Biol. Bull. Sign up for etocs!
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (19)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Santos, S. R.
Right arrow Articles by Coffroth, M. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Santos, S. R.
Right arrow Articles by Coffroth, M. A.
Related Collections
Right arrow Algae
Right arrow Cell Biology
Right arrow Cnidarians
Right arrow Evolution
Right arrow Population Biology
Right arrow Symbiosis
Right arrow Phylogeny
Biol. Bull. 204: 10-20. (February 2003)
© 2003 Marine Biological Laboratory

Molecular Genetic Evidence that Dinoflagellates Belonging to the Genus Symbiodinium Freudenthal Are Haploid

Scott R. Santos* and Mary Alice Coffroth{ddagger}

Department of Biological Science, State University of New York at Buffalo, Buffalo, New York 14260-1300

{ddagger} To whom correspondence should be addressed. E-mail: coffroth{at}buffalo.edu


   Abstract
TOP
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Literature Cited
 
Microscopic and cytological evidence suggest that many dinoflagellates possess a haploid nuclear phase. However, the ploidy of a number of dinoflagellates remains unknown, and molecular genetic support for haploidy in this group has been lacking. To elucidate the ploidy of symbiotic dinoflagellates belonging to the genus Symbiodinium, we used five polymorphic microsatellites to examine populations harbored by the Caribbean gorgonians Plexaura kuna and Pseudopterogorgia elisabethae; we also studied a series of Symbiodinium cultures. In 690 out of 728 Symbiodinium samples in hospite (95% of the cases) and in all 45 Symbiodinium cultures, only a single allele was recovered per locus. Statistical testing of the Symbiodinium populations harbored by P. elisabethae revealed that the observed genotype frequencies deviate significantly from those expected under Hardy-Weinberg equilibrium. Taken together, our results confirm that, in the vegetative life stage, members of Symbiodinium, both cultured and in hospite, are haploid. Furthermore, based on the phylogenetics of the dinoflagellates, haploidy in vegetative cells appears to be an ancestral trait that extends to all 2000 extant species of these important unicellular protists.


   Introduction
TOP
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Literature Cited
 
The ploidy of an organism can significantly affect genome evolution. For example, diploids carry twice as much DNA as haploids and may be expected to accumulate new beneficial mutations at a higher rate (Paquin and Adams, 1983). The genome of diploids may also evolve rapidly because they carry more than a single copy of an allele. These "extra" alleles, over time, may evolve new functions while "old" alleles continue to perform their original functions (Lewis and Wolpert, 1979). Haploids, on the other hand, tend to have deleterious mutations purged more rapidly from the population since they are not masked (Hughes and Otto, 1999). Furthermore, knowledge of ploidy is essential to the interpretation and understanding of population genetic data. Given the importance of ploidy to genome evolution and population genetics, it is surprising that questions pertaining to it remain for a number of organisms.

Dinoflagellates are a diverse and ecologically important group of unicellular protists. For example, some species are major photosynthetic or heterotrophic components of the plankton, and others are considered to be the causative agents of fish kills. Microscopic and cytological evidence from the species examined to date suggests that dinoflagellates (with the exception of Noctiluca spp.) possess a vegetative haploid nuclear phase (Pfiester and Anderson, 1987; Coats, 2002). However, ploidy has not been explicitly determined for a number of dinoflagellates, including the important genus Symbiodinium Freudenthal (Taylor, 1974). Members of Symbiodinium, commonly referred to as zooxanthellae, are intra- or intercellular symbionts of marine invertebrates, including foraminiferans, sponges, cnidarians, and molluscs (Glynn, 1996). Blank (1987) reconstructed the nucleus of Symbiodinium kawagutii and found that the chromosomes of this dinoflagellate could not be paired either by size, appearance, or distribution. This cytological result led to the speculation that the vegetative (coccoid) cells of Symbiodinium may be haploid (Blank, 1987). To date, the molecular genetic data necessary to establish the ploidy of these symbiotic dinoflagellates, or any dinoflagellate, are lacking. We resolve this lingering question by using microsatellites to elucidate the ploidy of Symbiodinium.

Microsatellites are simple, tandemly repeated DNA sequences (reviewed in Chambers and MacAvoy, 2000) that are distributed abundantly in the genomes of virtually all organisms (Bennett, 2000). These single-locus, multiallelic, codominant segments of DNA are highly versatile and accessible markers for population genetic studies. However, microsatellites have also been successfully applied as tools in determining an organism’s ploidy. For example, recovery of two alleles from a single locus in a single individual or isolate demonstrates a diploid nuclear phase. On the other hand, if a single allele is recovered from each of multiple loci, and the result is replicated over multiple individuals or isolates, the data suggest a haploid nuclear phase. This rationale has been applied to numerous organisms, including a parasitic protozoan, Trypanosoma cruzi (Oliveira et al., 1998); two parasitic fungi, Magnaporthe grisea (Brondani et al., 2000) and Aspergillus fumigatus (Bart-Delabesse et al., 1998); a diatom, Ditylum brightwellii (Rynearson and Armbrust, 2000); a bryophyte, Polytrichum formosum (van der Velde et al., 2001); and a false spider mite, Brevipalpus phoenicis (Weeks et al., 2001). In this investigation, we apply the same strategy to members of the genus Symbiodinium. Populations of Symbiodinium harbored by the gorgonians Plexaura kuna and Pseudopterogorgia elisabethae, as well as a series of Symbiodinium cultures, were screened with five polymorphic microsatellites. The results confirm that members of Symbiodinium are haploid in the vegetative life stage.


   Materials and Methods
TOP
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Literature Cited
 
Biological materials and nucleic acid isolation
Samples of Plexaura kuna were collected from colonies at depths of 1–7 m at 10 sites in the San Blas Islands, Republic of Panama (n = 142); two sites in the Florida Keys (n = 12); and three sites in the Bahamas (n = 6). For Pseudopterogorgia elisabethae, samples were collected from 575 colonies at depths of 10–27 m at 12 sites in the Bahamas (n = 43–50 per site). The 12 P. elisabethae sites were Sweetings Cay, Gorda Rock, Little San Salvador, Cat Island, Rum Cay, Hog Cay, and two sites each at Abacos, Eleuthera, and San Salvador Islands. Immediately after collection, samples were preserved in either 95% ethanol or salt-saturated DMSO (Seutin et al., 1991) or were frozen in a liquid nitrogen vapor shipper. Total nucleic acids were extracted and quantified from branch pieces (about 2 cm) of P. kuna according to the methods of Coffroth et al. (1992). Nucleic acids were extracted from branch pieces (about 3 cm) of P. elisabethae by first grinding the tissue in STE buffer (0.05 M Tris-HCl (pH = 8.0), 0.1 M EDTA, 0.1 M NaCl, 0.2% SDS) followed by a modified extraction protocol using the Prep-A-Gene DNA Extraction Kit (Bio-Rad Laboratories, Hercules, CA) (Shearer and Coffroth, unpubl.). These extraction methods extracted nucleic acids from both the gorgonian hosts and from their Symbiodinium populations in hospite.

Symbiodinium cultures, isolated from a range of invertebrate hosts and geographic locations (Table 1), were maintained as described in Santos et al. (2001). Total nucleic acids were extracted and quantified from fresh algal cultures (approximately 5 x 103 cells) as described above for the P. kuna samples. After extraction, all nucleic acid samples were stored at -20 °C.


View this table:
[in this window]
[in a new window]
  		Table 1 Information about the Symbiodinium cultures that amplified with at least one of the two microsatellites primer sets designed for the Symbiodinium populations of Pseudopterogorgia elisabethae

 
Phylogenetic identification of Symbiodinium populations and cultures
The phylogenetic identity of the Symbiodinium populations harbored by P. kuna and P. elisabethae, as well as the Symbiodinium cultures, was determined by length heteroplasmy in domain V of chloroplast large subunit (cp23S) rDNA as described in Santos et al. (2003). This technique rapidly genotypes Symbiodinium isolates and places them into a phylogenetic framework based on cp23S-rDNA (Santos et al., 2003).

Construction of Symbiodinium microsatellite libraries
Nucleic acids used in the construction of the microsatellite libraries for P. kuna and P. elisabethae Symbiodinium populations were obtained by different procedures. Symbiodinium was isolated from samples of P. kuna collected from around the Caribbean and purified of host tissue (described in Santos et al., 2001) before the nucleic acids were extracted and quantified as described above for P. kuna samples. On the other hand, nucleic acids for the Symbiodinium microsatellite library from P. elisabethae were extracted from colonies collected at Sweetings Cay (n = 2) and San Salvador (n = 4) as described above. These preparations contained nucleic acids from both P. elisabethae and their Symbiodinium populations in hospite, with no effort being made to enrich for Symbiodinium nucleic acids. After extraction, nucleic acids were pooled according to host source and used to construct the microsatellite libraries. The libraries were constructed and screened for microsatellites with dinucleotide repeats as described in Ciofi and Bruford (1998).

Screening of microsatellite loci polymerase chain reaction primers
To ensure that the microsatellite loci were not part of the host gorgonian genome, PCR amplifications were carried out on nucleic acids extracted from planulae of P. kuna and P. elisabethae, which are asymbiotic upon release from the maternal colony (Kinzie, 1974; Coffroth et al., 2001; see below). Nucleic acids were extracted from the planulae as described above for samples of P. kuna. To test for the presence of template nucleic acids (i.e., cnidarian host DNA), PCR amplifications were conducted using the universal n18S-rDNA primer set ss5 and ss3 (Rowan and Powers, 1991). The asymbiotic nature of the planulae was then confirmed by the absence of a PCR product when the zooxanthella-biased n18S-rDNA primers ss5 and ss3z were used (Rowan and Powers, 1991). Both sets of reactions were carried out under the conditions described by Rowan and Powers (1991). Details of the five Symbiodinium microsatellite loci, including the sequences of the PCR primers and GenBank accession numbers, are given in Table 2.


View this table:
[in this window]
[in a new window]
  		Table 2 Characterization and PCR amplification conditions of the five Symbiodinium clade B microsatellite loci; size refers to the predicted PCR product size with original sequenced clone as template

 
Microsatellite amplifications and allele detection for Symbiodinium populations
PCR reactions for each Symbiodinium microsatellite locus were performed in 10-µl volumes containing 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 0.001% gelatin, 200 µM dNTPs, 1 U Taq polymerase, approximately 10 ng of template DNA, and MgCl2 and primers at concentrations detailed in Table 2. Thermocycling conditions were as follows: initial denaturing step of 94°C for 2–3 min, 35–39 cycles of 94°C for 30–45 s, annealing temperature of X° (see Table 2) for 30–45 s, 72°C for 30 s, and a final extension of 72°C for 3–5 min.

Most Symbiodinium microsatellite alleles from P. kuna were separated in 2% 0.5X Tris-borate (TBE) agarose gels and visualized by ethidium bromide staining. In addition, some Symbiodinium microsatellite alleles from P. kuna were amplified in the presence of 33P-ATP (Sambrook et al., 1989), separated in 6% polyacrylamide / 0.5X TBE gels under denaturing (7 M urea) conditions, and visualized by exposure to X-ray film. The sizes of Symbiodinium microsatellite alleles from P. kuna were determined either with a 100-bp DNA ladder (MBI Fermentas, Hanover, MD) for 2% agarose gels, or for polyacrylamide gels, with a 33P-ATP puC19 plasmid sequencing reaction serving as a nucleotide size-ladder.

For P. elisabethae, Symbiodinium microsatellite alleles were labeled by incorporation of a 5'-IRD800 M13 Forward primer (see Table 2), and were separated in 25-cm-long, 0.25-mm-thick, 6.5% Long Ranger (FMC Bioproducts, Rockland, ME)/0.5X TBE gels under denaturing (7 M urea) conditions. Gel electrophoresis was performed at 1500 V, 40 W, 40 mA, 50 °C, and the default scan speed, with LI-COR’s NEN Global IR2 DNA sequencer system (LI-COR Biotechnology Division, Lincoln, NE). The sizes of Symbiodinium microsatellite alleles from P. elisabethae were determined with the fragment analysis program Gene ImagIR v3.55 (Scanalytics Inc, Fairfax, VA) using DNA ladders (ladder 97–147 bp or 172–272 bp, rungs at 25-bp increments, ladders loaded every fifth lane) as size references.

In addition, Symbiodinium microsatellite-loci primer-sets for each host species were tested on the Symbiodinium populations of the other host species. For the host species comparisons, PCR reaction conditions and detection of Symbiodinium microsatellite alleles were as previously described.

A total of 45 Symbiodinium cultures were also screened with the five Symbiodinium microsatellite loci primer sets using the methods described above.

Statistical testing of microsatellite data
Exact probabilities (analogous to Fisher’s exact test for 2 x 2 contingency tables) for the observed genotype frequencies and for those expected under Hardy-Weinberg equilibrium were used to determine whether populations of Symbiodinium harbored by P. elisabethae in the Bahamas deviate from Hardy-Weinberg equilibrium. Significance values were calculated with the computer program BIOSYS-2 (Swofford and Selander, 1981; modified by William C. Black, Department of Microbiology, Colorado State University; available at: ftp://lamar.colostate.edu/pub/wcb4/).


   Results
TOP
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Literature Cited
 
Phylogenetic identification of Symbiodinium populations and cultures
The numerically dominant Symbiodinium in association with Plexaura kuna and Pseudopterogorgia elisabethae belong to Symbiodinium clade B (sensu Rowan and Powers, 1991), and more specifically, to Symbiodinium B184 (sensu Santos et al., 2003). The Symbiodinium cultures also belong to clade B, with representatives from Symbiodinium B184, B211, and B223 (sensu Santos et al., 2003; Table 3).


View this table:
[in this window]
[in a new window]
  		Table 3 Microsatellite and chloroplast genotypes for the Symbiodinium cultures; a microsatellite genotype is defined as a unique combination of microsatellite allele sizes at loci CA4.86 and CA6.38

 
Genomic location of the microsatellite loci
Nucleic acids extracted from planulae of P. kuna and P. elisabethae failed to produce an amplicon with the zooxanthella-biased n18S-rDNA PCR primers ss5 and ss3z. On the other hand, an amplicon of approximately 1800 bp was generated from the same templates using the universal n18S-rDNA PCR primers ss5 and ss3 (data not shown). These results confirmed that the planulae were asymbiotic. Nucleic acids extracted from planulae and then used as templates for the microsatellite loci primer sets failed to produce PCR amplicons, whereas nucleic acids from adult tissue that contained Symbiodinium or nucleic acids isolated from Symbiodinium cultures produced PCR amplicons (see below) of the correct size (given in Table 2). Taken together, the data demonstrate that the microsatellite loci are located within the genome of Symbiodinium, and not that of the gorgonian hosts.

Microsatellites of Symbiodinium within a host species
For the Symbiodinium populations of P. kuna, four unique alleles were identified at locus CA1'.7, three at GA2.8, and three at GA4.84. The sizes of the alleles at each locus were CA1'.7 (approximately 360, 380, 400, 420 bp), GA2.8 (approximately 180, 190, 200 bp), and GA4.84 (approximately 280, 300, 310 bp). In 147 cases, only a single allele per locus could be detected from a P. kuna colony, but the Symbiodinium populations from 13 colonies produced two distinct alleles at one or more loci. Thus, 8.1% of the Symbiodinium populations sampled from P. kuna individuals produced more than a single allele at any microsatellite locus.

When the Symbiodinium populations of P. elisabethae were screened at loci CA4.86 and CA6.38, 8 and 10 unique alleles, respectively, were identified from 568 of the 575 colonies that were examined (nucleic acids from seven colonies failed to amplify at either locus and were excluded from further analysis). Allele sizes for loci CA4.86 and CA6.38 ranged between 185–207 bp and 96–122 bp, respectively (Fig. 1). In most of these cases, only a single allele per locus could be detected from the Symbiodinium population of a P. elisabethae colony (Fig. 2). In 25 cases (4.4%), the Symbiodinium population from a P. elisabethae colony possessed two distinct alleles at one or more loci (example in Fig. 2). This pattern is similar to that observed for Symbiodinium microsatellites isolated and amplified from P. kuna.



View larger version (17K):
[in this window]
[in a new window]
  		Figure 1. Size distribution of alleles at two microsatellite loci: (a) CA4.86 and (b) CA6.38. Solid bars, Symbiodinium populations of Pseudopterogorgia elisabethae (in hospite); diagonal strip bars, Symbiodinium cultures. Values for P. elisabethae are derived from all samples, include those that possessed two distinct alleles or a null allele at one or more loci.

 


View larger version (71K):
[in this window]
[in a new window]
  		Figure 2. Symbiodinium populations from Pseudopterogorgia elisabethae: Polyacrylamide gel electrophoresis analysis of microsatellite alleles from two loci: (a) CA4.86 and (b) CA6.38. Number above lane represents individual samples of P. elisabethae. L represents DNA ladder lanes. Vertical arrows denote samples in which more than a single allele was amplified per locus.

 
Symbiodinium microsatellites between host species
Amplifications using the Symbiodinium microsatellite loci primer sets for one host species produced mixed results when applied to the Symbiodinium populations of the other host species. In many cases, the primers failed to produce an amplicon. However, when PCR amplification did occur, only a single allele was detected per locus (see fig. 4 of Santos et al., 2001, for examples).

Microsatellites from Symbiodinium cultures
The Symbiodinium microsatellite loci CA1'.7, GA2.8, and GA4.84 were not detected in any of the Symbiodinium cultures. However, the 45 cultures did amplify with at least one of the two Symbiodinium microsatellites primer sets isolated from P. elisabethae (Table 2). At loci CA4.86 and CA6.38, 5 and 6 alleles, respectively, were identified (Table 3, Fig. 1). Allele sizes for loci CA4.86 and CA6.38 ranged between 179–193 bp and 98–112 bp, respectively (Fig. 1). In each case, however, only a single allele was detected per locus.

Test for deviations from Hardy-Weinberg equilibrium
The observed genotype frequencies of the Symbiodinium populations inhabiting Pseudopterogorgia elisabethae from most of the 12 sites in the Bahamas were significantly different from those expected under Hardy-Weinberg equilibrium (Table 4). An excess of "homozygote" (genotypes with a single allele per locus) genotypes was present in the populations compared to the expected number based on allelic frequencies. In four cases, a single monomorphic allele was recovered from the population (Table 4).


View this table:
[in this window]
[in a new window]
  		Table 4 Significant deviation of observed genotype frequencies from those expected under Hardy-Weinberg equilibrium for Symbiodinium populations inhabiting Pseudopterogorgia elisabethae in the Bahamas

 

   Discussion
TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Literature Cited
 
Microsatellite loci isolated from members of Symbiodinium clade B were amplified by PCR to assess ploidy in these symbiotic dinoflagellates. From most Symbiodinium populations harbored by Plexaura kuna and Pseudopterogorgia elisabethae (690 out of 728 samples; 95% of the cases), only a single allele was recovered per locus, and all 45 Symbiodinium cultures possessed a single allele per locus. Furthermore, an excess of "homozygous," or single allele per locus, genotypes in the Symbiodinium populations of P. elisabethae violated Hardy-Weinberg equilibrium. Nonconformity to the prediction of Hardy-Weinberg equilibrium indicates that one or more of its assumptions are not met in the population. The assumptions include (1) the organism is diploid; (2) reproduction is sexual; (3) the population is infinitely large; (4) mating occurs randomly; (5) generations are nonoverlapping; and (6) the population is free of genetic drift, migration, mutation, and natural selection (reviewed in Hartl and Clark, 1989). The Symbiodinium populations of P. elisabethae may violate many of these assumptions, but the predominance of "homozygous" genotypes in the Symbiodinium populations harbored by P. kuna, and in all Symbiodinium cultures, strongly suggests that at least the first assumption is being violated. If Symbiodinium were diploid in the vegetative life stage, more "heterozygous" genotypes should have been sampled. A possible explanation for the lack of heterozygous genotypes is that the Symbiodinium microsatellite loci are located in an organellar (i.e., mitochondrial or chloroplast) genome. If so, a similar pattern of a single allele per locus would be observed. Although a microsatellite locus has been described from the chloroplast genome of a free-living dinoflagellate, Heterocapsa triquetra (Zhang et al., 1999), it is highly unlikely that all five Symbiodinium microsatellite loci were isolated from the organellar genomes. Taken together, our results demonstrate that members of Symbiodinium clade B, both cultured and in hospite, possess a haploid nuclear phase. Moreover, since the genus is monophyletic (see, for example, Rowan and Powers, 1992; LaJeunesse, 2001; Santos et al., 2002), our findings corroborate Blank’s (1987) speculation that haploidy exists in S. kawagutii and extend it to all members of Symbiodinium.

If the vegetative life stage of Symbiodinium is haploid, why was more than a single allele sometimes recovered per locus in hospite? Among the samples of P. kuna and P. elisabethae, about 8.1% and 4.4%, respectively, gave this result. In these cases, the recovery of more than a single allele at a microsatellite locus suggests that the colonies harbored at least two genotypes of symbiotic algae. It has been demonstrated, using a combination of culturing and molecular techniques, that P. kuna and P. elisabethae can harbor more than a single genotype of Symbiodinium simultaneously (Goulet and Coffroth, 1997, 2003; Coffroth et al., 2001; Santos et al., 2001). Our data support this conclusion while suggesting that it is uncommon for colonies of these gorgonians to harbor more than one Symbiodinium genotype, at least at detectable concentrations. For these uncommon colonies, it remains to be determined whether the additional genotypes represent relicts of the initial symbiont uptake by the newly settled asymbiotic planulae or reflect populations that are secondarily acquired and contribute to the host in other ways.

Phylogenetic support for haploidy in Symbiodinium and other dinoflagellates
Our conclusion that Symbiodinium is haploid is consistent with data from most of the other dinoflagellates. The life cycle of nearly all dinoflagellates examined to date is dominated by asexual reproduction of haploid vegetative cells (Pfiester and Anderson, 1987). This observation, coupled with the monophyly of dinoflagellates (Saldarriaga et al., 2001), suggests that all members of the dinoflagellates should possess a vegetative haploid nuclear phase. Interestingly, the vegetative cells of the red-tide dinoflagellate genus Noctiluca are thought to be diploid (Zingmark, 1970; Pfiester and Anderson, 1987). In Noctiluca, the first divisions of the gamete mother-cell nucleus are believed to be meiotic (Zingmark, 1970), which would imply diploidy in these dinoflagellates. Recently, the conclusion of diploidy in Noctiluca has been challenged (Schnepf and Drebes, 1993), but no definitive data have been presented to establish the ploidy of these dinoflagellates. We hypothesize, based on our knowledge of other dinoflagellates, that Noctiluca spp. possess a vegetative haploid nuclear phase. Analyses of microsatellite loci, such as we have done here for Symbiodinium, would be one way to test this hypothesis and settle the question of ploidy in Noctiluca.

A haploid nuclear phase in the dinoflagellates is consistent with that of their closest relatives. The Apicomplexa, obligate intracellular parasites of many vertebrate and invertebrate hosts, are thought to have evolved from, or shared a common ancestor with, the dinoflagellates (Wolters, 1991; Cavalier-Smith, 1993) about 395-929 Mya (Escalante and Ayala, 1995). The apicomplexan Plasmodium falciparum, one of the causative agents of human malaria, is haploid in its human host and only briefly diploid in its mosquito vector (Campbell, 1993; Conway et al., 1999). Other apicomplexans, such as Cryptosporidium parvum and Toxoplasma gondii, also possess a haploid nuclear phase (Costa et al., 1997; Feng et al., 2002). Given the close evolutionary relationship between the two groups, the ancestral state in the progenitor of the apicomplexans and dinoflagellates was probably haploidy.

Evidence for fine-scale specificity in associations between host and Symbiodinium
Surprisingly, primer sets designed for amplification of the Symbiodinium microsatellite loci in one host species produced mixed results when applied to the Symbiodinium populations of the other host species or to Symbiodinium cultures. For example, primers designed for the symbiont populations of P. kuna were not very successful in amplifying the Symbiodinium populations of P. elisabethae colonies or algal cultures derived from a variety of hosts (Santos et al., 2001; unpubl. data). Typically, the utility of a microsatellite system (i.e., microsatellite primer sets) decreases as the phylogenetic distance between the samples being screened increases (Schlotterer, 1998). In these experiments, however, most (768 out of 773; 99.4%) of the samples belonged to a single lineage, Symbiodinium B184. Therefore, the microsatellite primer sets should have worked on all members of the group. This failure to amplify alleles from closely related, but non-focal, Symbiodinium samples is probably due to mutational changes in the flanking regions of the microsatellite array. Mutations in these regions can lead to primer-template mismatch and thus to inhibition of the PCR reaction. In support of this hypothesis, we have sequenced alleles from loci CA4.86 and CA6.38 and found mutations, such as nucleotide substitutions and insertion-deletions (indels), in the microsatellite flanking regions from members of Symbiodinium B184, B211, and B223 (the evolution of Symbiodinium microsatellites will be discussed in a subsequent paper).

Although microsatellite alleles were not always recovered in the host species comparisons, an important conclusion can be drawn from these data. The specificity exhibited by the different microsatellite primer sets to the population of Symbiodinium from which they were designed and the consistent presence of "null," or absent, alleles in the other host species suggest that the two Symbiodinium B184 populations are genetically distinct from each another and specific to a given host species. The genetic differences are probably spread across the Symbiodinium genome, but at the minimum they are confined to mutations in the flanking regions of the microsatellite loci. Unfortunately, internal transcribed spacer (ITS) sequences—one of the most useful genetic markers for identifying Symbiodinium types (LaJeunesse, 2001)—are identical, or nearly so, in the two populations (Santos et al., 2001); thus other genetic markers are needed to elucidate the relationship between them. Nevertheless, P. kuna and P. elisabethae appear to associate preferentially with genetically distinct Symbiodinium B184 populations, which provides evidence for fine-scale host-Symbiodinium specificity in these gorgonian species.

Microsatellites and Symbiodinium diversity
Consistent with other studies (Schoenberg and Trench, 1980; Colley, 1984; Goulet and Coffroth, 1997, 2003; Baillie et al., 1998, 2000; Belda-Baillie et al., 1999), our microsatellite data suggest an enormous amount of genotypic diversity within Symbiodinium, as illustrated by the following example. At loci CA4.86 and CA6.38, a total of 10 and 12 unique alleles, respectively, were recovered from samples belonging to Symbiodinium B184. Pairing alleles from each locus, under the assumption that there are no restrictions against particular combinations of alleles, generates 120 unique genotypes of Symbiodinium B184. However, we feel that this is a conservative estimate for several reasons. First, a minimal number of microsatellite loci are being employed. Data from other polymorphic microsatellite loci would distinguish more genotypes within Symbiodinium B184. Second, some alleles for CA4.86 and CA6.38 are missing from the data set because they have not yet been sampled (Fig. 1). The inclusion of any of these alleles would generate up to 210 unique genotypes of Symbiodinium B184. Third, the Symbiodinium B184 populations of hosts such as P. kuna possess "null" alleles at these loci, suggesting an additional level of diversity within the group (see above). Last it is extremely unlikely that this high level of genotypic diversity is confined to Symbiodinium B184. Thus, microsatellites will doubtlessly uncover high levels of genotypic diversity in most, if not all, of the 16 Symbiodinium lineages recognized in cp23S-rDNA phylogenies (Santos et al., 2002, 2003).

Evidence for recombination in Symbiodinium
The finding that vegetative cells of Symbiodinium possess a haploid nuclear phase does not preclude recombination within the life cycle of these symbiotic dinoflagellates. For example, other haploid organisms maintain some form of recombination during their life cycle, including the green alga Chlamydomonas and members of the Acrasiomycota (cellular slime molds), the Bryophyta (mosses), the Pterophyta (ferns), the Apicomplexa, and the Dinophyceae (dinoflagellates) (Pfiester and Anderson, 1987; Campbell, 1993). In fact, the high allelic variability observed for allozymes (Schoenberg and Trench, 1980; Baillie et al., 1998; Belda-Baillie et al., 1999), random-amplified-polymorphic DNA (RAPDs) (Belda-Baillie et al., 1999; Baillie et al., 2000), and DNA fingerprints (Goulet and Coffroth, 1997, 2003) suggests extensive recombination in Symbiodinium (Baillie et al., 2000; reviewed in LaJeunesse, 2001). This evidence for recombination, taken together with our finding of haploidy, lends strong support to Symbiodinium life cycle (a), as proposed by Fitt and Trench (1983). However, questions pertaining to recombination in these enigmatic dinoflagellates, such as the factors that induce it and whether it occurs inside or outside a host, remain to be answered.


   Acknowledgments
 
We thank D. Brancato and J. Weaver for maintaining the Symbiodinium cultures, C. Gutierrez-Rodriguez for access to P. elisabethae samples and primers, Dr. H. R. Lasker (State University of New York at Buffalo) for help in collecting P. kuna and P. elisabethae, and Dr. R. A. Kinzie III (University of Hawaii; Hawaii Institute for Marine Biology) for access to Symbiodinium cultures. We are also grateful to the Kuna Nation and the Republic of Panama, the Bahamas Department of Fisheries, and the Florida Keys National Marine Sanctuary for permission to collect and export samples from Panama, the Bahamas, and Florida, respectively. The technical and logistical support of the staff and scientists of the Smithsonian Tropical Research Institute, the Don Gerace Research Center, the Keys Marine Lab, and the National Undersea Research Center in Key Largo is greatly appreciated. We also thank H.R. Lasker and two anonymous reviewers for comments that improved the manuscript. This research was supported by a National Science Foundation (NSF) Minority Graduate Fellowship (SRS), NSF grants OCE-95-30057 and OCE-99-07319 (MAC), and a Caribbean Marine Research Center/NURC grant and New York State Sea Grant (MAC and H.R. Lasker).


   Footnotes
 
Received 18 July 2002; accepted 28 November 2002.

* Present Address: Department of Biochemistry and Molecular Biophysics, University of Arizona, Tucson, Arizona 85721. Back


   Literature Cited
TOP
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Literature Cited
 

    Baillie, B. K., V. A. Monje, V. Silvestre, M. Sison, and C. A. Belda-Baillie. 1998. Allozyme electrophoresis as a tool for distinguishing different zooxanthellae symbiotic with giant clams. Proc. R. Soc. Lond. B 256: 1949–1956.
    Baillie, B. K., C. A. Belda-Baillie, V. Silvestre, M. Sison, A. V. Gomez, E. D. Gomez, and V. Monje. 2000. Genetic variation in Symbiodinium isolates from giant clams based on random-amplified-polymorphic DNA (RAPD) patterns. Mar. Biol. 136: 829–836.
    Bart-Delabesse, E., J. F. Humbert, E. Delabesse, and S. Bretagne. 1998. Microsatellite markers for typing Aspergillus fumigatus isolates. J. Clin. Microbiol. 36: 2413–2418.[Abstract/Free Full Text]
    Belda-Baillie, C. A., M. Sison, V. Silvestre, K. Villamor, V. Monje, E. D. Gomez, and B. K. Baillie. 1999. Evidence for changing symbiotic algae in juvenile tridacnids. J. Exp. Mar. Biol. Ecol. 241: 207–221.
    Bennett, P. 2000. Microsatellites. J. Clin. Pathol. Mol. Pathol. 53: 177–183.[Abstract/Free Full Text]
    Blank, R. J. 1987. Cell architecture of the dinoflagellate Symbiodinium sp. inhabiting the Hawaiian stony coral Montipora verrucosa. Mar. Biol. 94: 143–155.
    Brondani, C., R. P. V. Brondani, L. R. Garrido, and M. E. Ferreira. 2000. Development of microsatellite markers for the genetic analysis of Magnaporthe grisea. Genet. Mol. Biol. 23: 753–762.
    Campbell, N. A. 1993. Biology. 3rd ed. Benjamin/Cummings Publishing, San Francisco, CA.
    Cavalier-Smith, T. 1993. Kingdom Protozoa and its 18 phyla. Microbiol. Rev. 57: 953–994.[Abstract/Free Full Text]
    Chambers, G. K., and E. S. MacAvoy. 2000. Microsatellites: consensus and controversy. Comp. Biochem. Physiol. 126: 455–476.
    Ciofi, C., and M. W. Bruford. 1998. Isolation and characterization of microsatellite loci in the Komodo dragon Varanus komodoensis. Mol. Ecol. 7: 133–135.
    Coats, D. W. 2002. Dinoflagellate life-cycle complexities. J. Phycol. 38: 417–419.[ISI]
    Coffroth, M. A., H. R. Lasker, M. E. Diamond, J. A. Bruenn, and E. Bermingham. 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., S. R. Santos, and T. L. Goulet. 2001. Early ontogenetic expression of specificity in a cnidarian-algal symbiosis. Mar. Ecol. Prog. Ser. 222: 85–96.
    Colley, N. J. 1984. The cell biology of dinoflagellate symbiosis in a coelenterate. Ph.D. dissertation. University of California, Santa Barbara, 174 pp.
    Conway, D. J., C. Roper, A. M. J. Oduola, D. E. Arnot, P. G. Kremsner, M. P. Grobusch, C. F. Curtis, and B. M. Greenwood. 1999. High recombination rate in natural populations of Plasmodium falciparum. Proc. Natl. Acad. Sci. USA 96: 4506–4511.
    Costa, J. M., M. L. Darde, B. Assouline, M. Vidaud, and S. Bretagne. 1997. Microsatellite in the beta-tubulin gene of Toxoplasma gondii as a new genetic marker for use in direct screening of amniotic fluids. J. Clin. Microbiol. 35: 2542–2545.[Abstract]
    Escalante, A. A., and F. J. Ayala. 1995. Evolutionary origin of Plasmodium and other Apicomplexa based on rDNA genes. Proc. Natl. Acad. Sci. USA 92: 5793–5797.[Abstract/Free Full Text]
    Feng, X., S. M. Rich, S. Tzipori, and G. Widmer. 2002. Experimental evidence for genetic recombination in the opportunistic pathogen Cryptosporidium parvum. Mol. Biochem. Parasitol. 119: 55–62.
    Fitt, W. K., and R. K. Trench. 1983. The relation of diel patterns of cell division to diel patterns of motility in the symbiotic dinoflagellate Symbiodinium microadriaticum Freudenthal in culture. New Phytol. 94: 421–432.
    Glynn, P. W. 1996. Coral reef bleaching: facts, hypotheses and implications. Global Change Biol. 2: 495–509.
    Goulet, T. L., and M. A. Coffroth. 1997. A within colony comparison of zooxanthellae genotypes in the Caribbean gorgonian Plexaura kuna. Proc. 8th Int. Coral Reef Symp. 2: 1331–1334.
    Goulet, T. L., and M. A. Coffroth. 2003. Genetic composition of zooxanthellae between and within colonies of the octocoral Plexaura kuna, based on small subunit rDNA and multilocus DNA fingerprinting. Mar. Biol. DOI 10.1007/s00227-002-0936-0.
    Hartl, D. L., and A. G. Clark. 1989. Principles of population genetics. 2nd ed. Sinauer Associates, Sunderland, MA.
    Hughes, J. S., and S. P. Otto. 1999. Ecology and the evolution of biphasic life cycles. Am. Nat. 154: 306–320.[Medline]
    Kinzie, R. A. 1974. Experimental infection of aposymbiotic gorgonian polyps with zooxanthellae. J. Exp. Mar. Biol. Ecol. 15: 335–345.
    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.[ISI]
    Lewis, J., and L. Wolpert. 1979. Diploidy, evolution, and sex. J. Theor. Biol. 78: 435–438.
    Oliveira, R. P., N. E. Broude, A. M. Macedo, C. R. Cantor, C. L. Smith, and S. D. J. Pena. 1998. Probing the genetic population structure of Trypanosoma cruzi with polymorphic microsatellites. Proc. Natl. Acad. Sci. USA 95: 3776–3780.[Abstract/Free Full Text]
    Paquin, C. E., and J. Adams. 1983. Frequency of fixation of adaptive mutations is higher in evolving diploid than haploid yeast populations. Nature 302: 495–500.[Medline]
    Pfiester, L. A., and D. M. Anderson. 1987. Dinoflagellate reproduction. Pp. 611–648 in The Biology of Dinoflagellates, F.J.R. Taylor, ed. Blackwell Scientific Publications, London.
    Rowan, R., and D. A. Powers. 1991. Molecular genetic identification of symbiotic dinoflagellates (zooxanthellae). Mar. Ecol. Prog. Ser. 71: 65–73.
    Rowan, R., and D. A. Powers. 1992. Ribosomal RNA sequences and the diversity of symbiotic dinoflagellates (zooxanthellae). Proc. Natl. Acad. Sci. USA. 89: 3639–3643.[Abstract/Free Full Text]
    Rynearson, T. A., and E. V. Armbrust. 2000. DNA fingerprinting reveals extensive genetic diversity in a field population of the centric diatom Ditylum brightwellii. Limnol. Oceanogr. 45: 1329–1340.
    Saldarriaga, J. F., F.J.R. Taylor, P. J. Keeling, and T. CavalierSmith. 2001. Dinoflagellate nuclear SSU rRNA phylogeny suggests multiple plastid losses and replacements. J. Mol. Evol. 53: 204–213.[ISI][Medline]
    Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular Cloning: a Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
    Santos, S. R., D. J. Taylor, and M. A. Coffroth. 2001. Genetic comparisons of freshly isolated vs. cultured symbiotic dinoflagellates: implications for extrapolating to the intact symbiosis. J. Phycol. 37: 900–912.[ISI]
    Santos, S. R., D. J. Taylor, R. A. Kinzie III, M. Hidaka, K. Sakai, and M. A. Coffroth. 2002. Molecular phylogeny of symbiotic dinoflagellates inferred from partial chloroplast large subunit (23S)-rDNA sequences. Mol. Phylogenet. Evol. 23: 97–111.[ISI][Medline]
    Santos, S. R., C. Gutierrez-Rodriguez, and M. A. Coffroth. 2003. Phylogenetic identification of symbiotic dinoflagellates via length heteroplasmy in domain V of chloroplast large subunit (cp23S)-rDNA sequences. Mar. Biotechnol. (In press).
    Schlotterer, C. 1998. Microsatellites. Pp. 237–261 in Molecular Genetic Analysis of Populations: A Practical Approach, A. R. Hoelzel, ed. IRL Press, Oxford.
    Schnepf, E., and G. Drebes. 1993. Anisogamy in the dinoflagellate Noctiluca? Helgol. Meeresunters. 47: 265–273.
    Schoenberg, D. A., and R. K. Trench. 1980. Genetic variation in Symbiodinium (=Gymnodinium) microadriaticum Freudenthal, and specificity in its symbiosis with marine invertebrates. I. Isozyme and soluble protein patterns of axenic cultures of Symbiodinium microadriaticum. Proc. R. Soc. Lond. B 207: 405–427.
    Seutin, G., B. N. White, and P. T. Boag. 1991. Preservation of avian blood and tissue samples for DNA analyses. Can. J. Zool. 69: 82–92.
    Swofford, D. L., and R. B. Selander. 1981. BIOSYS-1: a FORTRAN program for the comprehensive analysis of electrophoretic data in population genetics and systematics. J. Hered. 72: 281–283.[Abstract/Free Full Text]
    Taylor, D. L. 1974. Symbiotic marine algae: taxonomy and biological fitness. Pp. 245–262 in Symbiosis in the Sea, W. B. Vernberg, ed. University of South Carolina Press, Columbia, SC.
    van der Verde, M., H. J. During, L. van de Zande, and R. Bijlsma. 2001. The reproductive biology of Polytrichum formosum: clonal structure and paternity revealed by microsatellites. Mol. Ecol. 10: 2423–2434.[Medline]
    Weeks, A. R., F. Marec, and J.A.J. Breeuwer. 2001. A mite species that consists entirely of haploid females. Science 292: 2479–2482.[Abstract/Free Full Text]
    Wolters, J. 1991. The troublesome parasites: molecular and morphological evidence that Apicomplexa belong to the dinoflagellate-ciliate clade. Biosystems 25: 75–83.[ISI][Medline]
    Zhang, Z., B. R. Green, and T. Cavalier-Smith. 1999. Single gene circles in dinoflagellate chloroplast genomes. Nature 400: 155–159.[Medline]
    Zingmark, R. G. 1970. Sexual reproduction in the dinoflagellate Noctiluca miliaris Suriray. J. Phycol. 6: 122–126.[ISI]



This article has been cited by other articles:


Home page
 Biol. Bull.Home page
N. L. Kirk, J. R. Ward, and M. A. Coffroth
Stable Symbiodinium Composition in the Sea Fan Gorgonia ventalina During Temperature and Disease Stress
Biol. Bull., December 1, 2005; 209(3): 227 - 234.
[Abstract] [Full Text] [PDF]

Home page
 Mol Biol EvolHome page
J. R. Reichman, T. P. Wilcox, and P. D. Vize
PCP Gene Family in Symbiodinium from Hippopus hippopus: Low Levels of Concerted Evolution, Isoform Diversity, and Spectral Tuning of Chromophores
Mol. Biol. Evol., December 1, 2003; 20(12): 2143 - 2154.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (19)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Santos, S. R.
Right arrow Articles by Coffroth, M. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Santos, S. R.
Right arrow Articles by Coffroth, M. A.
Related Collections
Right arrow Algae
Right arrow Cell Biology
Right arrow Cnidarians
Right arrow Evolution
Right arrow Population Biology
Right arrow Symbiosis
Right arrow Phylogeny

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS