could include compounds that predispose most strains to be taken up during the early stages of infection (Coffroth et al., 2001; Rodriguez-Lanetty et al., 2006; Abrego et al., 2009a). In order for this mucilage layer to be involved in the establishment of the symbiosis there must be differences in the chemical composition of this layer that could be part of a specific host-symbiont recognition process. Using both the BF10 antibody created here and the PC3 antibody (Wakefield and Kempf, 2001), we detected antigenic variation among Symbiodinium strains. BF10 was highly specific, labeling only two cultures, both in clade F. PC3 antigen was found to be present only in clades A and B with some variation within these clades. In clade A, only one known cp23S-rDNA strain did not possess PC3 antigen (another culture did not label but the cp23S-rDNA strain type of this culture is unknown). In clade B, of the 4 distinct cp23S- rDNA types tested, 2 possessed the PC3 antigen while 2 did not. The other clades tested with 54 this antibody (clade C, D, E, and F) all showed no labeling. This is consistent with the finding that different isolates of Symbiodinium possess some of the same, but also many different, cell wall associated macromolecules (Markell et al., 1992). These results indicate that the mucilage layer possesses molecular variation among Symbiodinium strains and that the antigenic composition of this layer is, to some extent, correlated to genotype (Figure 3). The strain that exhibited the weakest specific labeling was a clade A198 culture while other clade A strains (and even another culture from a different host with the same cp23S-rDNA strain, A198) showed stronger labeling. Thus, the cp23S-rDNA type did not correlate with differences in intensity of fluorescent label. For example, the strain A198 cultures from different hosts (see Table 1 for host species and culture identity) show considerable variation in labeling intensity despite having the same cp23S-rDNA genotype (Figure 2). This indicates that while there is significant variation in the amount of antigen present in the mucilage, the variation is not related to the cp23S-rDNA strain identity. Instead the observed variation is almost certainly the result of as yet unknown genetic and/or physiological differences in mucilage secretion between cultures from different hosts instead of the genotype strain as determined by cp23S-rDNA. We have seen variation in antigenicity in the mucilage that forms the outermost surface layer of material surrounding the Symbiodinium cell. Since this layer is the part of the symbiont that actually contacts the host it would have to be the location for any recognition/communication that occurs between the host and the symbiont. While other investigations have recognized that the algal surface would have to perform this function, e.g., the cell wall (Lin et al., 2000), exuded macromolecules (Markell et al., 1992; Markell and Trench, 1993; Markell and Wood-Charlson, 2010) or simply cell-surface glycans (Logan et al., 2010), little is known about the molecular processes that function in cell-cell recognition 55 between host and Symbiodinium sp. Our study further confirms variation in cell surface molecules and demonstrates that there is variation within the mucilage component that demonstrates a correlation with symbiont clade. This is a first step in identifying components of the algal cell that may act in recognition/communication between host and symbiont. Lectin-carbohydrate reactions have been implicated in a number of cell-cell recognition processes (Weis and Drickamer, 1996), including those involving organisms from very different groups (such as the Rhizobium bacteria symbiosis with legumes and the immune system in animals; Bhuvaneswari et al., 1977). Due to the nature of cnidarian Symbiodinium symbioses it is very likely that a similar process occurs in these associations. A mannose-binding lectin has been found in Acropora millepora that binds to both bacterial pathogens as well as Symbiodinium (Kvennefors et al., 2008). Another group of studies investigated the presence of lectins in Sinularia lochmodes and discovered a D-galactose-binding lectin (SLL-2; Jimbo et al., 2000). Subsequent studies revealed that this lectin 1) localized to the cell surface, 2) arrested motility in Symbiodinium motile stages, and 3) bound D-galactose within the symbiont glycoconjugate (Koike et al., 2004; Jimbo et al., 2013). The presence of these types of molecules and their localization to the Symbiodinium cell surface suggest that this type of response may be involved in establishing and maintaining the association between Symbiodinium and their cnidarian hosts. It has been demonstrated that the Symbiodinium outer covering (sometimes referred to as exudate or cell wall, but here referred to as the mucilage) contains macromolecules with carbohydrate components (Markell and Trench, 1993; Lin et al., 2000). Symbiodinium strains have been shown to have reduced infectivity if surface glycoproteins either embedded in the cell wall or external to the cell wall are masked or digested (Lin et al., 2000; Wood-Charlson et al., 56 2006). Here we have tested a wider variety of Symbiodinium strains and determined that there is widespread variation in antigenicity that is consistent with previous studies (Markell et al., 1992; Markell and Trench, 1993; Logan et al., 2010) that examined usually only a few strains. This study gives support to previous work investigating the cell surface of Symbiodinium by using 2 antibodies to reveal widespread variation in the outermost layer of the Symbiodinium cell correlated to strain. Our results strengthen the argument that the mucilage layer has the required variation to establish specificity between host and symbiont, as well as function in host - symbiont communication in the intact symbiosis. While the exact process of recognition and maintenance of Symbiodinium symbioses and the molecules involved are still unknown, this study supports the hypothesis that the potential for cell-cell recognition between the host and symbiont is present in the outermost layer of the symbiont cell. Future studies will hopefully reveal the molecules that drive initiation and maintenance of Symbiodinium symbioses. 57 III. ULTRASTRUCTURAL VARIATION AMONGST SYMBIODINIUM ISOLATES ABSTRACT Symbiodinium are unicellular dinoflagellates that reside intracellularly in a variety of invertebrate hosts, including cnidarians. There is great diversity amongst isolates of this genus; however, very few ultrastructural studies have examined differences among them. Here, ultrastructural examinations using High Pressure Rapid Freezing were performed on 5 isolates (2 from clade A, and 1 from clade B, 1 from clade F, and a free-living isolate from clade E) to examine variation amongst these strains that could affect the ability of a particular symbiont to reside within a particular host. All 5 isolates studied exhibited the standard dinokaryon nucleus, a peripheral multi-lobed chloroplast with a single stalked pyrenoid surrounded by a starch cap, and dinoflagellate-type mitochondrion with tubular cristae. Electron dense inclusions were typically positioned near the cell periphery and may be related to mucocysts sometimes seen in dinoflagellates. Some isolates (CCMP421 and Y109) had a few cells that exhibited large electron dense structures containing calcium oxalate crystals. Interestingly, the cell size amongst different isolates was significantly different and consistent with previous studies on the same or similar isolates. Examination of the ultrastructure of 5 isolates of Symbiodinium revealed that there is very little variation in morphological characters. Therefore, it seems unlikely that isolates or species of Symbiodinium can be usefully differentiated on the basis of morphology alone. However, characterization of the morphology of the genus Symbiodinium as it relates to its symbiosis with the host may lead to a better understanding of how the association between host and symbiont is established and maintained. 58 INTRODUCTION Symbiodinium microadriaticum was originally described using cultured cells collected from Cassiopeia sp. (Freudenthal, 1962). Using light microscopy, the life cycle of Symbiodinium microadriaticum was found to begin with a vegetative cell which appears to contain numerous irregular shaped chloroplasts, an inconspicuous nucleus, and, as the cell ages, an increasingly large ?assimilation product?. This vegetative cell can divide forming two nearly identical daughter cells or thicken its cell wall and encyst. The vegetative cyst can then form a zoospore motile form or divide within the cyst. At the time it was thought possible that gametes could form from the cyst stage, but this was not actually observed (Freudenthal, 1962). An initial ultrastructural investigation described the periplast (cell wall) as a thick amorphous layer between 2 membranes (Kevin et al., 1969). The chloroplast was described as a single, multi- lobed, peripheral structure surrounded by an envelope of 3 membranes and having repetitive lamellae each consisting of 3 closely stacked thylakoids. All cells had a large nucleus enclosed by an envelope with pores and containing chromosomes having a typical dinoflagellate appearance, i.e., condensed and with a tightly coiled structure (Kevin et al., 1969). Additional identified structures included mitochondria, calcium oxalate crystals inside large vacuoles, and lipid droplets (Kevin et al., 1969). This description was done at a time when Symbiodinium microadriaticum was thought to be a single pandemic species (Taylor, 1974). However, further descriptive studies found significant variation between cultured Symbiodinium isolates from a wide variety of hosts. Differences included variation in isoenzyme and soluble protein patterns (Schoenberg and Trench, 1980a), morphology (Schoenberg and Trench, 1980b), specificity as measured by infectivity in hosts (Schoenberg and Trench, 1980c), motility rhythm (Fitt et al., 1981), and 59 chromosome volume (Blank and Trench 1985). These studies led to the understanding that there were a number of distinct species that were erroneously grouped into the single species Symbiodinium microadriaticum. With the understanding that there were many different isolates within the genus Symbiodinium new species began to be named based on biochemical, physiological, morphological, and behavioral differences (Trench and Blank, 1987). Three of the earliest novel species named after S. microadriaticum include S. goreauii, S. kawagutii, and S. pilosum (Trench and Blank, 1987). Despite the vast variety that was seen in the earliest genetic studies (Rowan and Powers, 1991a; Rowan and Powers, 1992), further formal species descriptions were very few (2) and initially still based on ultrastructure. The next formal species description was Gymnodinium linucheae (later reclassified as Symbiodinium linucheae; Trench and Thinh, 1995). Finally, Symbiodinium natans was described using a combination of light and electron microscopy combined with nuclear-encoded partial LSU rDNA sequence information (Hansen and Daugberg, 2009). While the naming of species may prove informative, what has been consistently used to understand the phylogeny of Symbiodinium is identification of type using a variety of genetic markers. Currently there are 9 clades into which isolates of this genus can be placed. Clades A, B and C were the first identified using restriction fragment length polymorphisms (RFLPs) (Rowan and Powers, 1991b). Later, clades D (Carlos et al., 1999), E (LaJeunesse and Trench, 2000; LaJeunesse, 2001), F (LaJeunesse, 2001), G (Pochon et al., 2001), H (Pochon et al., 2004), and I (Pochon and Gates, 2010) were added based on variation in the internal transcribed spacer (ITS) sequence (LaJeunesse, 2001; Pochon et al., 2004), 18S rDNA sequence (LaJeunesse, 2001), 28S rDNA sequence (Pochon et al., 2006), and cp23S rDNA sequence (Santos et al., 60 2001; Santos et al., 2002). All these studies and many more have come to the same conclusion, namely that there is a huge amount of variation between isolates within genus Symbiodinium (Coffroth and Santos, 2005). Along with the investigation into Symbiodinium genetics, further ultrastructural examinations were undertaken to examine morphological variation amongst isolates. Examination of Symbiodinium from Zoanthus sp. confirmed the presence of a lobed chloroplast and stalked pyrenoid (Van Thinh et al., 1986). Symbionts from Montipora verrucosa were found in situ to occur as a coccoid cell with an amphiesma of 5 membranous layers; a single, peripheral lobed chloroplast with parallel thylakoids and a stalked pyrenoid; and a dinokaryon nucleus with 26 chromosomes (Blank, 1987). Symbionts similar to Symbiodinium microadriaticum from 3 foraminiferans (Leutengger, 1977) and Tridacna maxima (Bishop et al., 1976) were examined ultrastructurally and were similar to previous descriptions of Symbiodinium. As further ultrastructural examinations of isolates symbiotic with cnidarians were performed, refined descriptions of the ultrastructure of some cell structures, such as the chloroplast (Hannack et al., 1997) and cell covering including thecal vesicles were made (Wakefield et al., 2000). Table 1: Genetic characterization of the Symbiodinium isolates in the current study. Species Culture Host Clade Cp23S Nr28S Nr18S ITS1/5.8S/ ITS2 Symbiodinium sp. Y109 Unknown host A191 AY035413 Symbiodinium sp. FLAp4 Aiptasia pallida A193 AY035404 AF427453 AF427441 AF427465 Symbiodinium pulchrorum HIAp or Ap Aiptasia pulchella B224 AY035421 AF427457 AF427445 AF360564 Symbiodinium sp. CCMP 421 None E202 AY055240 Symbiodinium kawagutii Mv Montipora verrucosa F178 AY035422 AF427462 AF427450, AY035413 AF360577 61 Since ultrastructural studies have all found similar cell structure while the genetic studies have all found a huge variety of molecular differences within the genus, it is difficult to determine if there are morphological differences that would be informative in regard to the great genetic variety that exists amongst isolates of the genus Symbiodinium. Along with all the genetic information and a variety of physiological studies examining cultured Symbiodinium it is important to begin parsing through the information and relating ultrastructural, genetic, and physiological traits to give a thorough understanding of the biology of the symbiont. Here we provide ultrastructural details of genetically well-characterized isolates in 4 clades in order to provide a basis for variations between previously undescribed isolates and to compare ultrastructure between clades. This work is meant to provide a baseline against which future ultrastructural analyses can be compared. METHODS METHODS FOR ASSESSMENT OF ULTRASTRUCTURAL DIFFERENCES IN SYMBIODINIUM ISOLATES. In looking for ultrastructural differences between Symbiodinium isolates, it is critical that all isolates be cultured under identical conditions and that they be sampled at the same time in the cell cycle. Similarly, it is also critical that all samples be processed in the same manner using the same fixation and embedding techniques. To that end, we suggest the following approach as a standardized method that can be followed by investigators doing future comparisons of these and other isolates. The culture methods are chosen because they are relatively standard among labs working with Symbiodinium. The high pressure rapid freezing technique for electron microscopy is chosen because it offers the best fixation with the fewest artifacts. 62 CULTURE GROWTH. Cultures were grown in F/2 media (Guillard and Ryther, 1962) in 125mL erlenmeyer flasks under 12/12 light dark cycle at about 24.5?C (? 0.5?) at Auburn University for 3 weeks. Cultures were transported to the culture room of Dr. Fitt at the University of Georgia and allowed to grow for 2 weeks under a 12/12 light dark cycle at 26.5?C (? 0.5?). Isolates subjected to ultrastructural analyses are listed in Table 1. HIGH PRESSURE RAPID FREEZING. Fixative solutions were prepared several days prior to fixation. 0.1% uranyl acetate in HPLC grade acetone was mixed at room temperature 2 days prior to freezing and the day before freezing was transferred to a -80?C freezer. A vial of 25 mL of HPLC grade acetone was pre-chilled in the -80?C freezer overnight and then 1 g of OsO4 was added (making a 4% OsO4 solution) while the solution was kept on dry ice for 10 minutes before storage in a -80?C freezer the day before freezing. Isolated samples were collected at virtual 'dawn' as defined by the light/dark cycle by pipetting 100?L of cells+media using a Gilson pipetman starting at the bottom of the flask (where the cells accumulate) and slowly drawing the pipette up about 1 cm from the bottom. The pipette was then removed from the flask and the sample was transferred to a 1.5 ml microcentrifuge tube and centrifuged briefly (MyFuge at 2000g) to produce a loose pellet. Most of the remaining F/2 media was removed (leaving the cell pellet in about 50 ?L of F/2 media) and 50 ?L of 20% dextran in F/2 media was added and the cells were re-suspended in a final concentration of 10% dextran in 100 ?L of F/2 medium. The 10% dextran encouraged the cells to remain clumped together after freezing. Cells were transferred to planchettes using a 2-10?L Gilson pipetman and then rapidly frozen in a High Pressure Freezing Machine (HPM 010 ABRA Fluid AG). After freezing, pellets were transferred to cryotubes submerged in liquid nitrogen and then transferred to a pre-cooled dewer and maintained in liquid nitrogen until transfer into fixative. The 0.1% uranyl acetate in acetone and 63 4% OsO4 in acetone were then mixed (all vials kept on dry ice) in a 1:1 ratio and aliquoted into enough vials for the frozen samples. After 2 hours in the -80?C freezer, the frozen samples in planchettes were quickly added to the vials of 1:1, 0.1% uranyl acetate: 4% OsO4 in acetone, all vials and cryotubes being kept on dry ice. Samples in fixative were then placed in a - 80?C freezer for 4 days. Samples were next moved to a -20?C freezer for 3 hours, then to a 4?C refrigerator for 2 hours, and finally into a fume hood at room temperature for 30 minutes. After this gradual return to room temperature the fixative was removed and samples were rinsed 3 times in 100% HPLC grade acetone for 15 minutes each. Samples were then removed from planchettes and infiltrated with Low viscosity Embedding Media Spurr's Kit (hard recipe; Electron Microscopy Sciences) by first adding the embedding resin as a 1:2 mix in acetone and leaving it overnight at room temperature. The next day the samples were changed to 2 parts embedding resin to 1 part acetone for 8 hours and then into 100% resin overnight. The next morning samples were placed in fresh 100% resin for 8 hours. Near the end of the day the samples were transferred to catalyzed resin in Beem capsules. After 1 hour at room temperature in the catalyzed resin the samples were placed in an oven at 60?C and left for 24-48 hours to ensure hardening. TRANSMISSION ELECTRON MICROSCOPY. Samples were sectioned using a Reichert- Jung Ultracut E Microtome and placed on copper 75 mesh grids (EMS). Samples were stained using 4% aqueous solution of uranyl acetate for 10 to 20 minutes and then stained with 0.1% aqueous solution of lead citrate for 5 minutes. Samples were viewed on a Zeiss EM 10C 10CR Transmission Electron Microscope and digital images were taken using MaxIm DL5 software and SIA-L3C 4.3 megapixel (Scientific Instruments & Applications, Duluth, GA 30096). MEASUREMENTS AND STATISTICS. In order to obtain accurate measurements of cell 64 size, only cells larger than 4 ?m in diameter were measured and added to a list to obtain 100 measured cells from each isolate. Since cells were often oblong in shape, the diameter was measured twice, at the largest point and the smallest; only one of those measurements had to be greater than 4 ?m. These cells were then sorted by size (based on the greatest of 2 diameter measurements) and only the largest 30% (n =30) were used to determine average cell size. Measurements were made first using MaxIMDL5 software and then confirmed using Image-Pro Plus (Version 7.0; Media Cybernetics, 2009). The first measurement made on each cell was a cross-sectional area measurement. This was obtained by setting the scale of the image and then manually encircling the entire cell to get an area. Cell size differences (both of the greatest diameter and of a cross-sectional area measurement) among isolates were compared using a one- way ANOVA run on SAS (Version 9.2) to ascertain any significant differences. Post-hoc multiple comparison tests were run with a Tukey-Kramer adjustment to examine which strains exhibited significant differences in size. Additionally, thylakoid size, cell membrane size, cell wall size, and outer membrane size were analyzed using a one-way ANOVA run on SAS (Version 9.2) followed by a Tukey-Kramer adjustment. DNA EXTRACTION, AMPLIFICATION AND SEQUENCING OF Cp23S rDNA. DNA extraction and sequencing were performed on all 5 isolates according to Santos et al. (2002). PCR success was confirmed by electrophoresis in a 1% agarose gel and visualized by ethidium bromide staining and ultraviolet (UV) light. Amplicons were sent to Genewiz, Inc. for sequencing. 65 Figure 1: Representative examples of 5 isolates of Symbiodinium showing characteristic ultrastructure. All diameters listed are of the greatest distance across the cell in the case of oblong cells. A. An average cell of Y109 with a diameter of 8.5 ?m. The peripheral chloroplast and central dinokaryon nucleus are visible along with some inclusions. B. An average cell of FLAp4 with a diameter of 7.1 ?m. Here you can see the peripheral lobed chloroplast and the starch cap around the pyrenoid is visible. The section also shows the dinokaryon nucleus near the center of the cell. C. An average cell of Ap with a diameter of 7.9 ?m. This cell is filled with starch granules and lipid droplets. The peripheral chloroplasts are visible as is the pyrenoid though the stalk of the pyrenoid is not included in this section. D. An average cell of CCMP421 with a diameter of 10.1 ?m. The cell has peripheral chloroplast lobes, a dinokaryon nucleus, starch granules and lipid droplets throughout. E. An average cell of Mv with a diameter of 9.1 ?m. Here the dinokaryon nucleus is clearly visible pushed a bit to the edge of center by the presence of a number of starch granules and lipid droplets. The peripheral chloroplast lobes are visible as well. All scale bars are 1?m. 66 RESULTS SYMBIODINIUM ISOLATE Y109. Y109 was originally cultured by M. Hidaka from an unknown host in Okinawa, Japan. Coccoid cells measured during the log phase of growth range in average size from 8?m to 10?m in diameter (Average 9.0 ?0.83 ?m; n=30; Figures1A and 2). The nucleotide sequence of the cp23S-rDNA was previously reported (AY035413; Santos et al., Figure 2: Cell size of coccoid cells grown in culture during log phase growth. Symbols indicate which isolates are not significantly different from each other (p<0.05). Error bars indicate the standard error. Cell diameters are of the greatest straight-line distance across a cell; cell cross-sectional area was measured in Image-Pro Plus which allowed the area to be measured very accurately. 67 2002) and confirmed here. The chloroplast is located peripherally and a single stalked pyrenoid with a starch cap was observed (Figure 3A). Thylakoids are arranged in groups of 3, each 9.4 ? 1.2 nm (n=10; Figure 4A and 5). The lamellae of 3 thylakoids are closely pressed together as is common in dinoflagellates and in Symbiodinium. The accumulation body is circular and centrally located (Figure 6A). During log phase growth, cells contain many medium to large lipid and starch granules (Figure 6A). The cell is bordered by an outer membranous layer that averages 10.5 ?1.3 nm (n=10, Figure 7A and 8). Inside the outer membranous layer, the cell wall averages 134.6 ?47.9 nm (n=10). The plasma membrane (7.3 ? 1.1 nm; n=6) is just outside other membranous or thecal vesicle structures that are sometimes seen just below the plasma membrane (Figure 7A). Some cells contain a large dark central structure that contains plentiful calcium oxalate crystals (Figure 9A, C, and E). At the periphery of the cell, usually between the cell membrane and chloroplast, there are many small dark staining granules (Figure10A and D). The nucleus has the standard dinokaryon nuclear structure and is located centrally (Figure11A). The mitochondria also have the tubular cristae typical in dinoflagellates (Figure 12A). ULTRASTRUCTURAL DESCRIPTION OF SYMBIODINIUM ISOLATE FLAP4. FLAp4 was originally isolated by S.R. Santos from Aiptasia pallida in the Florida Keys. Coccoid cells of isolate FLAp4 during log phase growth averaged 6.5?m to 8.5?m in diameter with an average diameter of 7.5 ?0.6 ?m (n=30) (Figures 1B and 2). The cells have a peripheral chloroplast attached to a single stalked pyrenoid with a starch cap (Figure 3B). Within the chloroplast, thylakoids are arranged in groups of 3, each approximately 10.5 ?1.8 nm (n=10; Figure 4B and 5). The lamellae of 3 thylakoids are stacked closely as is common in dinoflagellates and in Symbiodinium. The accumulation body is circular and centrally located (Figure 6B). The cell wall (122 ?51.7 nm) is bordered by an outer membranous layer 9.1 ? 0.9 nm across (n=10, 68 Figure 3: Examples of the pyrenoid with starch cap in 5 isolates of Symbiodinium. A. A cell of Y109 with the pyrenoid clearly indicated by the starch cap surrounding. B. An example of the pyrenoid in FLAp4 with the stalk visible. C. A cell of Ap with the classic pyrenoid and starch cap. D. A cell of CCMP421 the stalk only partially visible. E. An example from Mv with no visible stalk of the pyrenoid. F. A high magnification image of the pyrenoid from Ap; notice that there are no lamellae present within the pyrenoid. A-E have scale bars of 2?m. The scale bar in F is 500nm. 69 Figure 7B and 8). Inside the cell wall lies a plasma membrane (7.0?0.9 nm; n=8) and below that membrane other membranous structures are often seen (Figure 7B and F). Cells often have electron-dense structures near the periphery which may be related to mucocyts seen in other dinoflagellates (Figure 10 B and E; Hoppenrath and Leander, 2008). The nucleus has the standard dinokaryon structure and is usually central or close to central (Figure 11B). The mitochondrion also has the typical dinoflagellate appearance with tubular cristae (Figure 12B). Figure 4: An example of thylakoids in 5 Symbiodinium isolates. A. Isolate Y109 thylakoids. B. An example of the thylakoid in FLAp4. C. A stack of 3 thylakoids in isolate Ap. D. A cell of CCMP421 with they stack of 3 thylakoids clearly visible. E. An example from Mv of the thylakoid arrangement. Scale bars are 100nm. All grana were composed of 3 thylakoids. One stack of 3 thylakoids is indicated by brackets. 70 ULTRASTRUCTURAL DESCRIPTION OF SYMBIODINIUM ISOLATE AP. Isolate Ap was originally cultured by R. A. Kinzie from the host Aiptasia pulchella from Kaneohe Bay, Hawaii. Subsequently, this isolate was named as S. pulchrorum by R. K. Trench in Banaszak et al. (1993). During log phase growth, coccoid cells of S. pulchrorum vary in diameter from 6.8?m to 9?m in diameter with an average of 8 ? 0.86 ?m (n=30; Figures 1C and 2). The chloroplast is located peripherally with a single stalked pyrenoid with a starch cap (Figure 3C and F). Thylakoids are arranged in stacks of 3 with the average thickness of each thylakoid of 9.4 ?1.1 nm (n =10; Figure 4C and 5). In the interior of the cell, starch granules are relatively large while lipid droplets are smaller (Figure 6C). The cell wall (127.7 ?32.4 nm) is bordered by an outer membranous layer 10.0 ?2.0 nm (n=10, Figure 7C and 8). Interior to the cell wall is a plasma Figure 5: Thylakoid membrane size in 5 isolates of Symbiodinium sp. There was no significant difference in thylakoid size or in the organization of the stacks of thylakoids. All thylakoid membranes averaged between 8.5 and 10.5 nm. 71 membrane (7.4?1.4 nm; n=9) often bordering other membranous or thecal vesicle-like structures (Figure 7C). Some cells have small dark vesicles that may be related to mucocysts (Figure 9C; Hoppenrath and Leander, 2008). The nucleus has the standard dinokaryon appearance and is located centrally (Figure 11C) while the mitochondria also has the standard dinoflagellate appearance with tubular cristae (Figure 12C). ULTRASTRUCTURAL DESCRIPTION OF SYMBIODINIUM ISOLATE CCMP421. CCMP421 was originally cultured from water samples collected from Wellington Harbor, New Zealand and was assumed to be a free-living isolate of Symbiodinium. Coccoid cells measured during the log phase of growth range in size from 9 to 13?m in diameter (Average 10.2 ? 1.01 ?m, n=30; Figure 1D and 2). The nucleotide sequence of the cp23S-rDNA was previously reported (AY055240; Santos et al., 2002) and confirmed here. The cp23S-rDNA sequence indicates that this species of Symbiodinium is in clade E. The chloroplast has a single stalked pyrenoid with a starch cap (Figure 3D). The chloroplast is located peripherally and appears branched so it may be a single, large branched chloroplast. Thylakoids are in groups of 3 with the standard width of 10.7 ?1.4 nm per each thylakoid (n=10; Figure 4D and 5). There are many small to medium starch granules and lipid droplets located within cells in the log phase stage of growth (Figure 6D). The accumulation body is always circular and stains darkly (Figure 6D). The cell wall is bordered by an outer membranous layer 10.5 ? 1.9 nm across (n=10, Figure 7D and 8). The cell wall averages 117.5?37.7 nm (n=10). Inside the cell wall lies a plasma membrane (7.5?1.4nm; n=8; Figure 7D and 8). Cells often contain a large, amorphous, darkly staining, non-membrane bound structure within which calcium oxalate crystals are present (Figure 9B, D, and F; Franceschi and Nakata, 2005). The dinokaryon nucleus is located centrally (Figure 11D) and the mitochondria have tubular cristae common in dinoflagellates (Figure 12D). 72 Figure 6: Comparison of accumulation body size inclusions (starch granules and lipid droplets) amongst 5 species of Symbiodinium. Accumulation bodies indicated with AB, starch by S, and lipid droplets by L. A. Y109, B. FLAp4, C. Ap, D. CCMP421, E. Mv, F. High magnification of the accumulation body fromY109. Scale bars 1?m in B and C; 2 ?m in A, D, and E; and 500nm in F. 73 ULTRASTRUCTURAL DESCRIPTION OF SYMBIODINIUM ISOLATE MV. Mv was originally isolated by R. A. Kinzie from Montipora verrucosa in Hawaii. Subsequently, this isolate was described as S. kawaguti by Trench and Blank (1987). Coccoid cells of S. kawagutii are large with a diameter ranging from 8 ?m to 10.6?m during log phase growth for an average of 9.05 ? 0.74 ?m (n=30; Figures 1E and 2). The chloroplast is located peripherally and connected to a stalked pyrenoid with a starch cap (Figure 3E). Within the chloroplast, thylakoids are arranged in groups of 3, each approximately 10.8?1.2 nm (n=10; Figure 4E and 5). Lamellae of 3 thylakoids are stacked closely as is common in dinoflagellates and in Symbiodinium. The interior of the cell has many starch granules and lipid droplets. Calcium oxalate crystals are rare. Accumulation bodies were rare at the early log phase of growth, but were similar in structure to the other isolates (circular, darkly staining; Figure 6E). The cell wall is bordered by an outer membranous layer 10.5 ?0.9 nm across (n=10, Figure 6). The cell wall averages 169.7 ?41.7 nm (n=10) and just inside the cell wall lies a plasma membrane (7.2?0.9 nm; n=7; Figure 7E and 8). The nucleus has the standard dinokaryon appearance and is located centrally (Figure 11E). The mitochondria contain tubular cristae (Figure 12E). DISCUSSION The general ultrastructure for Symbiodinium isolates appears to be consistent. All isolates studied here and all previous ultrastructural studies of Symbiodinium note a peripheral chloroplast with a stalked pyrenoid surrounded by a starch cap. Likewise, these chloroplasts all contain thylakoids in stacks of 3 and the size of the thylakoids does not vary amongst isolates or from what is typically seen in dinoflagellates (Dodge and Greuet, 1987). Symbiodinium also possesses a standard dinokaryon nucleus with permanently condensed chromosomes (Dodge and 74 Figure 7: Images of the Symbiodinium cell periphery in 5 isolates. A. Y109, B. FLAp4 cell periphery which also contains a cross-section of a flegellum, C. Ap, D. CCMP421, E. Mv. F. FLAp4 example of a recently divided cell that currently has 2 cell walls. In the cyst stage cells can divide within the cell wall and remain 2 cells (each with their own cell wall) inside the parent cell wall for some time. Scale bars in A, B, D, E, and F are 100nm. Scale bar in C is 200nm. The outer double membrane is indicated by an arrowhead. The plasma membrane is indicated by the arrow. The bracket indicates the layer typically referred to as the cell wall or amorphous layer. The open arrowheads indicate membranous layers below the plasma membrane. These are best seen in panel F between the 2 cell walls. 75 Greuet, 1987). The cell periphery of Symbiodinium is interesting, the cell periphery is bounded by an outer membranous layer that is slightly wider than the plasma membrane (Average 10.2 ?0.78 nm, n=50; consistent with previous studies on Symbiodinium bermudense, Wakefield et al., 2000). The plasma membrane is the standard 7nm across. The cell wall varies greatly in thickness, probably due to the age of the individual cells as there is no significant difference between Symbiodinium isolates investigated here or any Symbiodinium species investigated previously (Symbiodinium sp., S. bermudense, and S. microadriaticum; Wakefield et al., 2000) . Isolate Y109 was previously used to elucidate the variation within the genus Symbiodinium using the cp23S-rDNA (Santos et al., 2002). Isolate Y109 is a clade A alga using the cp23S-rDNA method, but it has not previously been tested using other genetic methods or examined ultrastructurally or physiologically. From this study we note that while the Figure 8: Statistical analysis of the Symbiodinium cell periphery. The outer membrane size is indicated by the open bar using the left axis, there was no significant difference between isolates. The plasma membrane is indicated by bars with horizontal lines using the left axis. There was no significant difference between isolates. The size of the cell wall is indicated by the diagonal striped bars using the axis on the right, there was no significant difference. 76 ultrastructure is similar to other Symbiodinium isolates, there are some notable differences such as cell size. Isolate Y109 is significantly smaller than several of the others tested. Isolate FLAp4 was described using sequences from the cp23S-rDNA, nr28S, nr18S, and nrITS1/5.8S/ITS2 rDNA (Santos et al., 2002). Since there is very little ultrastructural or physiological data on this isolate, here we presented an ultrastructural description of the same isolate using the cp23S-rDNA sequence to confirm the culture identity. The cell size found in this study was consistent with the formal description of another species from the same host, Symbiodinium minutum (LaJeunesse et al., 2012). Here we saw that the ultrastructure of this isolate was very similar to other Symbiodinium isolates, but was significantly smaller in cell size and contained dark granules near the periphery. These electron dense structures are not membrane bound and are unidentified, but may have some relationship to mucocysts (Hoppenrath and Leander, 2008). Isolate Ap, Symbiodinium pulchrorum, was named based on a culture (Ap, the culture used here) isolated from Aiptasia pulchella in Hawaii (Kinzie and Chee, 1979). S. pulchrorum has not been formally described, however, genetic testing has revealed that this culture is a clade B1 Symbiodinium based on ITS2 sequencing (LaJeunesse, 2001). Sequencing of cp23S-rDNA, 18S-rDNA, n28S-rDNA, and ITS-rDNA supported the placement in clade B and identified the culture as a B224 (Santos et al., 2002; Santos et al., 2003; Pochon et al., 2006). The original isolate of Kinzie and Chee (1979) was also used to examine thermal tolerance and it was found that this isolate showed decreased growth rates at 29? and 31? C as compared to 27? C (Kinzie et al., 2001). Since this isolate has been well studied genetically it can be compared to many other new isolates. Interestingly, a different new species, Symbiodinium psygmophilum (LaJeunesse et al., 2012), is genetically identical to S. pulchrorum (isolate Ap) at the cp23S-rDNA sequence 77 Figure 9: A. Depictions of large amorphous dark structures (arrows) containing calcium oxalate crystals (asterisks) in cells of 2 isolates of Symbiodinium. A. C. and E. are isolate Y109. B. D. and F. are isolate CCMP421. A. and B. depict large, central electron dense structures. C and D depict smaller more peripheral examples of what appear to be similar electron dense structures. E. and F. are high magnification images of the calcium oxalate crystals contained in these structures. Scale bars in A, and C are 1?m. Scale bars in B and D. are 2?m. Scale bars in E and F are 500nm. 78 level, but different using the ITS2 sequence (LaJeunesse, 2001; Santos et al., 2002; Santos et al., 2003). The cell size is also similar between S. pulchrorum and S. psygmophilum possibly indicating that in some cases morphological characters are consistent with the genetic relationship between isolates. The ultrastructural description here finds that the isolate Ap is very similar to other Symbiodinium isolates, but is significantly smaller than some isolates and contains small electron dense structures at the periphery (similar to the structures identified in isolate FLAp4). Isolate CCMP421 is only the second free-living isolate of Symbiodinium described (the first is Symbiodinium natans; Hansen and Daugbjerg, 2009). This is also the only ultrastructural Figure10. Examples of peripheral electron dense structures. These structures are not present in all isolates and may be related to mucocysts. A and D. are isolate Y109. B. and E. are isolate FLAp4. C and F. are isolate Ap. Scale bars for A-C are 2?m. Scale bars for D-F are 500 nm, These putative mucocysts are indicated by black arrows. A white arrowhead was used to indicate a calcium oxalate crystal in E. 79 description of an isolate within the previously identified clade E. Like S. natans, CCMP421 is larger than the symbiotic species described and contains many storage products including lipid droplets and starch granules (Hansen and Daugbjerg, 2009). While S. natans and isolate CCMP421 are from two different clades within the genus Symbiodinium their shared life history pattern seems to have conferred upon them these similar morphological traits. If these two species are permanently free-living it is interesting that they seem to contain more storage products than symbiotic species and are significantly larger. Interestingly, when isolate CCMP421 and other Symbiodinium cultures were tested with a variety of lectin probes CCMP421 labeled less strongly with most of the lectin probes than the symbiotic cultures (Logan et al., 2010). This could indicate that lectins are involved in aspects of the symbiosis and such interactions are unnecessary for the free-living dinoflagellate. Isolate CCMP421 has been well studied genetically, beginning with an examination of the small ribosomal subunit (18S) RNA where it was found that the difference within the genus Symbiodinium was similar to other dinoflagellate orders and the first 3 clades, A, B, and C, were identified (Rowan and Powers, 1992). CCMP421, referred to as Gymnodinium varians in that study, was identified within clade B (though it was an outgroup from most of the clade - Rowan and Powers, 1992). Again, as G. varians, this culture was used to examine the LSU rRNA and grouped near clades B and C but not as a member of either (Wilcox, 1998). Subsequent work looking at the SSU rRNA found that this culture grouped with other Symbiodinium strains (Saldarriaga et al., 2001). Examinations of the chloroplast 23S-rDNA grouped this culture (now referred to simply by the culture identification code at the Provasoli-Guillard National Center for Marine Algae and Microbiota, CCMP421) in clade E (Santos et al., 2002; Santos et al., 2003). The cp 23S-rDNA and nr28S-rDNA sequences from this culture were used as the 80 only representative from clade E to examine evolutionary rates in Symbiodinium (Pochon et al., 2006). The cp 23S-rDNA sequence was then retested in 2012 to ensure that the culture housed at Auburn University had not been contaminated. Isolate Mv, Symbiodinium kawagutii, was originally described in 1987 (Trench and Blank). The size description given here for isolate Mv is consistent with the findings of Trench and Blank. While the algae isolated for that study were isolated from planulae of Montipora Figure 11: Examples of the dinokaryon nucleus in 5 isolates of Symbiodinium. A. The dinokaryon nucleus of isolate Y109 located centrally near an accumulation body. B. An example of the nucleus from FLAp4. C. An image of the nucleus from Ap highlighting the structure of the chromosomes. D. The dinokaryon nucleus from CCMP421 is also centrally located. E. The nucleus in isolate Mv is likewise centrally located and surrounded here by starch granules. All scale bars are 1?m. 81 verrucosa and the algae originally isolated and in culture for this and all other studies were isolated from a branch of the adult; it seems likely that Symbiodinium kawagutii described by Trench and Blank is the same type as those identified as S. kawagutii (Kinzie et al., 1984; Kinzie et al., 200) or Mv (Santos et al., 2001; LaJeunesse, 2001; Santos et al., 2002; Pochon et al., 2006) in subsequent studies. The cellular contents described here are also consistent with the original formal species description of Symbiodinium kawagutii. S. kawagutii has been examined for its growth properties under different wavelengths of light, growing better under blue or white light than under green or red lights (Kinzie et al., 1984). This species was also identified as a thermally tolerant Symbiodinium species exhibiting better growth at 29? and 31? C than other Symbiodinium strains in culture (Kinzie et al., 2001). Genetic studies have also investigated isolate Mv, this isolate was used as an outgroup from clade C in a study examining clade B strains cultivated from a variety of different host animals (Santos et al., 2001). This isolate was then identified as a strain in clade F1 using the ITS sequences (LaJeunesse, 2001). Using the cp23S-rDNA sequence along with sequences from the 18S-rDNA, n28S-rDNA, and ITS-rDNA genes this isolate was placed in Clade F (Santos et al., 2002). Later genetic examination of more cultures from clade F using the n28S-rDNA and cp23S-rDNA sequences has determined that this isolate is within clade F5 (Pochon et al., 2006). While most of the cellular structures are similar between different isolates of Symbiodinium, understanding the differences can help us both in understanding phylogenies and in understanding the ecology of these dinoflagellates. It is noteworthy that the size of the different isolates determined by our work is consistent with previous studies, some done many years ago on the same cultured isolate. This indicates that while we cannot state that the isolates do not change in culture we can state that cell size is a consistent characteristic that can be useful 82 in identification. It is also interesting that here and in the one previous description of a free- living Symbiodinium isolate the cell size is typically larger than symbiotic Symbiodinium isolates (Hansen and Daugbjerg, 2009). While the size and spacing of thylakoid membranes and the cell membrane and cell wall do not appear to be useful characters for identification, in this study we have provided an analysis of these characteristics for 5 isolates that any future work can use for comparison. The evidence presented here strongly suggests that there are not sufficient Figure 12: Examples of the mitochondrion structure in 5 isolates of Symbiodinium. A. The mitochondrion in Y109 are clustered and tubular cristae are located in the periphery of each. B. Isolate FLAp4 also contains mitochondria with tubular cristae. C. The mitochondria seen here in Ap is elongated with the characteristic peripheral tubular cristae seen in dinoflagellates. D. An example of a mitochondrion from CCMP421 with tubular cristae. E. Mv contains this elongated mitochondria with tubular cristae. Scale bars for panels A, B, D, and E are 1?m. Scale bar in panel C is 500nm. 83 ultrastructural differences amongst isolates to allow for identification of species or clade. Identification of isolates as members of a clade or species will have to rely solely on molecular identification methods. Despite the inability to use ultrastructural features for taxonomic identification, understanding of the ultrastructure of the symbiotic partners may suggest directions for a better understanding of the physiology of these symbioses as it relates to this extremely diverse genus of symbionts. 84 IV. ULTRASTRUCTURAL AND BIOCHEMICAL VARIATION AMONGST SYMBIODINIUM ISOLATES DEMONSTRATES VARIATION IN STORAGE MOLECULES ABSTRACT Symbiodinium is a diverse genus of unicellular dinoflagellates that often reside intracellularly in a variety of invertebrate hosts. Despite what is known about the genetic diversity of the genus, the physiological, morphological, and biochemical variation among members of this diverse group is poorly understood. One important area of research is determining how the symbiont establishes and maintains an association with a cnidarian host. Here we have looked at storage products within 5 isolates to determine any variation in the amount of lipid droplets and starch granules. Amongst the 5 isolates examined, the two smaller isolates from clade A contained the least amount of both starch and lipid. However, another small isolate from clade B contained some of the largest amounts of both starch and lipid. A clade F isolate also contained many storage products while the only free-living isolate examined, a clade E isolate, contained more lipid and starch than the clade A symbionts but less than the clade B and F isolates. Examination of exudate for the presence of carbohydrate and protein components revealed that in all 5 isolates, more carbohydrate than protein was secreted into the culture medium. Examining the cell wall associated compounds that may be involved in initiation and/or maintenance of the symbiosis, it was determined that a previously investigated antigen, PC3, is composed of carbohydrate structures and a probable protein component. This is consistent with the idea that a glycan/lectin interaction may be involved in the initiation and perhaps maintenance of this symbiosis. 85 INTRODUCTION Symbiodinium was originally described as a single pandemic species (Freudenthal, 1962), but has since been found to be very diverse based upon molecular homology. Through genetic techniques this genuswas separated into 9 clades (Rowan and Powers, 1991a; Carlos et al., 1999; LaJeunesse and Trench, 2000; LaJeunesse, 2001; Pochon et al., 2001; Santos et al., 2002; Pochon et al., 2004; Pochon and Gates, 2010). Additionally, ultrastructural (Taylor, 1968; Kevin et al., 1969; Bishop et al., 1976; Leutenegger, 1977; Schoenberg and Trench, 1980b; Van Thinh et al., 1986; Blank, 1987; Ambariyanto and Hoegh-Guldberg, 1996; Muller-Parker et al., 1996; Wakefield et al., 2000), physiological (Warner et al., 1996; Iglesias-Prieto and Trench, 1997; Rowan et al., 1997; Kinzie et al., 2001; Perez et al., 2001; Baker et al., 2004; Bhagooli and Hidaka, 2004; Rowan, 2004), and biochemical (Schoenberg and Trench, 1980a; Markell et al., 1992; Markell and Trench, 1993; Markell and Wood-Charlson, 2010) studies revealed differences amongst isolates within this genus. This diversity is important because it may allow for many potential combinations of host species and symbionts. Thus, some combinations of host and symbiont may be better adapted for specific environmental conditions, such as increases in sea surface temperature that can cause detrimental bleaching events, than others (Buddemeier and Fautin, 1993; Hoegh-Guldberg, 1999; Fitt et al., 2001; Rowan, 2004; Weis, 2010; Stat and Gates, 2011). The basis for this symbiosis is a nutritional one, meaning that the symbiont donates photosynthetically fixed carbon as a nutrient for the host while the host donates its waste products (such as CO2 and nitrogenous compounds) which are nutrients for the symbiont (Muscatine and Porter, 1977; Muscatine, 1990; Verde and McCloskey, 1998; Yellowlees et al., 2008). Both the type and the amount of translocated photosynthetically fixed carbon 86 compound(s) are important in determining whether the symbiont is mutualistic or veering into a parasitic lifestyle. Previous studies have shown that Symbiodinium isolates can contribute between 20% (Paylthoa townsleyi; Trench, 1971a) and 169% (Cassiopea xamachana; Verde and McCloskey, 1998) of carbon-based host metabolic needs. Thus, in many associations studied, the symbiont can transfer sufficient photosynthetically fixed carbon to provide for the hosts metabolic needs and often an excess above this level that can be used by the host for growth or reproductive efforts (Muscatine and McCloskey, 1981; Muscatine et al., 1984; Stambler and Dubinsky, 1987; Davy and Cook, 2001; Grottoli et al., 2006; Tanaka et al., 2006; Loram et al., 2007; Gordan and Leggat, 2010). While there are many studies looking at the amount of photosynthetically fixed carbon, fewer studies have examined the specific molecules translocated. Initial work indicated that the primary carbon compound produced is glycerol (Muscatine, 1967), but later investigations revealed that while glycerol was the major translocation product, other compounds such as alanine, glucose, fumaric acid, succinic acid, and glycolyic acid were also translocated(Trench, 1971b). Another study confirmed that glucose and succinate/fumarate were major transfer products in Anemonia viridis (Whitehead and Douglas, 2003). While there is some information about the compounds and amount of photosynthetically fixed carbon transferred to the host, there is less known about the pathway(s) by which the photosynthate moves between symbiont and host. Symbiodinium in culture produces a mucilaginous exudate (Markell et al., 1992; Markell and Trench, 1993) that in situ appears as a matrix that contains multiple layers of membranous material derived from the symbiont (Wakefield and Kempf, 2001). In culture the exudate is at least in part water soluble (and therefore present in the culture medium) and analysis of the culture medium and of extracted cell 87 wall indicate that it is composed of glycosylated polypeptides (Markell et al., 1992); it is usually composed of more carbohydrate than protein (Markell and Trench, 1993). These substances could be involved in recognition between host and symbiont, consistent with lectin-glycan recognition systems in other organisms and with lectin-like compounds found in cnidarian hosts (Kvennefors et al., 2008; Jimbo et al., 2013). Some of these carbohydrate containing compounds may be involved in recognition (Logan et al., 2010; Wood-Charlson et al., 2006) and/or host settlement cues (Markell and Wood-Charlson, 2010). Additionally, these compounds may be involved in translocation of photosynthetically fixed carbon from the symbiont to the host since some of the same compounds identified as components of the carbon translocated to the host are present in the exudate (Trench, 1971b; Markell and Trench, 1993). While it seems that much of the photosynthate is translocated to the host, storage products have been seen in ultrastructural studies of Symbiodinium. In the initial ultrastructural description, starch granules were not mentioned, but lipid droplets and vacuoles containing calcium oxalate were mentioned as inclusions (Kevin et al., 1969). In a 1968 study of symbiotic dinoflagellates the accumulation body containing mucopolysaccharide was identified along with the presence of starch grains throughout the cytoplasm and surrounding the pyrenoid (Taylor, 1968). Lipid droplets, also present in the symbiont's cytoplasm, have been well studied (Bishop et al., 1976) and found to mostly be galactosyldiacylglycerols (50%). Several later studies described symbionts from different hosts with the same inclusions as these earlier studies (Schoenberg and Trench, 1980; Van Thinh et al., 1986; Blank, 1987; Wakefield et al., 2000). In an ultrastructural examination of starch inclusions in normal and well-fed (N-enriched) symbioses between Tridacna maxima and its symbiont there was an increase in starch under normal conditions possibly due to N limitation on the reef (Ambariyanto and Hoegh-Guldberg, 88 1996). This is probably due to the carbon stores being utilized for greater amino acid synthesis and therefore, protein synthesis, under nitrogen enriched conditions (Ambariyanto and Hoegh- Guldberg, 1996). Likewise, an examination of symbionts within the host compared to those in culture revealed an increase in starch in cultured symbionts, again possibly due to N limitation in culture compared to within the host (Schoenberg and Trench, 1980). Lipid droplets behaved similarly and in well-fed anemones, Symbiodinium lipid content decreased from 30% to 15% (Muller-Parket et al., 1996). While there is some information about the starch grains and lipid droplets, there have been no comparisons of this content amongst different Symbiodinium isolates under the same environmental conditions. It is important to understand how the Symbiodinium-cnidarian association is initiated and maintained in order to recognize whether, during bleaching episodes, the host may be able to acquire new symbiont strains or switch to different symbiont strains that may be better suited for the new stress-inducing environmental conditions (the ?adaptive bleaching hypothesis?; Buddemeier and Fautin, 1993). We can then better understand the possibility for survival of some associations as sea surface temperatures increase (Baker et al., 2004; Stat and Gates, 2011). The fixation and donation to the host of photosynthetic carbon is vital to the maintenance of the association and the coral reef ecosystem. Here we examine the presence of storage molecules of five Symbiodinium isolates to see if there is variation between isolates that may correspond to physiological differences that could alter the efficiency of a particular host-symbiont association. Additionally, we examine the mucilaginous exudate for protein and carbohydrate composition to see if there is a correlation between the amount of stored starch and the compounds secreted to the exterior of the cell. 89 METHODS CULTURE OF SYMBIODIMIUM ISOLATES. Cultures were grown in F/2 media (Guillard and Ryther, 1962) in 125mL Erlenmeyer flasks under 12/12 light dark cycle at 24.5?C ? 0.5? at Auburn University for 2.5 weeks. Cultures were then transported to the culture room of Dr. William Fitt at the University of Georgia and allowed to grow for 2 weeks under a 12/12 light dark cycle at 26.5?C ? 0.5?. Cultures examined ultrastructurally in this investigation are listed in Table 1. HIGH PRESSURE RAPID FREEZING. Fixative solutions were made up several days prior to the freezing event. 0.1% Uranyl acetate in HPLC grade acetone was mixed at room temperature 2 days prior to freezing and the day before freezing was transferred to a -80?C freezer. A vial of 25 mL of HPLC grade acetone was pre-chilled in the -80?C freezer overnight and then 1 g of OsO4 was added (making a 4% OsO4 solution) while the solution was kept on dry ice for 10 minutes before being replaced in the -80?C freezer the day before freezing. Isolate samples were collected around 'dawn' by pipetting 100?L of cells and media using a Gilson pipetman starting at the bottom of the flask (where the cells accumulate) and slowly drawing the pipette up about 1 cm from the bottom. The sample was then transferred to a 1.5 ml microcentrifuge tube and centrifuged briefly to produce a loose pellet. Most of the remaining F/2 media was removed (leaving the cell pellet in about 50 ?L of F/2 media) and then 50 ?L of 20% dextran in F/2 media was added and the cells were re-suspended in a final concentration of 10% dextran in 100 ?L of F/2 medium. The 10% dextran encouraged the cells to remain clumped together after freezing. 5?L of cells were transferred to planchettes using a Gilson pipetman and then rapidly frozen in a High Pressure Freezing Machine (HPM 010 ABRA Fluid AG). After freezing, pellets were transferred to cryotubes submerged in liquid nitrogen and then 90 transferred to a pre-cooled dewer and maintained in liquid nitrogen until transfer into fixative. The 0.1% Uranyl acetate in acetone and 4% OsO4 in acetone were then mixed (all vials kept on dry ice) in a 1:1 ratio and aliquoted into enough vials for the frozen samples. After 2 hours in the -80?C freezer, the frozen samples in planchettes were quickly added to the vials of 1:1, 0.1% Uranyl acetate: 4% OsO4 in acetone, all vials and cryotubes being kept on dry ice. Samples in fixative were then placed in the - 80?C freezer for 4 days. Samples were next moved to the -20?C freezer for 3 hours, then to a 4?C refrigerator for 2 hours, and finally into the fume hood at room temperature for 30 minutes. After this gradual return to room temperature the fixative was removed and samples were rinsed 3 times in 100% HPLC grade acetone for 15 minutes each. Samples were then removed from planchettes and embedded using a Low viscosity Embedding Media Spurr's Kit (hard recipe; EMS) by first adding the embedding resin as a 1:2 mix in acetone and leaving it overnight at room temperature. The next day the samples were changed to 2 parts embedding resin to 1 part acetone for 8 hours and then into 100% resin overnight. The next morning samples were placed in fresh 100% resin for 8 hours. Near the end of the day the samples were transferred to catalyzed resin in Beem capsules. After 1 hour at room temperature in the catalyzed resin the samples were placed in an oven at 60?C and left for 24-48 hours to ensure hardening. Table 1: Information about the isolates of Symbiodinium studied here. Species Culture ID Host Clade (Cp23S rDNA) Avg Diameter Symbiodinium sp. Y109 Unknown host A191 9.0 ?0.83 ?m, n=30 Symbiodinium sp. FLAp4 Aiptasia pallida A193 7.5 ?0.6 ?m, n=30 Symbiodinium pulchrorum HIAp or Ap Aiptasia pulchella B224 8.0 ? 0.86 ?m, n=30 Symbiodinium sp. CCMP 421 None E202 Symbiodinium kawagutii Mv Montipora verrucosa F178 9.1 ? 0.74 ?m, n=30 91 TRANSMISSION ELECTRON MICROSCOPY. Sections were cut using a Reichert-Jung Ultracut E Microtome and placed on copper 75 mesh grids (EMS). They were then stained using 4% aqueous solution of uranyl acetate for 10 to 20 minutes, rinsed and then secondarily stained with 0.1% aqueous solution of lead citrate for 5 minutes. Samples were viewed on a Zeiss EM 10C 10CR Transmission Electron Microscope and digital images were taken using MaxIm DL5 software and an SIA-L3C 4.3 megapixel camera (Scientific Instruments & Applications, Duluth, GA 30096). ULTRASTRUCTURAL MEASUREMENTS AND STATISTICS. In order to obtain accurate measurements of cell size, only cells larger than 4 ?m in diameter were measured and added to a list to obtain 100 measured cells from each isolate. These cells were then sorted by size and only the largest 30% (n=30) were used to determine percent coverage of lipid and starch inclusions. This was done to ensure that comparisons were always made between largest cross sections and therefore approximately in the middle of each cell. Measurements of cell diameter were made first using the MaxIMDL5 software and then confirmed using Image-Pro Plus (Version 7.0; Media Cybernetics, 2009). Measurements of cell cross-sectional area and starch/lipid cross- sectional area were made using Image Pro Plus (Version 7.0; Media Cybernetics, 2009). The percent of lipid and starch cross-sectional area was determined by dividing the total cross- sectional area covered by either starch or lipid by the entire cell cross-sectional area and multiplying by 100. Lipid and starch inclusion differences among isolates were transformed using an arcsin transformation and then compared using a one-way ANOVA run on SAS (Version 9.2) to ascertain any significant differences. Post-hoc multiple comparison tests were run with a Tukey-Kramer adjustment to examine which strains exhibited significant differences. 92 COLLECTION OF EXUDATE FROM SYMBIODINIUM CULTURES. Symbiodinium isolates for exudate analysis were grown as described above at Auburn University for 4-5 weeks (until they were in the log phase of growth). 300?L of culture medium plus cells were then collected and placed in a microcentrifuge tube along with 300?L of a 45ppt salt solution. The samples were next centrifuged at high speed in a clinical centrifuge for 10 minutes. After centrifugation, the supernatant was collected and used for further analysis. Cell numbers were determined using a hemacytometer. BIOCHEMICAL ANALYSIS OF EXUDATE. 200?L of supernatant containing exudate was transferred to a clean microcentrifuge tube (Tube 1). To separate the carbohydrate and protein components, 100?L of 15% Trichloracetic acid (TCA) was added and the microcentrifuge tube was vortexed for 5 minutes followed by 10 minute incubation at 4? C and then centrifugation at 5000g for 10min. The supernatant was then transferred to another clean tube (Tube 2). 200?L of 5% TCA was then added to Tube 1 from which the supernatant had just been removed. This was vortexed as above, centrifuged for 5 minutes, and then the resulting supernatant was added to Tube 2 (now containing approximately 500?L of carbohydrate containing supernatant). 700?L of 1.0N NaOH in 1% Na-deoxycholate was added to Tube 1 which now had the carbohydrate containing supernatant removed and a protein pellet remaining. The tube was vortexed for 4 minutes, heated to 56? C for 30min, and centrifuged for 10 minutes. During this time BSA standards were prepared. The rest of the protein determination was carried out according to the Lowry method (Lowry et al., 1951). In order to determine the amount of carbohydrate present in the exudate, 200 ?L of supernatant from Tube 2 (see separation of carbohydrate and protein above) was transferred, to each of 2 new 1.5 ml microcentrifuge tubes. Next 20?L of 6.0N HCl was added to each 93 microcentrifuge tube. The tubes were then vortexed followed by heating for 2 hours at 95?C. After the 2 hours at 95?C, the following was used to determine the amount of carbohydrate in each sample (modified from Holland and Gabbot, 1971). 1. The tube was cooled for 5 minutes and centrifuged at 5000g for 2 minutes. 2. 20?L of 6.0N NaOH and 60?L of 5% TCA were added followed by additional vortexing. 3. 100?L of this solution was transferred to each of two glass reaction tubes along with 100?L of 5% TCA. 4. At this point glucose standard tubes were prepared containing 0, 2.5, 5, 10, 12, and 16 ?g of glucose. 5. To both the reaction tube and the glucose standard tubes, 500?L of 0.625% NaOH, 100?L of 0.2% potassium ferricyanide, and 200?L of the carbonate-cyanide reagent (500?L 16% NaCO3 + 30?L 5%KCN + 9.5 nanopureH2O) were added, vortexing in between each addition of a solution. 6. The reaction and standards tubes were heated at 95?C for 10 minutes and then cooled for 10 minutes. 7. 500?L of npH2O was added to each tube and vortexed. 8. The absorbance was read at 420nm on a Genesys 2 Spectrophotometer. The absorbance values for the glucose standards were used to prepare a standard absorbance versus total mass curve. The amount of carbohydrate in each reaction tube was determined from the standard glucose curve and the average value for the two reaction tubes analyzed was used to determine the amount of carbohydrate present in the volume of the original sample. 94 ENZYMATIC EXAMINATION OF EXUDATE. 200?L of cells of isolate FLAp4 were collected at dawn, centrifuged, media removed, and then dehydrated and rehydrated through PBS+, 30% ethanol, 50% ethanol, 70% ethanol, 50% ethanol, 30% ethanol, PBS+ centrifuged between each step to remove chlorophyll autofluorescence. Then each of 9 pellets was treated in Figure 1: Examples of cells from 5 Symbiodinium isolates containing the average cross-sectional percent area starch granules present in that isolate. A. Isolate Y109 contains 9.45% ? 5.5% cross-sectional area composed of starch. B. 8.82% ? 6.3% of cross-sectional area is starch in isolate FLAp4. C. Isolate Ap is composed of 16.84% ? 4.6% cross- sectional area of starch. D. 12.05% ? 5.2% cross-sectional area is composed of starch in isolate CCMP421. E. Isolate Mv contains 19.99% ? 6.7% cross-sectional area composed of starch. Scale bars = 2?m. 95 a different way. Treatments were always suspended in 200?L of the solution mentioned and included (1) incubation with 25 units of trypsin in 20mM phosphate buffered saline containing 0.1% sodium azide and 0.1% Triton X100 (PBS+) for 1 hour at 37?C; (2) incubation with 25 units of neuraminidase in PBS+ for 1 hour at 37?C; (3) incubation with 25 units of hyaluronidase in PBS+ for 1 hour at 37?C; (4) incubation with 25 units of chymotrypsin in PBS+ for 1 hour at 37?C; (5) media removal via centrifugation, resuspension in PBS+ and homogenization using the VirTis hand-held homogenizer; (6) incubation with 1% periodic acid in nanopure water for 1 hour; (7) incubation in nanopure water and boiled for 5 minutes; (8) incubated at 37?C for 1 hour in PBS+ (control + heat); (9) kept at 4?C for 1 hour (control). All treatments were then divided into 2 1.5mLmicrocentrifuge tubes, centrifuged at 2000g, supernatant removed and incubated 5% heat-inactivated goat serum in PBS+ (blocking medium) for 10 minutes at room temperature. Blocking medium was removed by centrifugation and 1 tube of each treatment was incubated with spent medium containing primary antibody PC3 (an antibody specific to the outer coat of Symbiodinium of isolate FLAp4) diluted 1:1 in blocking medium for 1 hour at room temperature (the second tube of each strain was incubated with only blocking medium for this hour as a negative control). Samples were then washed in PBS+ 3 times for 5 minutes each, followed by incubation in secondary antibody (Alexa fluor 488 Goat anti-mouse IgM (? chain) - Invitrogen, A-21042) diluted 1:300 in blocking medium for 30 minutes at room temperature in darkness (to prevent quenching). Samples were then rinsed 3 times in PBS+ for 5 minutes each and resuspended in a small amount of Nanopure-H2O (np-H2O). Cells were then put on slides and viewed on a Nikon Optiphot fluorescence microscope and imaged with a Canon Eos Rebel T2i camera. A minimum of 150 cells of each treatment were examined for the presence of fluorescence indicating that the antigen remained intact. Percent fluorescence data was 96 transformed using an arcsin transformation and then examined with an ANOVA using SAS 9.2 for statistical significance. Post-hoc multiple comparison tests were run with a Tukey-Kramer adjustment to examine which strains exhibited significant differences. Figure 2: Graph of statistical analysis of the percent cross- sectional area of starch granules (A) and lipid droplets (B). Error bars indicate the standard error. Symbols above bars indicate isolates that are not significantly different from each other. P<0.0001 97 RESULTS ULTRASTRUCTURAL STARCH VARIATION. Starch content varied dramatically and significantly amongst 5 different Symbiodinium isolates (Figure 1, 2). Isolates FLAp4 (8.82% ? 6.3%, n=30 cells, p <.0001), Y109 (9.45% ? 5.5%, n=30 cells, p <.0001 and CCMP421 (12.05% ? 5.2%, n=30 cells, p <.0001) were not significantly different from each other but were significantly different from isolates Ap (16.84% ? 4.6%, n=30 cells, p <.0001) and Mv (20.0% ? 6.7%, n=30 cells, p <.0001). ULTRASTRUCTURAL LIPID VARIATION. Lipid content varied amongst the 5 Symbiodinium isolates examined. The lipid droplets are identified as gray inclusions (Dodge and Greuet, 1987) within the cell cytoplasm (Figure 3). The average percent of lipid varied significantly amongst isolates (Figure 2). Isolate FLAp4 (6.20% ? 6.7%, n=30 cells, p < .0001) was not significantly different from Y109 (10.93% ? 7.13%, n=30 cells, p < .0001), but was significantly different from the other 3 isolates. Isolate CCMP421 (13.20% ? 9.8%, n=30 cells, p < .0001) was not significantly different from Y109 or Mv (17.07 ? 8.32%, n=30 cells, p < .0001), but was significantly different from both FLAp4 and Ap (19.24 ? 8.55%, n=30 cells, p < .0001). BIOCHEMICAL ANALYSIS OF EXUDATE. Examination of the exudate from isolates of Symbiodinium used in this study revealed that in all cases the exudate contained both protein and carbohydrate components (Table 2). In every isolate, cells secreted more carbohydrate than protein. There was no significant difference between isolates in the amount of either protein or carbohydrate present in the exudate. ENZYMATIC DIGESTION OF EXUDATE. Incubating cells of isolate FLAp4 with various enzymes showed a slight (but not significant) decrease in labeling with the PC3 antibody 98 for the enzymes trypsin, and hyaluronidase (Figure 4). Neuraminidase showed a significant decrease in number of cells that labeled with PC3 antibody (Figure 4; p=0.03) indicating the presence of glycosidic linkages. There was also a significant decrease in antibody labeling in treatments with 1% periodic acid and boiling (Figure 4; p<0.0001). Figure 3: Examples of cells from 5 Symbiodinium isolates containing the average percent cross-sectional percent coverage of lipid droplets. A. Isolate Y109 contains 10.93 ? 7.13% cross-sectional area of lipid. B. Isolate FLAp4 contains an average of 6.20 ? 6.7% cross-sectional area composed of lipid. C. 19.24 ? 8.55% of cross- sectional area is composed of lipid in isolate Ap. D. Isolate CCMP421 contains 13.20 ? 9.8% cross-sectional area of lipid. E. Isolate Mv contains 17.07 ? 8.32% cross-sectional area composed of lipid. Scale bars are all 2?m. 99 DISCUSSION Since the symbiosis between Symbiodinium and its cnidarian hosts is nutritionally based, the production, storage, and excretion of photosynthetically fixed carbon by the symbiont is vital to the understanding of the association. Any variation in production, storage or excretion of photosynthetically fixed carbon could lead to differences in the potential for a successful association between a specific host and a specific Symbiodinium isolates. Isolates that can either photosynthesize more efficiently or translocate a higher percentage of their photosynthetically fixed carbon to the host should be better partners for the host (Muscatine et al., 1984). Here we have examined the presence of storage products in 5 isolates of Symbiodinium. Isolates Ap and Mv contain significantly more starch granules than other isolates. Interestingly, isolate Mv (a clade F5 isolate; Pochon et al., 2006) was identified as a thermally tolerant strain with increased growth rates at 29? and 31?C (Kinzie et al., 2001). Conversely, isolate Ap showed decreased thermal tolerance (Kinzie et al., 2001), but under the culture conditions here stored similar amounts of starch granules. Table 2: Biochemical Analysis of Exudate. Isolates Protein (per 1,000,000 cells) Carbohydrate (per 1,000,000 cells) Y109 0.133 ? 0.056 ?g n=14 1.632 ? 0.775 ?g n=8 FLAp4 0.149 ? 0.086 ?g n=14 3.785 ? 1.532 ?g n=8 Ap 0.087 ? 0.032 ?g n=10 2.123 ? 1.030 ?g n=8 CCMP421 0.093 ? 0.042 ?g n=8 2.691 ? 1.173 ?g n=10 Mv 0.339 ? 0.083 ?g n=14 2.405 ? 1.142 ?g n=10 100 Isolates FLAp4, Y109 and CCMP421 all contained less starch. Interestingly, both FLAp4 and Y109 are isolates from clade A that tend to be somewhat smaller (especially FLAp4) than all of the other studied isolates (Chapter 3, pg 66). Since statistics were performed on a percent of total cross-sectional area, the small cell size should not impact this conclusion. While there is little known about these isolates physiologically, a variety of different isolates within clade A have exhibited traits of both thermal tolerance and thermal susceptibility (Robison and Warner, 2006; Diaz-Almeyeda et al., 2011; Steinke et al., 2011). Without knowing about the thermal tolerance of isolates Y109 and FLAp4 we cannot make any suggestions about the connection between lower amounts of starch and thermal conditions. Isolate CCMP421 is a free- living isolate of Symbiodinium and the only other free-living isolate described is Symbiodinium natans (Hansen and Daugbjerg, 2009). S. natans has been reported to contain lipid and starch inclusions, but, unfortunately, only a qualitative description was given. Storage of lipid droplets also showed significant variation amongst the Symbiodinium isolates investigated. Based on our % area calculations, isolate Ap contained the greatest amount of lipid (significantly different from clade A isolates FLAp4 and Y109). This isolate is a thermally intolerant member of clade B. In previous studies, Symbiodinium in nitrogen starved conditions tended to store more lipid droplets and starch granules (Ambariyanto and Hoegh- Guldberg, 1996; Muller-Parker et al., 1996). The differences seen here occur in identical, healthy, nutrient rich cultures which suggest that they are due to physiological differences among the isolates tested. Thus, it is likely that isolate Ap has different lipid metabolism requirements than isolates FLAp4 and Y109. Isolate Mv also contained a large amount of lipid (17%) that was significantly different from the amount present in FLAp4, as well as a large amount of starch. Since Mv is known to be thermally tolerant (Pochon et al., 2006) it would be interesting to 101 examine whether the large amounts of storage product accumulated relate to its ability to withstand higher temperatures. Isolate CCMP421 contained 13.2 ? 9.8% lipid and 12.1 ? 5.2% starch. Perhaps the relatively high amount of both types of storage products seen in the free-living strain indicates that it would not be as efficient at translocation of photosynthate under symbiotic conditions. As a free-living isolate there is essentially no need to translocate photosynthate and thus such carbon may be channeled into storage products. This would be consistent with the life history of this isolate and with previous descriptions of free-living isolates that report a number of inclusions (Hansen and Daugbjerg, 2009), but very little in the way of excreted carbon compounds (as evidenced by a lack of labeling with lectin probes; Logan et al., 2010). Clade A isolates FLAp4 Figure 4: Number of cells of isolate FLAp4 exhibiting fluorescence after a variety of different treatments. 1%PA = 1% Periodic acid, boil was boiled for 5 minutes, neur = neuraminidase treatment, PBS+ = homogenized in PBS+, Tryp = trypsin treatment, Chymo = chymotrypsin treatment, C = Control, C + H = Control with the same heat treatment as the enzymes. Identical symbols above the bars indicate treatments that are not significantly different from one another. 102 and Y109 again had fewer lipid droplets (FLAp4 significantly fewer than CCMP421, Mv and Ap; Y109 significantly fewer than Ap). Isolates FLAp4 and Y109 are both physically smaller (FLAp4 especially) however, little is known of their physiology. Clade A contains members that are both thermally tolerant and intolerant, however, there is no information about the thermal tolerance of these two isolates in particular. The fewer inclusions in these strains could mean that they have different nutrient requirements than the other isolates or that they do not store food reserves as efficiently. As above, the differences in amount of stored lipid probably reflect differences in the physiology of these isolates. Such differences could have important consequences for the efficiency of translocation and possibly thermal and light tolerance as well as for the suitability of specific isolates as symbionts for specific hosts. While more needs to be understood about the physiology of these isolates, differences in the production and storage of lipid and/or starch could be important factors in maintaining a successful association with specific hosts or under specific environmental conditions. While the storage of products derived from photosynthetically fixed carbon may have implications for the health of the association between Symbiodinium and their host cnidarian, it is also likely that any compounds exuded by the symbiont are important in the establishment and/or the maintenance of the association. Here we saw that all 5 isolates exuded some protein and carbohydrate into the culture media. This is consistent with previous work that found a variety of compounds exuded into the media, including protein, neutral sugars, glucosamine, galactosamine, and uronic acid (Markell and Trench, 1993). There could be differences in the amounts of secreted compounds and it is likely that the components in the exudate would vary amongst Symbiodinium isolates. 103 These secreted compounds are similar to the compounds others have noted as being associated with the cell wall of Symbiodinium (Markell et al., 1992; Lin et al., 2000). It has been suggested that cell wall associated glycoproteins may be involved in cell-cell recognition during the establishment of the symbiosis (Lin et al., 2000). Such being the case, the glycans that are associated with the symbiont cell wall would interact with lectins associated with the cnidarian host cell to initiate the symbiosis (Wood-Charlson et al., 2006; Jimbo et al., 2012). Some lectins have been identified in cnidarian hosts including a mannose-binding lectin in Acropora millepora (Kvennefors et al., 2008) and a D-galactose-binding lectin from Sinularia lochmodes (Jimbo et al., 2000). The glycans identified thus far have shown no pattern relative to clade, but regardless of Symbiodinium clade, isolates from the same host have shown similar profiles (isolates from clade A and B from the same host had similar profiles; Logan et al., 2010). Logan found that all 8 isolates tested had exudates containing glycans with mannose residues while 6 out of 8 isolates contained glycans with N-acetyl groups (2010). Other glycans were present in smaller concentrations and associated with fewer isolates and it is possible that these play an important role in specificity (Logan et al., 2010). Here examination of an antigen present in the exudate of Symbiodinium isolate FLAp4 (as well as some other isolates; Chapter 2, pg 47) that is recognized by the PC3 antibody (Wakefield and Kempf, 2001) also showed that binding of the antibody is lessened when chemicals that damage the structure of glycans are added (such as periodic acid, which showed a very strong significant effect; and neuraminidase which also showed significant effects). The variation seen here in the storage of lipid and starch is likely due to physiological differences amongst isolates that could impact the transfer of photosynthetically fixed carbon and therefore the host-symbiont association. The biochemical analysis of exudate from a number of 104 solates is consistent with previous work (Markell and Trench, 1993; Markell and Wood- Charlson, 2010). Also, our examination of the cell wall associated antigen PC3 (Wakefield and Kempf, 2001) indicated that it is composed of both carbohydrate and protein components which agrees with cell wall associated compounds examined in previous studies (Markell et al., 1992; Lin et al., 2000; Wood-Charlson et al., 2006; Kvennefors et al., 2008; Logan et al., 2010; Jimbo et al., 2012). These results are consistent with the idea that cell surface associated compounds are exuded by isolates of Symbiodinium and could be involved in both the initiation and the maintenance of the Symbiodinium-cnidarian association. 105 REFERENCES Abrego, D., van Oppen, M.J.H., & Willis, B.L. 2009a. Highly infectious symbiont dominates initial uptake in coral juveniles. Mol. Ecol. 18:3518-3531. Abrego, D., van Oppen, M. J. H., & Willis, B. L. 2009b. Onset of algal endosymbiont specificity varies among closely related species of Acropora corals during early ontogeny. Mol. Ecol. 18:3532-3542. Ambariyanto, & Hoegh-Guldberg, O. 1996. Nutrient enrichment and the ultrastructure of zooxanthellae from the giant clam Tridacna maxima. Mar. Biol. 125:359-363. Backhed, F., Ley, R. E., Sonnenburg, J. L., Peterson, D. A., & Gordon, J. I. 2005. Host-bacterial mutualism in the human intestine. Science. 307: 1915-1920. Baker, A. C., Romanski, A. M. 2007. Multiple symbiotic partnerships are common in scleractinian corals, but not in octocorals: Comment on Goulet (2006). Mar Ecol Prog Ser. 335:237-242. Baker, A.C., Rowan, R., & Knowlton, N. 1997. Symbiosis ecology of two Caribbean acroporid corals. Proc. 8th Int Coral Reef Sym. 2:1295-1300. Baker, A.C., Starger, C.J., McClanahan, T.R., & Glynn, P.W. 2004. Corals' adaptive response to climate change. Nature. 430:741. Baillie, B. K., Belda-Baillie, C. A. & Maruyama, T. 2000b. Conspecificity and Indo-Pacific distribution of Symbiodinium genotypes (Dinophyceae) from giant clams. J. Phycol. 36:1153-1161. Baillie, B. K., Belda-Baillie, C. A., Silvestre, V., Sison, M.V., Gomez, E. D., & Monje, V. 2000a. Genetic variation in Symbiodinium isolates from giant clams based on random- amplified-polymorphic DNA (RAPD) patterns. Mar. Biol. 136:829-836 Baillie, B. K., Monje, V., Silvestre, V., Sison, M., & Belda-Baillie, C. A. 1998. Allozyme electrophoresis as a tool for distinguishing different zooxanthellae symbiotic with giant clams. Proc. R. Soc. Lond. B 265:1949-1956. Banaszak, A.T., Iglesias-Prieto, R. & Trench, R.K. (1993). Scrippsiella velellae sp. nov. (Peridiniales) and Gloeodinium viscum sp. nov. (Phytodiniales), dinoflagellate symbionts of two hydrozoans (Cnidaria). J. Phycol. 29(4): 517 -528 Barbrook, A. C., Visram, S., Douglas, A. E., & Howe, C. J. 2005. Molecular diversity of dinoflagellate symbionts of cnidaria: the psbA minicircle of Symbiodinium. Protist. 157:159-171. 106 Barneah, O., Brickner, I., Hooge, M., Weis, V. M., & Benayahu, Y. 2007b. First evidence of maternal transmission of algal endosymbionts at an oocyte stage in a triploblastic host, with observations on reproduction in Waminoa brickneri (Acoelomorpha). Invert Biol. 126:113-119. Barneah, O., Brickner, I., Hooge, M., Weis, V. M., LaJeunesse, T. C., & Benayahu, Y. 2007a. Three party symbiosis: acoelomorph worms, corals and unicellular algal symbionts in Eilat (Red Sea). Mar Biol. 151: 1215-1223. Barneah, O., Weis, V. M., Perez, S., & Benayahu, Y. 2004. Diversity of dinoflagellate symbionts in Red Sea soft corals: mode of symbiont acquisition matters. Mar Ecol Prog Ser. 275:89-95. Barnes, D.J. & Chalker, B.E. 1990. Calcification and Photosynthesis in Reef-building Corals and Algae. In Dubinsky Z (ed) Ecosystems of the World 25: Coral Reefs. Elsevier Science, Amsterdam 109-113. Bay, L. K., Cumbo, V. R., Abrego, D., Kool, J. T., Ainsworth, T. D. & Willis, B. L. 2011. Infection dynamics vary between Symbiodinium types and cell surface treatments during establishment of endosymbiosis with coral larvae. Diversity. 3:356-374. Belda-Baillie, C.A., Baillie, B.K., & Maruyama, T. 2002. Specificity of a model cnidarian- dinoflagellate symbiosis. Biol Bull. 202:74-85. Benayahu, Y., Achituv, Y., & Berner, T. 1988. Embryogenesis and acquisition of algal symbionts by planulae of Xenia umbellate. Mar Biol. 100:93-101. Benayahu, Y., Achituv, Y., & Berner, T. 1989. Metamorphosis of an octocoral primary polyp and its infection by algal symbiosis. Symbiosis. 7:159-169. Berkelmans, R., De?ath, G., Kininmonth, S., & Skirving, W. J. 2004. A comparison of the 1998 and 2002 coral bleaching events on the Great Barrier Reef: spatial correlation, patterns, and predictions. Coral Reefs. 23:74-83. Berkelmans, R., & van Oppen, M.J.H. 2006. The role of zooxanthellae in the thermal tolerance of corals: a 'nugget of hope' for coral reefs in an era of climate change. Proc. R. Soc. Biol. Sci. Ser. B 273:2305-2312. Berner, T., Wishkovsky, A., & Dubinsky, Z. 1986. Endozoic algae in shelled gastropods ? a new symbiotic association in coral reefs? I. Photosynthetically active zooxanthellae in Strombus tricornis. Coral Reefs. 5:103-106. Bhagooli, R., & Hidaka, M. 2004. Photoinhibition, bleaching susceptibility and mortality in two scleractinian corals, Platygyra ryukyuensis and Stylophora pistillata, in response to thermal and light stresses. Comp. Biochem. Physiol A. 137: 547-555. 107 Bhuvaneswari, T., Pueppke, S., & Bauer, W. 1977. Role of lectins in plant-microorganism interactions: I. binding of soybean lectin to rhizobia. Plant Physiol. 60:486-491. Bishop, D. G., Bain, J. M., & Downton, W. J. S. 1976. Ultrastructure and lipid composition of zooxanthellae from Tridacna maxima. Aust. J. Plant Physiol. 3:33-40. Blank, R. J. 1987. Cell architecture of the dinoflagellate Symbiodinium sp. inhabiting the Hawaiian stony coral Montipora verrucosa. Mar. Biol. 94: 143-155. Blank, R. J., & Trench, R. K. 1985. Speciation and Symbiotic Dinoflagellates. Science. 229: 656-658. Brandt, K. A. H. 1885. Die koloniebildenden Radiolarien (Sphaerozoein) des Golfes von Neapel. Fauna und Flora des Golfes von Neapel XIII, Neapel. Breckels, M.N., Roberts, E.C., Archer, S.D., Malin, G., & Steinke, M. 2011. The role of dissolved infochemicals in mediating predator-prey interactions in the heterotrophic dinoflagellate Oxyrrhis marina. J. Plankton Res. 33:629-639. Buddemeier, R. W., & Fautin, D. G. 1993. Coral bleaching as an adaptive mechanism. BioScience 43: 320-326. Burris, R.H. 1983. Uptake and assimilation of 15NH4 by a variety of corals. Mar. Biol. 75:151- 155. Cantin, N. E., van Oppen, M. J. H., Willis, B. L., Mieog, J. C., & Negri, A. P. 2009. Juvenile corals can acquire more carbon from high-performance algal symbionts. Coral Reefs. 28:405-414. Carlos, A. A., Baillie, B. K., Kawachi, M., & Maruyama, T. 1999. Phylogenetic position of Symbiodinium (Dinophyceae) isolates from Tridacnids (Bivalvia), Cardiids (Bivalvia), a sponge (Porifera), a soft coral (Anthozoa), and a free-living strain. J Phycol. 35: 1054- 1062. Carlos, A. A., Baillie, B. K., & Maruyama, T. 2000. Diversity of dinoflagellate symbionts (zooxanthellae) in a host individual. Mar Ecol Prog Ser. 195: 93-100. Catt, J. W., Hills, G. J., & Roberts, K. 1976. A structural glycoprotein, containing hydroxyproline, isolated from the cell wall of Chlamydomonas. Planta. 131:165-171. Chen, C. A., Lam, K. K., Nakano, Y., & Tsai, W. S. 2003. A stable association of the stress- tolerant zooxanthellae, Symbiodinium Clade D, with the low-temperature-tolerant coral, Oulastrea crispate (Scleractinia: Faviidae) in subtropical non-reefal coral communities. Zool Stud. 42: 540-550. 108 Chen, M. C., Cheng, Y. M., Hong, M. C., & Fang, L. S. 2004. Molecular cloning of Rab5 (ApRab5) in Aiptasia pulchella and its retention in phagosomes harboring live zooxanthellae. Biochemical and Biophysical Research Communications. 324:1024- 1033. Chen, M. C., Cheng, Y. M., Sung, P. J., Kuo, C. E., & Fang, L. S. 2003. Molecular identification of Rab7 (ApRab7) in Aiptasia pulchella and its exclusion from phagosomes harboring zooxanthellae. Biochemical and Biophysical Research Communications. 308:586-595. Chen, M. C., Hong, M. C., Huang, Y. S., Liu, M. C., Cheng, Y. M., & Fang, L. S. 2005. ApRab11, a cnidarian homologue of the recycling regulatory protein Rab11, is involved in the establishment and maintenance of the Aiptasia-Symbiodinium endosymbiosis. Biochemical and Biophysical Research Communications. 338:1607-1616. Coffroth, M.A., Lasker, H.R., Diamond, M.E., Bruenn, J.A.,& Bermingham, E., 1992. DNA fingerprinting of a gorgonian coral: A method for detecting clonal structure in a vegetative species. Mar. Biol. 114:317?325. Coffroth, M. A., & Santos, S. R. 2005. Genetic diversity of symbiotic dinoflagellates in the genus Symbiodinium. Protist. 156:19-34. Coffroth, M.A., Santos, S.R., & Goulet, T.L. 2001. Early ontogenetic expression of specificity in a cnidarian-algal symbiosis. Mar. Ecol. Prog. Ser. 222:85-96. Colley, N.J., & Trench, R.K. 1983. Selectivity in phagocytosis and persistence of symbiotic algae by the scyphistoma stage of the jellyfish Cassiopeia xamachana. Proc. R. Soc. Lond. B Biol. Sci. 219:61-82. Damaskos, D., & Kolios, G. 2008. Probiotics and prebiotics in inflammatory bowel disease: microflora ?on the scope?. British Journal of Clinical Pharmacology. 65:453-467. Davy, S. K., & Cook, C. B. 2001. The relationship between nutritional status and carbon flux in the zooxanthellate sea anemone Aiptasia pallida. Mar Biol. 139:999-1005. Davy, S. K., & Turner, J. R. 2003. Early development and acquisition of zooxanthellae in the temperate symbiotic sea anemone Anthopleura ballii (Cocks). Biol Bull. 205: 66-72. Diaz-Almeyda, E., Thome, P. E., Hafidi, M. E., & Iglesias-Prieto, R. 2011. Differential stability of photosynthetic membranes and fatty acid composition at elevated temperature in Symbiodinium. Coral Reefs. 30: 217-225. Diekmann, O.E., Olsen, J.L., Stam, W.T., & Bak, R.P.M. 2003. Genetic variation within Symbiodinium clade B from the coral genus Madracis in the Caribbean (Netherlands Antilles). Coral Reefs. 22:29-33. 109 Dodge, J. D. 1966. The dinophyceae. In Godward, M. B. E. [Ed.] The Chromosomes of Algae. London. Dodge, J. D. & Greuet, C. 1987. Dinoflagellate ultrastructure and complex organelles. In Taylor, F. J. R. [Ed.] The Biology of Dinoflagellates. Blackwell, Oxford, UK, pp. 92-142. Donner, S. D., Skirving, W. J., Little, C. M., Oppenheimer, M., & Hoegh-Guldberg, O. 2005. Global assessement of coral bleaching and required rates of adaptation under climate change. Global Change Biol. 11:2251-2265. Dunn, S. R., & Weis, V. M. 2009. Apoptosis as a post-phagocytic winnowing mechanism in a coral-dinoflagellate mutualism. Environmental Microbiol. 11:268-275. Fabricius, K. E., Miego, J. C., Colin, P. L., Idip, D., & van Oppen, M. J. H. 2004. Identity and diversity of coral symbionts (zooxanthellae) from three Palauan reefs with contrasting bleaching, temperature and shading histories. Mol. Ecol. 13: 2445-2458. Falkowski, P.G., Dubinsky, Z., Muscatine, L., & Porter, J.W. 1984. Light and the bioenergetics of a symbiotic coral. Bioscience. 34:705-709. Fang, F. C. 2004. Antimicrobial reactive oxygen and nitrogen species: concepts and controversies. Nature Reviews. 2: 820-832. Fautin, D. G. 2002. Reproduction of Cnidaria. Canadian Journal of Zoology. 80:1735-1754. Fitt, W.K. 1984. The role of chemosensory behavior of Symbiodinium microadriaticum, intermediate hosts, and host behavior in the infection of coelenterates. Mar. Biol. 81:9-17. Fitt, W. K. 1985. Chemosensory responses of the symbiotic dinoflagellate Symbiodinium microadriaticum (Dinophyceae). J Phycol. 21:62-67. Fitt, W.K., Brown, B.E., Warner, M.E., and Dunne, R.P. 2001. Coral bleaching: interpretation of thermal tolerance limits and thermal thresholds in tropical corals. Coral Reefs. 20: 51-65. Fitt, W.K., Chang, S.S., & Trench, R.K. 1981. Motility patterns of different strains of the symbiotic dinoflagellate Symbiodinium (=Gymnodinium) microadriaticum (Freudenthal) in culture. Bull. of Mar Sci. 31: 436-443. Fitt, W. K., & Trench, R. K., 1983. Endocytosis of the symbiotic dinoflagellate Symbiodinium microadriaticum Freudenthal by endodermal cells of the scyphistomae of Cassiopeia xamachana and resistance of the algae to host digestion. J Cell Sci. 64:195-212. Franceschi, V. R. & Nakata, P. A. 2005. Calcium oxalate in plants: formation and function. Annu. Rev. Plant Biol. 56:41-71. 110 Freudenthal, H. D. 1962. Symbiodinium gen. nov. and Symbiodinium microadriaticum sp. nov., a zooxanthella: taxonomy, life cycle, and morphology. J Euk. Microbiol. 9:45-52. Furla, P., Allemand, D., Shick, J. M., Ferrier-Pages, C., Richier, S., Plantivaux, A., Merle, P. L., & Tambutte, S. 2005. The symbiotic anthozoan: a physiological chimera between alga and animal. Integrative and Comparative Biology. 45:595-604. Garcia-Ramos, M., & Banaszak, A.T. 2007. The distribution of the dinoflagellate Symbiodinium in the conch Strombus gigas. Gulf and Carib. Fisheries. Inst. 58:403-406. Gast, R. J., & Caron, D. A. 2001. Photosymbiotic associations in planktonic foraminifera and radiolaria. Hydrobiologia. 461:1-7. Gates, R. D., Bagiidasarian, G., & Muscatine, L. 1992. Temperature stress causes host cell detachment in symbiotic cnidarians: implications for coral bleaching. Biol. Bull. 182: 324-332. Glynn, P. W. 1991. Coral reef bleaching in the 1980s and possible connections with global warming. Trends in Ecol. Evol. 6: 175-179. Glynn, P. W., Mate, J. L., Baker, A. C., & Calderon, M. O. 2001. Coral bleaching and mortality in Panama and Ecuador during the 1997-1998 El Nino-Southern Oscillation event: spatial/temporal patterns and comparisons with the 1982-1983 event. Bull. Mar. Sci. 69: 79-109. Goodson, M. S., Whitehead, L. F., & Douglas, A. E. 2001. Symbiotic dinoflagellates in marine Cnidaria: diversity and function. Hydrobiol. 461:79-82. Gordan, B. R., & Leggat, W. 2010. Symbiodinium-Invertebrate symbioses and the role of metabolomics. Mar. Drugs. 8:2546-2568. Goulet, T. L. 2006. Most corals may not change their symbionts. Mar Ecol Prog Ser. 321:1-7. Goulet, T. L. 2007. Most scleractinian corals and octocorals host a single symbiotic zooxanthella clade. Mar Ecol Prog Ser. 335:243-248. Goulet, T. L., & Coffroth, M. A. 1997. A within colony comparison of zooxanthella genotypes in the Caribbean gorgonian Plexaura kuna. Proc. 8th Int. Coral Reef Sym. 2:1331-1334. Goulet, T.L., & Coffroth, M.A. 2003a. Genetic composition of zooxanthellae between and within colonies of the octocoral Plexaura kuna, based on small subunit rDNA and multilocus DNA fingerprinting. Mar. Biol. 142:233-239. Goulet, T.L., & Coffroth, M.A. 2003b. Stability of an octocoral-algal symbiosis over time and space. Mar. Ecol. Prog. Ser. 250:117-124. 111 Goulet, T.L., & Coffroth, M.A. 2004. The genetic identity of dinoflagellate symbionts in Caribbean octocorals. Coral Reefs. 23:117-124. Goulet, T. L., LaJeunesse, T. C., & Fabricius, K. E. 2008. Symbiont specificity and bleaching susceptibility among soft corals in the 1998 Great Barrier Reef mass coral bleaching event. Mar Biol. 154:795-804. Grottoli, A. G., Rodrigues, L. J., & Palardy, J. E. 2006. Heterotrophic plasticity and resilience in bleached corals. Nature. 440:1186-1189. Guillard, R.R.L., & Ryther, J.H. 1962. Studies of marine planktonic diatoms: I. Cyclotella nana Hustedt, and Detonula confervacea (Cleve) Gran. Can. J. Microbiol. 8:229-239. Hallock, P. 2001. Coral reefs, carbonate sediments, nutrients and global change. In Stanley Jr. GD (ed) The History and Sedimentology of Ancient Reef Systems. Springer-Verlag, New York, 387-427. Hallock, P., Lidz, B. H., Cockey-Burkhard, E. M., & Donnelly, K. B. 2003. Foraminifera as bioindicators in coral reef assessment and monitoring: The FORAM Index. Environmental Monitoring and Assessment. 81:221-238. Hanes, S. D. & Kempf, S. C. 2013. Host autophagic degradation and associated symbiont loss in response to heat stress in the symbiotic anemone, Aiptasia pallida. Invertebr. Biol. doi: 10.1111/ivb.12018. Hannack, K., Kestler, P., Sicken, O., & Westheide, W. 1997. The influence of UV radiation on number and ultrastructure of the endosymbiotic dinoflagellates in the sea anemone Cereus pedunculatus (Anthozoa: Actiniaria). Helgol. Mar. Res. 51: 487-502. Hansen, G. & N. Daugbjerg. 2009. Symbiodinium natans sp. nov.: a "free-living" dinoflagellate from Tenerife (Northeast-Atlantic Ocean). J. Phycol. 45:251-263. Harii, S., Kayanne, H., Takigawa, H., Hayashibara, T., & Yamamoto, M. 2002. Larval survivorship, competency periods and settlement of two brooding corals, Heliopora coerulea and Pocillopora damicornis. Mar Biol. 141:39-46. Hill, M. S., 1996. Symbiotic zooxanthellae enhance boring and growth rates of the tropical sponge Anthosigmella varians forma varians. Mar Biol. 125:649-654. Hirose, M., & Hidaka, M. 2006. Early development of zooxanthella-containing eggs of the corals Porites cylindrica and Montipora digitata: The endodermal localization of zooxanthellae. Zooligical Science. 23:873-881. 112 Hirose, M., Kinzie, R. A. III., & Hidaka, M. 2000. Early development of zooxanthella-containing eggs of the corals Pocillopora verrucosa and P. eydouxi with special reference to the distribution of zooxanthellae. Biol Bull. 199: 68-75. Hoegh-Guldberg, O. 1999. Climate change, coral bleaching and the future of the world's coral reefs. Mar. Freshwater. Res. 50: 839-866. Hoegh-Guldberg, O., & Hinde, R. 1986a. Studies on a nudibranch that contains zooanthellae I. Photosynthesis, respiration and the translocation of new fixed carbon by zooxanthellae in Pteraeolidia ianthina. Proceedings of the Royal Society of London Series B. 228:493- 509. Hoegh-Guldberg, O., & Hinde, R. 1986b. Studies on a nudibranch that contains zooxanthellae II. Contribution of zooxanthellae to animal respiration (CZAR) in Pteraeolidia ianthina with high and low densities of zooxanthellae. Proceedings of the Royal Society of London Series B. 228: 511-521. Hoegh-Guldberg, O., Mumby, P. J., Hooten, A. J., Steneck, R. S., Greenfield, P., Gomez, E., Harvell, C. D., Sale, P. F., Edwards, A. J., Caldeira, K., Knowlton, N., Eakin, C. M., Iglesias-Prieto, R., Muthiga, N., Bradbury, R. H., Dubi, A., & Hatziolos, M. E. 2007. Coral reefs under rapid climate change and ocean acidification. Science 318:1737-1742. Hofman, D.K. & Kremer, B.P. 1981. Carbon metabolism and strobilation in Cassiopea andromedea (Cnidaria: Scyphozoa): Significance of endosymbiotic dinoflagellates. Mar Biol. 65: 25-33. Holland, D., & Gabbott, P. 1971. A micro-analytical scheme for determination of protein, carbohydrate, lipid and RNA levels in marine invertebrate larvae. J. Mar. Biol. Ass. U.K. 51: 659-668. Hollingsworth, L. L., Kinzie, R. A. III, Lewis, T. D., Krupp, D. A., & Leong, J-A. C. 2005. Phototaxis of motile zooxanthellae to green light may facilitate symbiont capture by coral larvae. Coral Reefs. 24: 523. Holstein, T., & Tardent, T. 1984. An ultrahigh-speed analysis of exocytosis: nematocyst discharge. Science. 223:830-833. Hoppenrath, M. & Leander, B. S. 2008. Morphology and molecular phylogeny of a new marine sand-dwelling Prorocentrum species, P. Tsawwassenense (Dinophyceae, Prorocentrales), from British Columbia, Canada. J. Phycol. 44:451-466. Huang, H. L., Wang, L. H., Chen, W. N. U., Fang, L. S., & Chen, C. S. 2008. Developmentally regulated localization of endosymbiotic dinoflagellates in different tissue layers of coral larvae. Coral Reefs. 27:365-372. 113 Hughes, T. P., Baird, A. H., Bellwood, D. R., Card, M., Connolly, S. R., Folke, C., Grosberg, R., Hoegh-Guldberg, O., Jackson, J. B. C., Kleypas, J., Lough, J. M., Marshall, P., Nystrom, M., Palumbi, S. R., Pandolfi, J. M., Rosen, B., Roughgarden, J. 2003. Climate change, human impacts, and the resilience of coral reefs. Science. 301:929-933. Hunter, R. L., LaJeunesse, T. C., & Santos, S. R. 2007. Structure and evolution of the rDNA internal transcribed spacer (ITS) region 2 in the symbiotic dinoflagellates (Symbiodinium, Dinophyta). J Phycol. 43: 120-128. Huss, V. A. R., Holweg, C., Seidel, B., Reich, V., Rahat, M., & Kessler, E. 1994. There is an ecological basis for host/symbiont specificity in Chlorella/Hydra symbioses. Endocytobiosis and Cell Research. 10:35-46. Iglesias-Prieto, R., Beltran, V.H., LaJeunesse, T.C., Reyes-Bonilla, H., & Thome, P.E. 2004. Different algal symbionts explain the vertical distribution of dominant reef corals of the eastern Pacific. Proc. R. Soc. Lond. B Biol. Sci. 271:1757-1763. Iglesias-Prieto, R., & Trench, R. K. 1997. Acclimation and adaptation to irradiance in symbiotic dinoflagellates. II. Response of chlorophyll-protein complexes to different photon-flux densities. Mar Biol. 130: 23-33. Jeong, H.J., Lee, C.W., Yih, W.H., & Kim, J.S. 1997. Fradilidium cf. Mexicanum, a thecate mixotrophic dinoflagellate which is prey for and a predator on co-occurring thecate heterotrophic dinoflagellate Protoperidinium cf, divergens. Mar. Ecol. Prog. Ser. 151:299- 305. Jimbo, M., Suda, Y., Koike, K., Nakamura-Tsuruta, S., Kominami, J., Kamei, M., Hirabayashi, J., Sakai, R., & Kamiya, H. 2013. Possible involvement of glycolipids in lectin-mediated cellular transformation of symbiotic microalgae in corals. J. Exp. Mar. Biol. Ecol. 439:129-135. Jimbo, M., Yanohara, T., Koike, K., Koike, K., Sakai, R., Muramoto, K., & Kamiya, H. 2000. The D-galactose-binding lectin of the octocoral Sinularia lochmodes: characterization and possible relationship to the symbiotic dinoflagellates. Comp. Biochem. Physiol. B Comp. Biochem. 125: 227-236. Jokiel, P.L. & Coles, S.L. 1977. Effects of temperature on the mortality and growth of Hawaiian reef corals. Mar. Biol. 43:201-208. Jones, A.M., Berkelmanns, R., van Oppen, M.J.H., Mieog, J.C., &Sinclair, W. 2008. A community change in the algal endosymbionts of a scleactinian coral following a natural bleaching event: field evidence of acclimatization. Proc. R. Soc. Biol. Sci. Ser. B 275:1359- 1365. 114 Kempf, S. C. 1984. Symbiosis between the zooxanthella Symbiodinium (= Gymnodinium) microadriacticum (Freudenthal) and four species of nudibranchs. Biol Bull. 166:110-126. Kempf, S. C. 1991. A ?Primitive? symbiosis between the Aeolid nudibranch Berghia verrucicornis (A. Costa, 1867) and a zooxanthella. J. Mollus. Stud. 57(Supplement 4):75-85. Kevin, M. J., Hall, W. T., McLaughlin, J. J. A., & Zahl, P. A. 1969. Symbiodinium microadriaticum Freudenthal, a revised taxonomic description, ultrastructure. J. Phycol. 28:82-89. Kinzie, R. A. III. 1974. Experimental infection of aposymbiotic gorgonian polyps with zooxanthellae. J Exp Mar Biol Ecol. 15:335-345. Kinzie, R. A. III., Araki, G., & Kin, L. 1977. The specificity of the intracellular symbiosis of zooxanthellae from a range of invertebrate hosts. Abstracts from the Annual Meeting of the American Society of Zoologists and Society of Systematic Zoology, hosted by the Canadian Society of Zoologists. 1977. 53. Kinzie, R. A. III., & Chee, G. S. 1979. The effect of different zooxanthellae on the growth of experimentally reinfected hosts. Biol Bull. 156:315-327. Kinzie, R.A., Jokiel, P.L. & R. York. 1984. Effects of light of altered spectral composition on coral zooxanthellal associations and on zooxanthellae in vitro. Mar. Biol. 78: 239-248. Kinzie, R. A., Takayama, M., Santos, S. R., & Coffroth, M. A. 2001. The adaptive bleaching hypothesis: experimental tests of cricial assumptions. Biol. Bull. 200: 51-58 Koike, K., Jimbo, M., Sakai, R., Kaeriyama, M., Muramoto, K., Ogata, T., Maruyama, T., & Kamiya, H. 2004. Octocoral chemical signaling selects and controls dinoflagellate symbionts. Biol Bull. 207:80-86. Kvennefors, E.C.E., Leggat, W., Hoegh-Guldberg, O., Degnan, B.M., & Barnes, A.C. 2008. An ancient and variable mannose-binding lectin from the coral Acropora millepora binds both pathogens and symbionts. Dev. Comp. Immunol. 32:1582-1592. LaJeunesse, T.C. 2001. Investigating the biodiversity, ecology, and phylogeny of endosymbiotic dinoflagellates in the genus Symbiodinium using the ITS region: in search of a ?species? level marker. J. Phycol. 37:866-880. LaJeunesse, T.C. 2002. Diversity and community structure of symbiotic dionflagellates from Caribbean coral reefs. Mar. Biol. 141:387-400. LaJeunesse, T.C., Loh, W.K.W., van Woesik, R., Hoegh-Guldberg, O., Schmidt, G.W., & Fitt, W.K. 2003. Low symbiont diversity in southern Great Barrier Reef corals, relative to those 115 of the Caribbean. Limnol. Oceanogr. 48:2046-2054. LaJeunesse, T.C., Parkinson, J.E., & J.D. Reimer. 2012. A genetics-based description of Symbiodinium minutum sp. nov. and S. psygmophilum sp. nov. (Dinophyceae), two dinoflagellates symbiotic with cnidaria. J. Phycol. 48: 1380-1391. LaJeunesse, T.C., Smith, R., Walther, M., Pinzon, J., Pettay, D.T., McGinley, M., Aschaffenburg, M., Medina-Rosas, P., Cupul-Magana, A.L., Perez, A.L., Reyes-Bonilla, H., & Warner, M.E. 2010. Host-symbiont recombination versus natural selection in the response of coral- dinoflagellate symbioses to environmental disturbance. Proc. R. Soc. Biol. Sci. Ser. B 277:2925-2934. LaJeunesse, T. C. & Trench, R. K. 2000. Biogeography of two species of Symbiodinium (Freudenthal) inhabiting the intertidal sea anemone Anthopleura elegantissima (Brandt). Biol. Bull. 199:126-134. Lang, J. C., Lasker, H. R., Gladfelter, E. H., Hallock, P., Jaap, W. C., Losada, F. J.,& Muller, R. G. 1992. Spatial and temporal variability during periods of ?recovery? after mass bleaching on western atlantic coral reefs. Society for Integrative and Comparative Biology. 32:696-706. Lesser, M.P. 1989. Photobiology of Natural Populations of Zooxanthellae from the Sea Anemone Aiptasia pallida: assessment of the host?s role in protection against ultraviolet radiation. Cytometry. 10:653-658. Lesser, M. P. 2006. Oxidative stress in marine environments: biochemistry and physiological ecology. Annu. Rev. Physiol. 68:253 -278. Leutenegger, S. 1977. Ultrastructure and motility of dinophyceans symbiotic with larger, imperforated foraminifera. Mar. Biol. 44:157-164. Lewin, R. A. 1982. Symbiosis and parasitism: definitions and evaluations. BioScience. 32:254- 260. Lewis, D.H. & Smith, D.C. 1971. The autotrophic nutrition of symbiotic marine coelenterates with special references to hermatypic corals. I. Movement of photosynthetic products between the symbionts. Proc. R. Soc. Lond. B Biol. Sci. 178:111-129. Lin, K.-L., Wang, J.-T., & Fang, L.-S. 2000. Participation of glycoproteins on zooxanthellal cell walls in the establishment of a symbiotic relationship with the sea anemone, Aiptasia pulchella. Zoological Studies. 39:172-178. Logan, D.D.K., LaFlamme, A.C., Weis, V.M., & Davy, S.K. 2010. Flow-cytometric characterization of the cell-surface glycans of symbiotic dinoflagellates (Symbiodinium spp). J. Phycol. 46:525-533. 116 Loram, J. E., Trapido-Rosenthal, H. G., & Douglas, A. E. 2007. Functional significance of genetically different symbiotic algae Symbiodinium in a coral reef symbiosis. Mol Ecol. 16:4849-4857. Lowry, O. H., Rosebrough, N. J., Farr, A. L., & Randall, R. J. 1951. Protein measurement with the folin phenol reagent. J. Biol. Chem. 193: 265-275. Loya, Y., Sakai, K., Yamazato, K., Nakano, Y., Sambali, H., & van Woesik, R. 2001. Coral bleaching: the winners and losers. Ecology Letters 4: 122-131. Magalon, H., Flot, J. F., & Baudry, E. 2007. Molecular identification of symbiotic dinoflagellates in Pacific corals in the genus Pocillopora. Coral Reefs. 26:551-558. Markell, D. A., & Trench, R. K. 1993. Macromolecules exuded by symbiotic dinoflagellates in culture; amino acid and sugar composition. J Phycol. 29:64-68. Markell, D. A., Trench, R. K., & Igleasias-Prieto, R. 1992. Macromolecules associated with the cell walls of symbiotic dinoflagellates. Symbiosis. 12:19-31. Markell, D.A. & Wood-Charlson, E.M. 2010. Immunocytochemical evidence that symbiotic algae secrete potential recognition signal molecules in hospite. Mar. Biol. 157:1105-1111. Martel, C.M. 2006. Prey location, recognition and ingestion by the phagotrophic marine dinoflagellate Oxyrrhis marina. J. Exp. Mar. Biol. Ecol. 335:210-220. Maynard, J. A., Anthony, K. R. N., Marshall, P. A., & Masiri, I. 2008. Major bleaching events can lead to increased thermal tolerance in corals. Mar. Biol. 155:173-182. McClanahan, T. R. 2004 The relationship between bleaching and mortality of common corals. Mar. Biol. 144: 1239-1245 McNally, K. L., Govind, N. S., Thome, P. E., & Trench, R. K. 1994. Small-subunit ribosomal DNA sequence analyses and a reconstruction of the inferred phylogeny among symbiotic dinoflagellates (Pyrrophyta). J Phycol. 30:316-329. McNeil, P.L. 1981. Mechanisms of nutritive endocytosis I. Phagocytic versatility and cellular recognition in Chlorohydra digestive cells, a scanning electron microscope study. J. Cell Sci. 49:311-339. Media Cybernetics, Inc. 2009. Image Pro Plus 7.0. Rockville, MD. Meints, R.H., & Pardy, R.L. 1980. Quantitative demonstration of cell surface involvement in a plant-animal symbiosis: lectin inhibition of reassociation. J. Cell Sci. 43:239-251. 117 Merle, P.-L., Sabourault, C., Richier, S., Allemand, D. & Furla, P. 2007. Catalase characterization and implication in bleaching of a symbiotic sea anemone. Free Radic. Biol. Med. 42,236 -246. Miller, D. H., Mellman, I. S., Lamport, D. T. A., & Miller, M. 1974. The chemical composition of the cell wall of Chlamydomonas gymnogama and the concept of a plant cell wall protein. J Cell Biol. 63:420-429. Muller-Parker, G., & Davy, S. K. 2001. Temperate and tropical algal-sea anemone symbioses. Invert Biol. 120:104-123. Muller-Parker, G. & D'Elia, C. 1997. Interactions between corals and their symbiotic algae. In Birkeland C. (ed) Life and Death of Coral Reefs. Chapman & Hall, New York, 96-113. Muller-Parker, G., Lee, K. W., & Cook, C. B. 1996. Changes in the ultrastructure of symbiotic zooxanthellae (Symbiodinium sp., Dinophyceae) in fed and starved sea anemones maintained under high and low light. J. Phycol. 32: 987-994. Muscatine, L. 1967. Glycerol excretion by symbiotic algae from corals and Tridacna and its control by the host. Science. 156: 516-518. Muscatine, L. 1990. The role of symbiotic algae in carbon and energy flux in reef corals. In Dubinsky Z. (ed) Coral Reefs Ecosystems of the World. Elsevier, New York, 75-87. Muscatine, L., Falkowski, P.G., Porter, J.W., & Dubinsky, Z. 1984. Fate of photosynthetically fixed carbon in light and shade-adapted colonies of the symbiotic coral, Stylophora pistillata. Proc. R. Soc. Lond. B Biol. Sci. 222:181-202. Muscatine, L., & McCloskey, L. R. 1981 Estimating the daily contribution of carbon from zooxanthellae to coral animal respiration. Limnol Oceanogr. 26:601-611. Muscatine, L., McCloskey, L.R., & Marian, R.E. 1981. Estimating the daily contribution of carbon from zooxanthellae to coral animal respiration. Limnol. Oceanogr. 26: 601-611. Muscatine, L. & Porter, J.W. 1977. Reef corals: mutualistic symbioses adapted to nutrient-poor environments. Bioscience. 27:454-460. Nikon Corporation. 2006. NIS-Elements BR Version 3.2. Nikon Corporation. Tokyo, Japan. Nyholm, S. V., & McFall-Ngai, M. J. 2004. The winnowing establishing the squid-Vibrio symbiosis. Nat. Rev. Microbiol. 2:632-642. Ogunlana, M. V., Hooge, M. D., Tekle, Y. I., Benayahu, Y., Barneah, O., & Tyler, S. 2005. Waminoa brickneri n. sp. (Acoela: Acoelomorpha) associated with corals in the Red Sea. Zootaxa. 1008: 1-11. 118 Oke, V., & Long, S.R. 1999. Bacteroid formation in the Rhizobium-legume symbiosis. Current Opinion in Microbiol. 2:641-646. Oliver, T. A. & Palumbi S. R. 2011. Many corals host thermally resistant symbionts in high- temperature habitat. Coral Reefs. 30: 241-250. Pacher, P., Beckman, J. S. & Liaudet, L. 2007. Nitric oxide and peroxynitrite in health and disease. Physiol. Rev. 87,315 -424. Pasternak, Z., Bachar, A., Abelson, A., & Achituv, Y. 2004. Initiation of symbiosis between the soft coral Heteroxenia fuscescens and its zooxanthellae. Marine Ecology-Progress Series. 279:113-116. Pasternak, Z., Blasius, B., Abelson, A., & Achituv, Y. 2006. Host-finding behavior and navigation capabilities of symbiotic zooxanthellae. Coral Reefs. 25:201-207. Pawlowski, J., Holzmann, M., Fahrni, J. F., Pochon, X., & Lee, J. J. 2001. Molecular identification of algal endosymbionts in large miliolid foraminifera: 2. dinoflagellates. J Euk Microbiol. 48:368-373. Pearse, V.B. 1970. Incorporation of metabolic CO2 into coral skeleton. Nature. 228:383. Pearse, V.B. & Muscatine, L. 1971. Role of symbiotic algae (zooxanthellae) in coral calcification. Biol. Bull. 141:955-958. Perez, S. F., Cook, C. B., & Brooks, W. R. 2001. The role of symbiotic dinoflagellates in the temperature-induced bleaching response of the subtropical sea anemone Aiptasia pallida. J. Exp. Mar. Biol. Ecol. 256: 1-14. Perez, S. & Weis, V. 2006. Nitric oxide and cnidarian bleaching: an eviction notice mediates breakdown of a symbiosis. J. Exp. Biol. 209: 2804-2810. Pochon, X. & Gates, R. D. 2010. A new Symbiodinium clade (Dinophyceae) from soritid foraminifera in Hawai'i. Mol. Phylogenet. Evol. 56:492-497. Pochon, X., LaJeunesse, T.C., & Pawlowski, J. 2004. Biogeographic partitioning and host specialization among foraminiferan dinoflagellate symbionts (Symbiodinium, Dinophyta). Mar. Biol. 146:17-27. Pochon, X., Montoya-Burgos, J.I., Stadelmann, B., & J. Pawlowski. 2006. Molecular phylogeny, evolutionary rates, and divergence timing of the symbiotic dinoflagellate genus Symbiodinium. Mol. Phylog. and Evol. 38: 20-30. 119 Pochon, X., Pawlowski, J., Zaninetti, L., Rowan, R. 2001. High genetic diversity and relative specificity among Symbiodinium-like endosymbiotic dinoflagellates in soritid foraminiferans. Mar. Biol. 139: 1069-1078. Porter, J. W., Fitt, W. K., Spero, H. J., Rogers, C. S., & White, M. W. 1989. Bleaching in reef corals: Physiological and stable isotopic responses. Proc. Natl. Acad. Sci. USA 86: 9342- 9346. Reisser, W., Radunz, A., & Wiessner, W. 1982. Participation of algal surface structures in the cell recognition process during infection of aposymbiotic Paramecium bursaria with symbiotic chlorellae. Cytobios 33:39-50. Richier S, Furla P, Plantivaux A, Merle P-L, & Allemand D 2005. Symbiosis-induced adaptation to oxidative stress. J Exp.Biol. 208:277?285. Richmond, R.H., & Wolanski, E. 2011. Coral Research: Past Efforts and Future Horizons. In ed. Dubinsky, Z. and Stambler, N. 2011. Coral Reefs: An ecosystem in transition. Springer Press. 3-10. Riggs, L. L. 1988. Feeding behavior in Aiptasia tagetes (Duchassaing and Michelotti) planulae: a plausible mechanism for zooxanthellae infection of aposymbiotic planktotrophic planulae. Caribbean Journal of Science. 24:201-206. Robison, J.D. & Warner, M.E. 2006. Differential impacts of photoacclimation and thermal stress on the photobiology of four different phylotypes of Symbiodinium (Pyrrhophyta). J. Phycol. 42:568-579. Rodriques, L. J., & Grottoli, A. G. 2007. Energy reserves and metabolism as indicators of coral recovery from bleaching. Limnol Oceanogr. 52:1874:1882. Rodriguez-Lanetty, M., Krupp, D.A., & Weis, V.M. 2004. Distinct ITS types of Symbiodinium in Clade C correlate with cnidarian/dinoflagellate specificity during onset of symbiosis. Mar. Ecol. Prog. Ser. 275:97-102. Rodriguez-Lanetty, M., Loh, W., Carter, D., & Hoegh-Guldberg, O. 2001. Latitudinal variability in symbiont specificity within the widespread scleractinian coral Plesiatrea versipora. Mar. Biol. 138:1176-1181. Rodriguez-Lanetty, M., Wood-Charlson, E.M., Hollingsworth, L.L., Krupp, D.A., & Weis, V.M. 2006. Temporal and spatial infection dynamics indicate recognition events in the early hours of a dinoflagellate/coral symbiosis. Mar. Biol. 149:713-719. Roth, L. 1989. Bacterium release into host-cells of nitrogen-fixing soybean nodules- the symbiosome membrane comes from 3 sources. Eur. J. Cell Biol. 49:13-23. 120 Roth, L. E. & Stacey, G. 1989. Bacterium release into host cells of nitrogen fixing soybean nodules: The symbiosome membrane comes from three sources. Eur. J. Cell Biol. 49:13- 23. Rowan, R. 1998. Diversity and ecology of zooxanthellae on coral reefs. J. Phycol. 34:407-417. Rowan, R. 2004. Thermal adaptations in reef coral symbionts. Nature. 430:742. Rowan, R., & Knowlton, N. 1995. Intraspecific diversity and ecological zonation in coral algal symbiosis. Proc. Natl. Acad. Sci. U.S.A. 92:2859-2853. Rowan, R., Knowlton, N., Baker, A., & Jara, J. 1997. Landscape ecology of algal symbionts creates variation in episodes of coral bleaching. Nature 388:265-269. Rowan, R. & Powers, D.A. 1991a. A molecular genetic identification of zooxanthellae and the evolution of animal ? algal symbioses. Science 251:1348 ? 1351. Rowan, R., & Powers, D.A. 1991b. Molecular genetic identification of symbiotic dinoflagellates (zooxanthellae). Mar. Ecol. Prog. Ser. 71:65-73. Rowan, R. & Powers, D.A. 1992. Ribosomal-RNA sequences and the diversity of symbiotic dinoflagellates (zooxanthellae). Proc. Natl. Acad. Sci. U.S.A. 89:3639-3643. Rudman, W. B. 1991. Further studies on the taxonomy and biology of the octocoral-feeding genus Phyllodesmium Ehrenberg, 1831 (Nudibranchia: Aeolidoidea). Journal of Molluscan Studies. 57:167-203. Rudman, W. B., Bergquist, P. R. 2007. A review of feeding specificity in the sponge-feeding Chromorididae (Nudibranchia: Mollusca). Molluscan Research: Malacological Society of Australia. 27:60-88. Ruppert, E. E., Fox, R. S., Barnes, R. D. 2004. Invertebrate Zoology. 7th Edition. Brooks/Cole. Sachs, J. L., & Wilcox, T. P. 2006. A shift to parasitism in the jellyfish symbiont Symbiodinium microadriaticum. Proceedings of the Royal Society of London Series B. 273:425-429. Saldarriaga, J.F., Taylor, F.J.R., Keeling, P.J. & T. Cavalier-Smith. 2001. Dinoflagellate nuclear SSU rRNA phylogeny suggests multiple plastid losses and replacements. J. Mol. Evol. 53: 204-213. Santos, S.R. 2012. Symbiodinium culture list. Available at: http://www.auburn.edu/~santosr/phplabware.htm (last accessed 16 August 2012). Santos, S. R., & Coffroth, M. A. 2003. Molecular Genetic Evidence that dinoflagellates belonging to the genus Symbiodinium Freudenthal are haploid. Biol Bull. 204: 10-20. 121 Santos, S.R., Gutierrez, R.C., & Coffroth, M.A. 2003. Phylogenetic identification of symbiotic dinoflagellates via length heteroplasmy in domain V of chloroplast large subunit (cp23S)- ribosomal DNA sequences. Mar. Biotechnol. 5:130-140. Santos, S.R., Taylor, D.J., Coffroth, M.A. 2001. Genetic comparisons of freshly isolated versus cultured symbiotic dinoflagellates: implications for extrapolating to the intact symbiosis. J. Phycol. 37: 900-912. Santos, S.R., Taylor, D.J., Kinzie, R.A. III, Hidaka, M., Sakai, K., & Coffroth, M.A. 2002. ?Molecular phylogeny of symbiotic dinoflagellates inferred from partial chloroplast large subunit (23S)-rDNA sequences?. Mol. Phylogenet. Evol. 23:97-111. SAS Institute Inc. 2009. SAS? 9.2 Macro Language: Reference. Cary, NC: SAS Institute Inc. Schmid, V., Ono, S. I., & Reber-Muller, S. 1999. Cell substrate interactions in Cnidaria. Microscopy Research and Technique. 44:254-268. Schoenberg, D.A. & Trench R.K. 1980a. Genetic variation in Symbiodinium (=Gymnodinium) microadriaticum Freudenthal, and specificity in its symbiosis with marine invertebrates. I. Isoenzyme and soluble-protein patterns of axenic cultures of Symbiodinium microadriaticum. Proc. R. Soc. Lond. B Biol. Sci. 207:405-427. Schoenberg, D.A. & Trench R.K. 1980b. Genetic variation in Symbiodinium (=Gymnodinium) microadriaticum Freudenthal, and specificity in its symbiosis with marine invertebrates. II. Morphological variation in Symbiodinium microadriaticum. Proc. R. Soc. Lond. B Biol. Sci. 207:429-444. Schoenberg, D.A. & Trench R.K. 1980c. Genetic variation in Symbiodinium (=Gymnodinium) microadriaticum Freudenthal, and specificity in its symbiosis with marine invertebrates. III. Specificity and infectivity of Symbiodinium microadriaticum. Proc. R. Soc. Lond. B Biol. Sci. 207:445-460. Schwarz, J.A., Krupp, D.A., & Weis, V.M. 1999. Late larval development and onset of symbiosis in scleractinian coral Fungia scutaria. Biol. Bull. 196:70-79. Schwarz, J.A., Weis, V.M., & Potts, D.C. 2002. Feeding behavior and acquisition of zooxanthellae by the planulae larvae of the sea anemone Anthopleura elegantissima. Mar. Biol. 140:417-478. Secord, D.L. 1995. Host specificity and symbiotic interactions in sea anemones (Anthopleura elegantissima, Symbiodinium, Chorella). PhD Dissertation, University of Washington. 88pp. 122 Stambler, N., & Dubinsky, Z. 1987. Energy relationships between Anemonia sulcata and its endosymbiotic zooxanthellae. Symbiosis. 3:233-248. Stat, M., Carter, D., & Hoegh-Guldberg, O. 2006. The evolutionary history of Symbiondinium and scleractinian hosts?symbiosis, diversity, and the effect of climate change. Perspectives in Plant Ecology, Evolution and Systematics. 8:23-43. Stat, M., & Gates, R.D. 2011. Clade D Symbiodinium in scleractinian corals: A ?nugget? of hope, a selfish opportunist, an ominous sign, or all of the above? J. Mar. Biol. 2011. 1-9. Steele, R. D. 1977. The significance of zooxanthella-containing pellets extruded by sea anemones. Bull. Mar. Sci. 27:591-594. Steinke, M., Brading, P., Kerrison, P., Warner, M.E., & Suggett, D.J. 2011. Concentrations of dimethylsulfoniopropionate and dimethylsulfide are strain-specific in symbiotic dinoflagellates (Symbiodinium sp., Dinophyceae). J. Phycol. 47:775-783. Sturz, A. V., Christie, B. R., Matheson, B. G., & Nowak, J. 1997. Biodiversity of endophytic bacteria which colonize red clover nodules, roots, stems and foliage and their influence on host growth. Biol Fertil Soils 25:13-19. Takabayashi, M., Santos, S. R., & Cook, C.B. 2004. Mitochondrial DNA phylogeny of the symbiotic dinoflagellates (Symbiodinium, Dinophyta). J Phycol. 40: 160-164. Takeda, K., Kaisho, T., & Akira, S. 2003. Toll-like receptors. Annu. Rev. Immunol. 21:335-376. Tanaka, Y., Miyajima, T., Koike, I., Hayashibara, T., & Ogawa, H. 2006. Translocation and conservation of organic nitrogen within the coral-zooxanthella symbiotic system of Acropora pulchra as demonstrated by dual isotope-labeling techniques. J Exp Mar Biol Ecol. 336:110-119. Taylor, D. L. 1968. In situ studies on the cytochemistry and ultrastructure of a symbiotic marine dinoflagellate. J. Mar. Biol. Ass. U.K. 48:349-366. Taylor, D. L. 1969. Identity of zooxanthellae isolated from some Pacific Tridacnidae. J. Phycol. 5:336-340. Taylor, D. L. 1974. Symbiotic marine algae: taxonomy and biological fitness. In Vernberg W.B. (ed) Symbiosis in the Sea. University of South Carolina Press, South Carolina, 245-291. Taylor, M. W., Radax, R., Steger, D., & Wagner, M. 2007. Sponge-associated microorganisms: evolution, ecology, and biotechnological potential. Microbiol Mol Biol Rev. 71:295-347. 123 Tchernov, D., Gorbunov, M. Y., de Vargas, C., Yadav, S. N., Milligan, A. J., Haggblom, M., & Falkowski, P. G. 2004. Membrane lipids of symbiotic algae are diagnostic of sensitivity to thermal bleaching in corals. PNAS. 101(37): 13531-13535. Thornhill, D. J., Fitt, W. K., & Schmidt, G. W. 2006. Highly stable symbioses among western atlantic brooding corals. Coral Reefs. 25:515-519. Thornhill, D.J., LaJeunesse, T.C., Kemp, D.W., Fitt, W.K., & Schmidt, G.W. 2006. Multi-year, seasonal genotypic surveys of coral-algal symbioses reveal prevalent stability or post- bleaching reversion. Mar. Biol. 148:711-722. Thornhill, D. J., LaJeunesse, T. C., & Santos, S. R. 2007. Measuring rDNA diversity in eukaryotic microbial systems: how intragenic variation, pseudogenes, and PCR artifacts confound biodiversity estimates. Mol Ecol. 16:5326-5340. Tibayrenc, M., Kjellberg, F., Arnaud, J., Oury, B., Breniere, S. F., Darde, M. & Ayala, F. J. 1991. Are eukaryotic microorganisms clonal or sexual? A population genetics vantage. Proc. Natl. Acad. Sci. USA 88:5129-5133. Trapido-Rosenthal, H.G., Sharp, K.H., Galloway, T.S., & Morrall, C.E. 2001. Nitric oxide and cnidarian-dinoflagellate symbioses: pieces of a puzzle. Am. Zool. 41:247-257. Trench, R. K. 1971a. The physiology and biochemistry of zooxanthellae symbiotic with marine coelenterates. I. The assimilation of photosynthetic products of zooxanthellae by two marine coelenterates. Proc. Roy. Soc. Lond. B. 177:225-235. Trench, R. K. 1971b. The physiology and biochemistry of zooxanthellae symbiotic with marine coelenterates. II. Liberation of fixed 14C by zooxanthellae in vitro. Proc. Roy. Soc. Lond. B. 177:237-250. Trench, R. K. 1971c. The physiology and biochemistry of zooxanthellae symbiotic with marine coelenterates. III. The effect of homogenates of host tissues on the excretion of photosynthetic products in vitro by zooxanthellae from two marine coelenterates. Proc. Roy. Soc. Lond. B. 177:251-264. Trench, R. K. 1979. The cell biology of plant-animal symbiosis. Annual Review of Plant Physiology. 30: 485-531. Trench, R. K. 1993. Microalgal-invertebrate symbioses: a review. Endocytobiosis & Cell Res. 9:135-175. Trench, R. K., & Blank, R. J. 1987. Symbiodinium microadriaticum Freudenthal, S. goreauii sp. nov., S. kawagutii sp. nov., and S. pilosum sp. nov.: Gymnodinoid dinoflagellate symbionts of marine invertebrates. J Phycol. 23:469-481. 124 Trench, R.K., Colley, N.J., & Fitt, W.K. 1981. Recognition phenomena in symbioses between marine invertebrates and ?zooxanthellae?; uptake, sequestration and persistence. J. Plant Biol. 94:529-545. Trench, R. K., & Thinh, L. V. 1995. Gymnodinium linucheae sp. nov.: the dinoflagellate symbiont of the jellyfish Linuche unguiculata. Eur J Phycol. 30: 149-154. Trench, R. K., Wethey, D. S., & Porter, J. W. 1981. Observations on the symbiosis with zooxanthellae among the tridacnidae (Mollusca, Bivalvia). Biol Bull. 161:180-198. Ulstrup, K.E., & van Oppen, M.J.H. 2003. Geographic and habitat partitioning of genetically distinct zooxanthellae (Symbiodinium) in Acropora corals on the Great Barrier Reef. Mar. Ecol. 12: 3477-3484. Van Oppen, M. J. H. 2004. Mode of zooxanthellae transmission does not affect zooxanthella diversity in acroporid corals. Mar Biol. 144:1-7. van Oppen, M.J.H., Palstra, F.P., Piquet, A.M.T., & Miller, D.J. 2001. Patterns of coral- dinoflagellate associations in Acropora: significance of local availability and physiology of Symbiodinium strains and host-symbiont selectivity. Proc. R. Soc. Lond. B Biol. Sci. 268:1759-1767. Van Thinh, L., Griffiths, D. J. & Winsor, H. 1986. Ultrastructure of Symbiodinium microadriaticum (Dinophyceae) symbiotic with Zoanthus sp. (Zoanthidea). Phycologia. 25:178-184. Verde, E.A. & McCloskey, L.R. 1998. Production, respiration, and photophysiology of the mangrove jellyfish Cassiopea xamachana symbiotic with zooxanthellae: effect of jellyfish size and season. Mar Ecol Prog Ser. 168: 147-162. Wakefield, T. S., Farmer, M. A. & Kempf, S. C. 2000. Revised description of the fine structure of in situ ?Zooxanthellae? Genus Symbiodinium. Biol. Bull. 199:76-84. Wakefield, T. S. & Kempf, S. C. 2001. Development of host- and symbiont-specific monoclonal antibodies and confirmation of the origin of the symbiosome membrane in a cnidarian- dinoflagellate symbiosis. Biol. Bull. 200: 127-143. Warner, M. E., Fitt, W. K., & Schmidt, G. W. 1996. The effects of elevated temperature on the photosynthetic efficiency of zooxanthellae in hospite from four different species of reef coral: a novel approach. Plant Cell Environ. 19: 291-299. Watson, G. M., & Hessinger, D. A. 1989. Cnidocyte mechanoreceptors are tuned to the movements of swimming prey by chemoreceptors. Science. 243:1589-1591. 125 Weis, V. 2008. Cellular mechanisms of cnidarian bleaching: stress causes the collapse of symbiosis. J. Exp. Biol. 211: 3059-3066 Weis, V. M. 2010. The susceptibility and resilience of corals to thermal stress: adaptation, acclimatization or both? Mol. Ecol. 19: 1515-1517. Weis, W.I., & Drickamer, K. 1996. Structural basis of lectin-carbohydrate recognition. Annu. Rev. Biochem. 65:441-473. Weis, V. M., Reynolds, W. S., deBoer, M. D. & Krupp, D. A. 2001. Host-symbiont specificity during onset of symbiosis between the dinoflagellates Symbiodinium spp. and planula larvae of the scleractinian coral Fungia scutaria. Coral Reefs. 20:301-308. Whitehead, L. F., & Douglas, A. E. 2003. Metabolite comparisons and the identity of nutrients translocated from symbiotic algae to an animal host. J Exp Biol. 206:3149-3157. Wilcox, T.P. 1998. Large-subunit ribosomal RNA systematics of symbiotic dinoflagellates: morphology does not recapitulate phylogeny. Mol. Phylog. and Evol. 10: 436-448. Wood-Charlson, E.M., Hollingsworth, L.L., Krupp, D.A., & Weis, V.M. 2006. Lectin/glycan interactions play a role in recognition in a coral/dinoflagellate symbiosis. Cell. Microbiol. 8:1985-1993. Yacobovitch, T., Benayahu, Y., & Weis, V. M. 2004. Motility of zooxanthellae isolated from the Red Sea soft coral Heteroxenia fuscescens (Cnidaria). J. Exp. Mar. Biol. And Ecol. 298:35-48. Yellowlees, D., Rees, T. A. V., & Leggat, W. 2008. Metabolic interactions between algal symbionts and invertebrate hosts. Plant, Cell and Environ. 31:679-694. Young, J. P. W., & Johnston, A. W. B. 1989. The evolution of specificity in the legume- Rhizobium symbiosis. Trends in Ecol and Evol. 4: 341-349. Zerial, M., & McBride, H. 2001. Rab proteins as membrane organizers. Nature Reviews: Molecular Cell Biology. 2:107-118. Zhang, Z., Green, B.R., & Cavalier-Smith, T., 2000. Phylogeny of ultra- rapidly evolving dinoflagellate chloroplast genes: A possible common origin for sporozoan and dinoflagellate plastids. J. Mol. Evol. 51:26?41.