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1 Marine Biology Research Division 0202, Scripps Institution of Oceanography, University of California San Diego, La Jolla, California 92093-0202
2 University of Guam Marine Laboratory, Mangilao, Guam 96923
3 Smithsonian Tropical Research Institute, Apartado 2072, Balboa, Republic of Panama
* To whom correspondence should be addressed. E-mail: rrowan{at}uog9.uog.edu
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Abstract |
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 TOP Abstract IntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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Abbreviations: RFLP, restriction fragment length polymorphism • srDNA, small subunit ribosomal RNA gene • YBD, yellow-blotch disease
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Introduction |
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TOPAbstractIntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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In contrast to this view, we found that individual colonies of coral in the Montastraea annularis species complex often contain more than one taxon of Symbiodinium (Rowan and Knowlton, 1995; Rowan et al., 1997; Toller et al., 2001). At Aguadargana reef in the San Blas Islands of Panama (see fig. 1 in Toller et al., 2001), colonies of M. annularis host Symbiodinium B (or rarely, Symbiodinium A) in tissues exposed to high irradiance, and they host Symbiodinium C in tissues exposed to low irradiance. Colonies of M. faveolata exhibit a similar pattern except that Symbiodinium A and B are both common at high irradiance in these corals (Rowan and Knowlton, 1995; Rowan et al., 1997). Nearby at Río Cartí (a near-shore habitat; see fig. 1 in Toller et al., 2001), members of the Montastraea annularis complex host Symbiodinium E in tissues exposed to high irradiance and host Symbiodinium C otherwise (Toller et al., 2001). Thus on these two reefs, corals at shallower depths, which experience both high (on the colony top, exposed to downwelling irradiance) and low (on colony sides) irradiance, typically host both high- (Symbiodinium A and/or B or Symbiodinium E) and low- (Symbiodinium C) irradiance-associated zooxanthellae simultaneously. (On another offshore reef, Symbiodinium E also occurs in some of the deepest colonies of M. franksi, possibly as a result of sediment-associated stress [Toller et al., 2001]).
Several observations suggest that interactions among different taxa of Symbiodinium within one colony of Montastraea may be dynamic. First, coral growth causes slow changes in irradiance microenvironments (e.g., corallites moving from tops to sides of M. annularis columns), and the specificity of different zooxanthellae for different irradiance environments (above) implies that zooxanthellar communities will change in response to these irradiance changes. Second, experimental manipulations of irradiance gradients within colonies of M. annularis hosting Symbiodinium B and C resulted in changes in the distribution of these zooxanthellae (Rowan et al., 1997). Finally, the proportions of Symbiodinium A, B, and C in Montastraea changed during a coral bleaching event (Rowan et al., 1997).
The present study tested the ability of zooxanthellar symbioses in M. annularis and in M. faveolata to reestablish typical patterns of association after being disturbed. Because zooxanthellae in unmanipulated corals have such environmentally predictable patterns of distribution (above), we hypothesized that disturbed zooxanthellar populations would re-establish the same patterns of association, directly. To disturb zooxanthellae, we treated corals with low light (e.g., Franzisket, 1970) or with high light (e.g., Dustan, 1979), both of which caused corals to lose zooxanthellae (to bleach). Corals were then allowed to recover. We also studied the zooxanthellar communities of unmanipulated corals that exhibited yellow-blotch disease and associated reductions in zooxanthellar numbers.
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Materials and Methods |
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TOPAbstractIntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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A second experiment was done (starting in January 1998) at a different location (ca. 0.5 km away). The second experiment differed from the first one only as follows: five colonies (15 coral columns) of M. annularis were used, treatment was for 4 rather than 7 weeks, Recovery samples were obtained after 13 rather than 17 weeks, and Final samples were obtained (from all columns) after a total of 33 rather than 37 weeks. Experiment II (Fig. 1B). In October 1997, one core was removed from each of 12 large (> 1.5 m tall and wide) colonies of M. faveolata living at depths of 1–9 m, using a pneumatic drill fitted with a 44-mm hole saw (resulting cores had ca. 12.6 cm2 of live tissue and were ca. 5 cm in height). At this time, tissue samples (Initial samples) were taken immediately adjacent to the coring sites with a steel hole punch (#12). Coral cores were then transplanted among three small caves (Low Light, as above) on the reef (7–11 m depth), where they were secured with plastic cable ties to masonry nails pounded into reef framework. Cores occupied the back (darkest) portion of the caves and were mounted upside-down on the cave ceilings.
After 6 weeks of low-light treatment in caves, coral cores were collected and assessed visually; tissue samples were taken from each coral core at a haphazardly selected location away from the core’s perimeter (#6 hole punch; Treatment samples). Cores were then attached to cleared reef substrate at 9 m depth with epoxy putty (Z-Spar Splash Zone, Kop-Coat, Inc., Pittsburgh, PA), facing upward under unobstructed natural irradiance. After 8 weeks in this common garden and then again after another 16 weeks, coral cores were assessed and sampled again (#6 hole punch; Recovery samples and Final samples, respectively). In the latter case (Final samples), only eight coral cores were sampled—the four others were lost. Experiment III (Fig. 1C). In January 1998, individual columns of M. annularis were each half-covered with a shield of aluminum foil. This treatment bisected each column vertically into two morphologically equivalent halves, one of which was covered by foil and therefore low-light treated (Treated) and the other of which was exposed to natural irradiance (Untreated). Each shield was molded to its column, lifted off slightly (<0.5 cm), and secured to the column’s nonliving base with nylon cable ties. A shield was placed on one column of each of 15 colonies living at depths of 2–4 m (shallow group), and on one column of each of 15 colonies living at depths of 7-9 m (deep group). Shields were removed after 4 weeks, at which time treatment boundaries were marked by gently tapping two small steel nails into opposite sides of each column. After another 12 weeks, columns were assessed visually and a pair of tissue samples was taken from the top of each, 2 cm apart and on either side of the treatment boundary (#6 hole punch; Treated and Untreated samples, Fig. 1C).
Yellow-blotch disease
In October 1997 and January 1998 at Cayos Limones, San Blas (see fig. 1 in Toller et al., 2001), we found a number of colonies of Montastraea that appeared to have "yellow-blotch disease" (YBD; Santavy et al., 1999). Some colonies had only one or two small lesions (ca. 10–30 cm wide), which usually consisted of a patch of exposed skeleton surrounded by a halo (typically ca. 1–3 cm wide) of yellow living tissue, which in turn was surrounded by a halo (typically 
2 cm wide) of white (bleached) tissue; lesions were surrounded by apparently healthy tissue (see Fig. 1D). Other colonies were mostly dead, in which case a patch of normal tissue was surrounded by a band of bleached tissue inside a band of yellow tissue.
Using a steel hole punch (#12), we took samples from five colonies of M. franksi (one lesion per colony), from six colonies of M. faveolata (one or two lesions per colony), and from one colony of M. annularis (two lesions). Two samples were taken at every lesion—one of normally pigmented tissue and one of yellow tissue nearby (3 cm apart; Normal and Yellow, respectively; see Fig. 1D). At five lesions we also sampled the white tissue that was between yellow and normal tissue (White; see Fig. 1D).
Progression of YBD was monitored in 12 colonies of M. faveolata. On 25 January 1998, two small nails were driven into the bare skeleton next to one YBD lesion in each colony. The two nails defined a line parallel to the lesion edge, and the distance between that line and the lesion edge (living, yellow tissue) was measured with a pair of calipers. We also measured the distance to normally pigmented tissue, along the same vector. These measurements were repeated 5 months later (28 May 1998).
Laboratory methods
Zooxanthellae were isolated from frozen samples as described previously (Rowan and Powers, 1991b; Rowan and Knowlton, 1995), except that skeletal cores or fragments, after being stripped of tissue, were broken apart with a steel spatula and then washed with isolation buffer. That wash was combined with the tissue that had been stripped from the sample previously. At this point, one-tenth of each sample was fixed in 10% formalin and stored at 4 °C for cell counts, which were obtained from eight subsamples of each sample by hemacytometry. The rest of each sample was used to prepare DNA as described previously (Rowan and Powers, 1991b; Rowan and Knowlton, 1995).
Zooxanthellae in each sample were identified by restriction fragment length polymorphism (RFLP) genotypes of small ribosomal subunit RNA genes (srDNA), as described previously (Rowan and Powers, 1991b; Toller et al., 2001). Each sample was analyzed at least twice—once by srDNA amplification with universal PCR primers (ss3 and ss5; Rowan and Powers, 1991b), and once by srDNA amplification with host-excluding PCR prim (ss3Z and ss5; Rowan and Powers, 1991b; Toller et al., 2001). All amplified srDNAs were digested with Dpn II and with Taq I, and then compared to standard srDNA genotypes of Symbiodinium A, B, C, and E (srDNA clones A0, B0, C0, and E0–1, amplified and digested the same way; see Toller et al., 2001). RFLP genotypes C2 (in two samples) and C (in 12 samples) were compared in greater detail using a total of 12 restriction enzymes: Alu I, BstN I, BstU I, Dpn II, Hae III, Hha I, Hinf I, Mbo I, Mse I, Msp I, Sau96 I, and Taq I.
Samples that contained more than one zooxanthellar RFLP genotype were compared to a series of synthetic mixtures of cloned srDNAs (srDNA clones A0, B0, C0, and E0–1 and srDNA clone C20–1, see below) to estimate the relative abundance of each genotype in the sample (Rowan et al., 1997; Toller et al., 2001). For graphical presentation, these estimates were multiplied by the sample’s total cell number (see above) to estimate the number of cells of each genotype in the sample, and these values were then converted to numbers of cells per square centimeter of live coral surface (number of zooxanthellae/cm2 in Figs. 3– 7).
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srDNA with an RFLP genotype distinct from Symbiodinium A0, B0, C0, and E0–1, here called C2, was cloned and sequenced using methods described previously (Toller et al., 2001). It was amplified with host-excluding PCR primers from a colony of M. annularis in Experiment III (clone C20–1) and from an unmanipulated colony of the coral Siderastrea siderea (clone C20–2). DNA sequences were deposited in GenBank [http://www.ncbi.nlm.nih.gov/; accession numbers AF238259 (C20–1), AF238260 (C20–2)].
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Results |
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srDNA of genotype C2 appears to lack a Dpn II restriction site relative to srDNA of genotype C (Fig. 2). Defined by this character, genotype C2 was found previously in various other species of host (R. Rowan and W. Toller, unpubl. obs.). Cloned srDNAs C0 (which represents Symbiodinium C; Toller et al., 2001), C20–1 (genotype C2 from Experiment III), and C20–2 (genotype C2 from Siderastrea siderea, collected nearby) differed from one another in nucleotide sequence by about 0.9% (not shown; see also Toller et al., 2001). That amount of srDNA sequence difference could imply that these three clones represent three species of Symbiodinium (e.g., McNally et al., 1994), or one species of Symbiodinium in which srDNA is heterogeneous (see Toller et al., 2001).
We further compared the two samples from which clones C20–1 and C20–2 were obtained to 12 samples of Symbiodinium C (samples in fig. 9 in Toller et al., 2001) by digesting srDNAs with 12 restriction enzymes (listed in Materials and Methods). The two samples of genotype C2 were indistinguishable and differed from Symbiodinium C only in Dpn II (above) and Mse I digests (not shown). This analysis also showed that srDNA was heterogeneous in all samples, which means that zooxanthellae in the samples cannot be described precisely by sequences of cloned srDNA (i.e., clones C0, C20–1, and C20–2; see above and Toller et al., 2001). Nevertheless, RFLP data indicate that genotype C2 represents a taxon of zooxanthella, Symbiodinium C2, that is distinct from the Symbiodinium C that occurs commonly in M. annularis and M. faveolata.
Examples of RFLP genotypes that we interpreted as mixtures of taxa of Symbiodinium are shown on the right side of Figure 2 (Samples). The figure compares data obtained by amplifying srDNAs with host-excluding (Fig. 2A) versus universal (Fig. 2B) PCR primers because, using Dpn II and Taq I, both sets of data are needed to distinguish mixtures of genotypes A and C2 (e.g., Fig. 2, A > C2 and C2 > A) from mixtures of genotypes A, C2, and E (e.g., Fig. 2, E > A
C2). Universal PCR primers also amplify coral host srDNA (Fig. 2B, clone H0), but none was detected in the samples of zooxanthellae shown in Figure 2B.
Experiment I
The experimental units were individual coral columns taken from 15 donor colonies of M. annularis (see Fig. 1A). Below, a column is identified by the colony from which it came and by its treatment group (e.g., Colony 1, High Light). Columns 1–10 were in the first experimental group; Columns 11–15 were in the second group (see Materials and Methods). Before treatment, samples from most coral columns contained Symbiodinium C (Fig. 3, Initial). Exceptions were two donor colonies that yielded only Symbiodinium B (Fig. 3, Initial; Colonies 14 and 15 in all treatment groups), and one column that yielded Symbiodinium B with a small amount of Symbiodinium E (Fig. 3, Initial; Colony 7, Low Light). Initial zooxanthellar numbers in the three treatment groups (Control, High Light, and Low Light; Fig. 3) were indistinguishable (31.5 ± 10.1 x 105, 31.9 ± 9.80 x 105, and 31.4 ± 7.10 x 105 zooxanthellae/cm2 of coral, respectively).
Zooxanthellar numbers decreased after treatment. Samples from high-light-treated coral columns (Fig. 3, High Light, Treatment) had, on average, 29% as many zooxanthellae as did samples from controls (10.7 ± 9.0 x 105 versus 36.6 ± 17.5 x 105 zooxanthellae/cm2 [means ± standard deviations]; Wilcoxon signed rank test, P < 0.001). Two high-light-treated columns (Colonies 14 and 15; with Symbiodinium B before treatment) appeared normal; the other 13 (with Symbiodinium C before treatment) were pale or bleached, but only on their tops and south-facing (sun-facing) sides. All low-light-treated columns (Fig. 3, Low Light, Treatment) were white, and samples from them had, on average, only about 2.5% as many zooxanthellae as did samples from controls (0.90 ± 1.4 x 105 versus 36.6 ± 17.5 x 105 zooxanthellae/cm2; Wilcoxon signed rank test, P < 0.001). Zooxanthellar identities did not change in samples from the tops of coral columns in the control group after 4 or 7 weeks (Fig. 3, Control, Treatment versus Initial), with one exception. That exception was Colony 14, which yielded only Symbiodinium B initially but yielded Symbiodinium B with a small amount of Symbiodinium E 4 weeks later. In high-light-treated columns one change was observed (Fig. 3, High Light, Treatment versus Initial): Colony 5 initially yielded Symbiodinium C but yielded roughly equal parts of Symbiodinium C and Symbiodinium E immediately after treatment.
Identities of zooxanthellae were difficult to determine in low-light-treated coral columns at the end of treatment, presumably because these columns contained so few zooxanthellae (above). Two rounds of PCR amplification (see Materials and Methods) allowed 13 samples to be analyzed (Fig. 3, Low Light, Treatment); no srDNA was obtained from the other two samples. Six samples contained Symbiodinium C, one contained Symbiodinium E (Colony 7, which contained some Symbiodinium E initially), and six yielded only a non-Symbiodinium RFLP genotype (genotype N; see above).
After a total time of 17 or 24 weeks, zooxanthellar numbers and RFLP genotypes in samples from the control group were similar to initial conditions; Colony 14 once again yielded only Symbiodinium B (Fig. 3, Control; Initial, Treatment, Recovery). High-light-treated columns, which had then spent 13 or 17 weeks in their original, deeper environment (Fig. 3, High Light, Recovery) regained coloration (11 normal, 4 pale on tops only) and zooxanthellar numbers (Recovery versus Treatment: 45.4 ± 34.2 x 105 versus 10.7 ± 9.0 x 105 zooxanthellae/cm2; Wilcoxon signed rank test, P < 0.001). At this time, zooxanthellar numbers in samples from high-light-treated columns were similar to those in samples from controls (45.4 ± 34.2 x 105 versus 35.2 ± 18.5 x 105 zooxanthellae/cm2, respectively; Wilcoxon signed rank test, P > 0.1).
Thirteen or 17 weeks after the end of treatment, 10 of 15 high-light-treated coral columns had the same taxa of Symbiodinium that they contained before treatment, but 5 of 15 coral columns apparently contained different taxa of Symbiodinium than they began with (Fig. 3, High Light, Recovery). One of these taxonomic differences had been observed at the end of treatment (Colony 5). The other four changes (Fig. 3, High Light, Initial versus Recovery) were Symbiodinium C to Symbiodinium A and C2 (Colony 1); Symbiodinium C to Symbiodinium A (Colony 2); Symbiodinium C to Symbiodinium C, A, and E (Colony 3); and Symbiodinium C to Symbiodinium C and E (Colony 4). These four columns represented four of the five columns with the lowest numbers of zooxanthellae following treatment.
All low-light-treated coral columns experienced major changes in zooxanthellar populations after 13 or 17 weeks back in their original environment (Fig. 3, Low Light, Recovery). Zooxanthellar numbers were about 100-fold higher than after treatment (Recovery versus Treatment: 111 ± 45.2 x 105 versus 0.90 ± 1.4 x 105 zooxanthellae/cm2; Wilcoxon signed rank test, P < 0.001), and were about 3-fold higher than in the control group (35.2 ± 18.5 x 105 zooxanthellae/cm2; Wilcoxon signed rank test, P < 0.001). Only Colony 2 appeared normal; the other 14 columns, despite their large numbers of zooxanthellae, were still pale at this time (May 1998).
No low-light-treated coral column contained the same zooxanthellae that it had originally (Fig. 3, Low Light, Recovery versus Initial). Eight of them contained mixtures of taxa, and Symbiodinium A was predominant, followed by Symbiodinium E and C2; Symbiodinium C and B were not detected (Fig. 3, Low Light, Recovery). The predominance of Symbiodinium A was observed primarily in Colonies 11–15 (sampled 13 weeks after treatment); among Colonies 1–10 (sampled 17 weeks after treatment) only four samples contained more than 50% Symbiodinium A.
Twenty-four of the above coral columns (n = 8 colonies) were sampled again, for the last time, 33 or 37 weeks after treatment. At this time (18 October 1998), many unmanipulated colonies of M. annularis, M. faveolata, and M. franksi living at depths of 8–14 m at our study site were pale or bleached on their upper surfaces. Weekly mean sea-surface temperatures near our study site ranged between 29.4 °C and 29.9 °C from 26 August to 7 October (9.5 °N, 78.5 °W; data from Integrated Global Ocean Services System, http://ingrid.ldgo.columbia.edu/SOURCES/.IGOSS). Historically, temperatures this high are associated with coral bleaching at our study site (see fig. 3e in Rowan et al., 1997). Moreover, most of the summer of 1998 was unusually warm: from the first week of May through the first week of October (23 weeks) in the years 1981 to 1997 (but excluding 1983 and 1995, when corals bleached), there were an average of 3.5 weeks of average sea-surface temperature at or above 29.0 °C near our study site; for this period in 1998, there were 16 such weeks (data from IGOSS, as above). However, bleaching appeared to be less severe than in October 1995 (Rowan et al., 1997; pers. obs.) in terms of the number of colonies of Montastraea affected, the number of species of coral affected, and the extent to which individual corals were bleached.
Results are presented in Figure 4 (Final), in comparison to Recovery samples (13 or 17 weeks after treatment) from the same columns (data from Fig. 3). When final samples were collected, six of eight control coral columns appeared pale or slightly pale [labeled (P) and (SP), respectively, in Fig. 4] on top, as were six of eight high-light-treated columns (Fig. 4, Final); the other two columns in each group hosted Symbiodinium B and appeared normal [labeled (N)]. Twenty weeks earlier (Recovery) only one of these 16 columns appeared pale (Colony 1, High Light) and all others appeared normal. As suggested by the increase in numbers of pale colonies, average zooxanthellar numbers decreased and were about 5-fold lower than in the previous samples (Final versus Recovery: control columns, 6.7 ± 4.0 versus 35.2 ± 18.5 x 105 zooxanthellae/cm2; high-light-treated columns, 9.4 ± 4.3 versus 45.4 ± 34.2 x 105 zooxanthellae/cm2; Wilcoxon signed rank tests, P = 0.01).
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Final samples from control columns contained the same taxa of Symbiodinium that were observed previously in those columns (Fig. 4; Control, Recovery). This was also true for most high-light-treated columns, although in one of these Symbiodinium B was found with Symbiodinium C (Fig. 4, High Light, Final; Colony 7), whereas only Symbiodinium C was detected in that column previously. In contrast, we found different zooxanthellae (relative to Recovery) in six of eight low-light-treated columns (Fig. 4, Low Light, Final versus Recovery). In final samples from low-light-treated corals overall, Symbiodinium A declined, Symbiodinium E increased to become predominant, and Symbiodinium C2 appeared in different columns, compared to samples taken 20 weeks earlier. Symbiodinium C and B were not detected.
Experiment II
Cores taken from colonies of M. faveolata (see Fig. 1B; these cores are hereafter referred to as "corals") living at depths between 1 and 9 m contained a variety of zooxanthellar taxa before treatment (Symbiodinium A, B, C; Fig. 5, Initial), as expected based on earlier surveys (Rowan and Knowlton, 1995; Rowan et al., 1997). After low-light treatment, all 12 corals were white and contained, on average, about 2.6% as many zooxanthellae as they began with (Fig. 5, Treatment versus Initial; 0.8 ± 1.0 x 105 versus 30.8 ± 10.2 x 105 zooxanthellae/cm2, respectively; Wilcoxon signed rank test, P = 0.002). Zooxanthellae were identified in only two samples; both contained Symbiodinium A, and were from corals that had mixtures of A and B before treatment (Corals 3 and 4; Fig. 5).
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Eight corals were sampled after a further 16 weeks in their common garden (24 weeks total time after the end of treatment, at the end of May 1998, prior to the bleaching event noted above). Six of them appeared normal and two (Corals 5 and 6) were pale. Zooxanthellar numbers remained high on average (111.1 ± 55.6 x 105 zooxanthellae/cm2). Samples from six corals contained only Symbiodinium A; samples from the other two corals contained Symbiodinium A and E (Fig. 5, Final).
Experiment III
When the foil treatment shields (see Fig. 1C) were removed, all treated tissues were white. Adjacent tissues that had not been covered appeared normal, and borders between the white (treated) tissue and the normal (untreated) tissue were sharp. Many corals had suffered partial mortality in covered areas during treatment; further observations were made only on those in which more than 50% of the treated tissue appeared healthy (n = 5 in the shallow group; n = 9 in the deep group). To avoid additional stress to the corals, no samples were taken immediately after treatment.
The corals had different appearances 12 weeks after treatment. In four shallow corals and one deep coral (Fig. 6, asterisks; Corals 1, 2, 3, 5, and 6) it appeared that normal pigment had spread from untreated into treated tissue by about 2–3 cm, so that treatment boundaries were no longer apparent. In these corals, samples taken from either side of the treatment boundary (see Fig. 1C, Treated and Untreated) were normally pigmented; they also had similar numbers of zooxanthellae, of the same taxon of Symbiodinium. The taxa were those expected in shallower (Corals 1, 2, 3, 5; Symbiodinium B or A) and deeper (Coral 6; Symbiodinium C) colonies of M. annularis at this location (Rowan and Knowlton, 1995; Rowan et al., 1997).
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Disease-associated disturbance of zooxanthellae
We marked YBD lesions in 12 colonies of M. faveolata and observed that mortality progressed by 11 ± 6 mm (mean ± standard deviation) during 5 months. Yellow and white halos (see Fig. 1D) progressed in concert with mortality. Thus, as YBD spreads across a coral, it appears that tissue first loses zooxanthellae (white), then partially recovers zooxanthellae (yellow), and then dies. Average numbers of zooxanthellae in samples of normal (31.8 ± 10.1 x 105 zooxanthellae/cm2), white (3.7 ± 1.8 x 105 zooxanthellae/cm2), and yellow (24.6 ± 16.1 x 105 zooxanthellae/cm2) tissues confirmed that hypothesis.
With one exception (Fig. 7; Colony 9), normal and yellow samples from the same lesion contained different taxa of Symbiodinium. Samples of normal tissues contained the taxa that are common in unaffected corals at these depths (Rowan and Knowlton, 1995; Rowan et al., 1997; Toller et al., 2001)—predominantly Symbiodinium C—and yellow tissues contained predominantly Symbiodinium A (estimated at 
50% of the total in samples from 12 of 15 lesions; Fig. 7). Yellow tissue also often (9 of 15 samples; Fig. 7) contained the zooxanthellae found in the adjacent normal tissue. We could identify zooxanthellae in two samples of white tissue (Colony 3 and Lesion 7-1, Fig. 7); they contained mixtures of the taxa found in the adjacent normal (Symbiodinium C) and yellow (Symbiodinium A) tissues.
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Discussion |
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TOPAbstractIntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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Our study compared zooxanthellae that were identified by Dpn II- and Taq I-generated RFLP genotypes of srDNA, and this method does not discriminate all species of Symbiodinium (Rowan, 1998; Toller et al., 2001). For example, three known species of Symbiodinium A—S. microadriaticum (GenBank accession number M88521), S. pilosum (X62650), and S. corculorum (L13717)—would be indistinguishable in this analysis. Nevertheless, where the same zooxanthellar RFLP genotype was detected in a coral both before and after treatment (e.g., Fig. 3, High Light, Colonies 6–15), parsimony argues that the coral hosted the same species of Symbiodinium throughout the experiment. On the other hand, no taxonomic uncertainty affects our observation that many re-established symbioses involved changes; when compared samples of zooxanthellae differed with respect to RFLP genotypes A, B, C, or E (e.g., Fig. 3, Low Light; Fig. 5), it is clear that the samples contained different species of Symbiodinium (Rowan, 1998; Toller et al., 2001).
Disturbance and re-establishment of zooxanthellar symbioses
Where our experiments were conducted, Symbiodinium B predominates in Montastraea annularis at higher irradiance, Symbiodinium A and B predominate in M. faveolata at higher irradiance, and Symbiodinium C predominates in both species of coral at lower irradiance (Rowan and Knowlton, 1995; Rowan et al., 1997). This predictable pattern suggests that host-symbiont specificity is defined largely by the interaction of each host-zooxanthella combination with its environment (sensu Buddemeier and Fautin, 1993). This led us to hypothesize that, under a constant environment, host-symbiont specificity should be directly re-established following acute disturbance.
We found some evidence for this in Experiment I, in that most (10 of 15) of high-light-treated columns of M. annularis were repopulated with the same zooxanthellae that existed prior to treatment (Symbiodinium C or B; Fig. 3, High Light, Colonies 6–15). However, the host-zooxanthella specificity was not re-established in the other high-light-treated columns, which contained at least some different zooxanthellae (Symbiodinium A, E, and C2; Fig. 3, High Light, Colonies 1–5) after recovery. In these five corals, treatment had led to significantly fewer zooxanthellae than in the other 10 corals (5.0 x 105 cells/cm2 vs. 13.6 x 105 cells/cm2, respectively; P < 0.05, Mann-Whitney test). This suggests that coral-zooxanthella associations may or may not be re-established following disturbance, depending on the magnitude of zooxanthellar depletion.
This conclusion is supported by the results of Experiment I, in which none of the previously observed coral-zooxanthella associations were re-established in low-light-treated M. annularis. Zooxanthellae were severely depleted in these corals during treatment (to ca. 1 x 105 cells/cm2 on average), and all corals were repopulated by completely different zooxanthellae (Symbiodinium A, E, and C2; Fig. 3, Low Light, Recovery), even 9 months after treatment (Fig. 4). A similar result was obtained by low-light treatment of M. faveolata in Experiment II, although in that experiment the re-establishment of symbioses was not tested under a constant environment; after being treated with low light, most corals were also transplanted to a new environment. In that new environment (9 m depth), unmanipulated M. faveolata host Symbiodinium C (Rowan and Knowlton, 1995; unpubl. obs.), whereas re-established symbioses involved Symbiodinium A or A and E (Fig. 5).
We hypothesized that new taxa (Symbiodinium A, E, or C2) following severe bleaching (Experiments I and II) would not become established if untreated zooxanthellae (resident Symbiodinium B or C populations) were abundant near bleached tissues (zooxanthellae are thought to be translocated within colonies, among coral polyps, via their gastrovascular systems [e.g., Gladfelter, 1983; Gateño et al., 1998]). In Experiment III, the results from 5 of the 14 half-bleached columns were consistent with this hypothesis: bleached tissues were repopulated with zooxanthellae that apparently originated from untreated tissues (Fig. 6, Corals 1, 2, 3, 5, 6). Together with the observations on the spread of pigmentation (Fig. 6, Results), this indirect evidence suggests that zooxanthellae are translocated into bleached tissues in some cases. However, in the majority of tested cases (9 of 14), new zooxanthellae did become established: Symbiodinium A, E, and/or C2 repopulated treated tissues (Fig. 6, Corals 4, 7–14), despite the proximity (7 polyps away; see Weil and Knowlton, 1994) of untreated zooxanthellae (Symbiodinium C, with one exception). When new zooxanthellae became established, they were observed more frequently in the deeper habitat (8 of 9 columns in the deep group vs. 1 of 5 in the shallow group), and when resident zooxanthellae were Symbiodinium C (rather than Symbiodinium B)—our data do not resolve which factor had the greater influence. Nevertheless, these observations clearly show that a reservoir of adjacent zooxanthellae, whether Symbiodinium C or B, is not sufficient to prevent the establishment of new host-zooxanthella associations in bleached tissues.
We do not know where the new Symbiodinium in re-established symbioses came from. For most experimental corals, the fact that these zooxanthellae were not detected initially or after treatment is not good evidence that they were truly absent. This is because an srDNA genotype must be at least 5% of the total to be detected reliably (e.g., for values of ca. 12%, see fig. 4 in Toller et al., 2001; fig. 2B in Rowan et al., 1997). Thus, corals that had ca. 1 x 105 cells/cm2 of Symbiodinium C after low-light treatment (Fig. 3, Low Light, Treatment; also see Results) also may have contained up to about 5 x 103 cells/cm2 of Symbiodinium A, E, or C2 that went undetected. With a hypothetical doubling time of 5 days (e.g., Wilkerson et al., 1988), 5 x 103 zooxanthellae/cm2 become 150 x 105 zooxanthellae/cm2 after only 8 weeks. Thus, even where only Symbiodinium C was detected right after treatment (Fig. 3, Low Light, Treatment; Colonies 1, 3, 4, 11, 12, and 14), there might have been enough Symbiodinium A, E, and/or C2 present to found the established symbioses observed 13 or 17 weeks later.
On the other hand, no data show that treated corals did not acquire Symbiodinium A, E, and C2 for the first time during recovery. Free-living Symbiodinium may be attracted specifically to hosts lacking zooxanthellae (Fitt, 1985b). Juveniles of host species that do not transmit zooxanthellae vertically (e.g., Montastraea; Szmant, 1991) must be colonized, and the ability of adult hosts to pick up Symbiodinium from the environment has been documented for bleached anemones (Kinzie et al., 2001) and for juvenile giant clams originally inoculated with cultured zooxanthellae (Belda-Baillie et al., 1999; also see Fitt, 1984).
Regardless of where Symbiodinium A, E, and C2 came from, they fared well compared to any Symbiodinium C or B that remained in corals after low-light treatment. For example, the about 1 x 105 cells/cm2 of Symbiodinium C that six corals in Experiment I contained after treatment (Fig. 3, Low Light, Treatment; Colonies 1, 3, 4, 11, 12, and 14) would have been observed in re-established symbioses if they had doubled only three or four times during 13 or 17 weeks (doubling times of 23–29 days; a slow rate of growth for zooxanthellae in general [Wilkerson et al., 1988]). Thus, the identities of re-established symbioses in these six corals resulted not only from the proliferation, acquisition, or both of Symbiodinium A, E, or C2, but also from the failure of Symbiodinium C to proliferate.
Competition and succession in zooxanthellar communities
In general, the first phototrophs to colonize disturbed habitat are transient and eventually replaced by competitively superior species that dominate thereafter. This process is called succession (Odum, 1969; Connell and Slatyer, 1977; Huston and Smith, 1987), and it might eventually have led from Symbiodinium A, E, and C2 to Symbiodinium C or B, and thus restored the host-symbiont specificity observed in nature. We did not observe this hypothetical succession of zooxanthellae. However, the only corals we followed for more than 17 weeks after treatment (Fig. 4, Final; 33 or 37 weeks after treatment) experienced a natural bleaching event (Results) that apparently reduced populations of Symbiodinium C by about 80% in control and high-light-treated M. annularis; Symbiodinium B may have been affected also (Fig. 4, Final vs. Recovery). It seems unlikely that Symbiodinium C or B would have proliferated in low-light-treated corals during the same period of time. Thus, unfavorable conditions might explain why succession was not observed.
Hypotheses on the mechanisms of plant succession invoke genetic differences in the abilities of species to compete for resources such as water, light, and nutrients, supplies of which decrease as succession proceeds (e.g., Huston and Smith, 1987; Tilman, 1988; Wilson, 1999). Zooxanthellae in unmanipulated corals cannot be water-limited, nor can they extensively shade one another (Drew, 1972), but they probably are nutrient-limited (Rees, 1991; Falkowski et al., 1993). In contrast, severely bleached corals may be nutrient-rich zooxanthellar habitats because the waste products of coral heterotrophy go largely unutilized (e.g., Szmant- Froelich and Pilson, 1977; Muscatine and D’Elia, 1978); competition among zooxanthellae for nutrients may be minimal in this case. Competition should increase, however, as zooxanthellar biomass increases, and the zooxanthellar genotype that competes for nutrients best should ultimately prevail, regardless of its rate of growth in the absence of competition or its initial abundance (e.g., Huston and Smith, 1987; Tilman, 1988).
In M. annularis and M. faveolata living at 9 m depth at our study site, those efficient, specialized, but comparatively slowly growing zooxanthellae might be Symbiodinium C and B. In contrast, Symbiodinium A, E, and C2 seem to have played the role of early successional, rapidly proliferating opportunists in our experiments. In Experiment II, Symbiodinium A reached large numbers in only 8 weeks (Fig. 5, Recovery; see also Fig. 3, Low Light). Opportunistic behavior by Symbiodinium A has also been observed in M. annularis and M. faveolata during a natural coral bleaching event (Rowan et al., 1997) and in mixed in vitro cultures of Symbiodinium (Rowan, 1998; Carlos et al., 1999). Our observation of Symbiodinium E in Montastraea living in a marginal habitat near our study site (Río Cartí; see Toller et al., 2001) also suggests a weed-like ecology. Similarly, the unusual association of Symbiodinium C2 with M. annularis (Results) suggests that our experimental treatment enabled this zooxanthella to exploit a host species with which it does not commonly associate. We note, however, in the cases of both Symbiodinium A and E, it is unknown whether different observations involved one or several species of zooxanthella (see discussion on taxonomy, above).
Stability of zooxanthellar communities and coral bleaching
During natural episodes of coral bleaching (reviewed in Brown, 1997), even severely bleached colonies of M. annularis retain at least 10% of their pre-bleaching population of zooxanthellae (Porter et al., 1989; Fitt et al., 2000), which represents at least four times as many zooxanthellae as our low-light-treated corals had. Thus, the dramatic changes in species of Symbiodinium that we observed following low-light treatment of M. annularis and M. faveolata are unlikely to be common in nature. Mild coral bleaching or seasonal fluctuations in numbers of zooxanthellae in normal years (Stimson, 1997; Fagoonee et al., 1999; Fitt et al., 2000) involve lesser depletions of zooxanthellae, comparable to those observed in high-light-treated M. annularis in which communities of Symbiodinium did not change (Fig. 3, High Light, Colonies 6–10). Nevertheless, two high-light-treated corals that retained about 15% and 20% (Fig. 3, Colonies 4 and 5, respectively) of their zooxanthellae after treatment apparently did acquire detectable amounts of Symbiodinium E as a result of treatment. This suggests that severe natural bleaching episodes might modify communities of Symbiodinium in M. annularis, at least in part.
If infrequent, natural bleaching events at our study site (e.g., Lasker et al., 1984; Rowan et al., 1997) do allow Symbiodinium E or C2 to proliferate in M. annularis and M. faveolata from time to time, the effect must be transient or slight. These host-zooxanthella combinations are rarely encountered in unmanipulated corals from this reef (Rowan and Knowlton, 1995; Rowan et al., 1997; this study), despite occasional coral bleaching. On the other hand, at a nearby coastal site (Río Cartí), where stresses that can induce coral bleaching may be severe and frequent (Toller et al., 2001), disturbance appears to have a widespread effect on host-zooxanthella association. In high-irradiance habitats, M. faveolata and M. annularis at Río Cartí associate predominantly with Symbiodinium E—an opportunistic taxon of zooxanthella (above) that may also be stress-resistant (Toller et al., 2001). In this environment, these host-zooxanthella associations resemble the persistence of early-to-mid successional phototrophs under conditions of chronic disturbance (Odum, 1969; Horn, 1974; Huston and Smith, 1987).
Buddemeier and Fautin (1993) proposed that bleaching allows corals to replace their zooxanthellae with different, better ones (see also Baker, 1999, 2001; Kinzie et al., 2001). According to this "adaptive bleaching hypothesis," such replacements are driven by environmental change, which simultaneously makes some host-zooxanthella combinations less well adapted and other combinations better adapted than they had been. Baker (2001) found that the mortality of corals challenged with prolonged environmental change (transplantation) was reduced when they acquired new zooxanthellae, but these new host-zooxanthella associations were only established after coral bleaching. Similarly, our experiments indicate that severe bleaching allowed corals to associate with new zooxanthellae: Symbiodinium A, E, or C2 replaced Symbiodinium C or B in Montastraea (see above). However, in our experiments, environmental change was not a prerequisite; instead, Symbiodinium A, E, or C2 proliferated in the very environment that Symbiodinium C or B apparently thrive in (Figs. 3 and 5; Recovery). Although our experiments did not include prolonged environmental change and therefore was not a direct test of the adaptive bleaching hypothesis, they do show that testing this hypothesis may not be straightforward-severe coral bleaching may favor zooxanthellar replacements, irrespective of environmental change.
Disease-related disturbance of zooxanthellar symbioses
Symbiodinium A especially and also Symbiodinium E proliferated in the bleached tissues of corals with yellow-blotch disease (Fig. 7, Yellow). These zooxanthellae apparently gave many lesions of YBD their distinctive yellow color, and they were fugitives in the strict sense because their habitat was ephemeral. Our measurements suggest that their habitat lasted an average of about 5 months. Nonetheless, continuous progression of YBD across a coral would provide a large amount of habitat for these fugitive zooxanthellae to occupy.
Our findings explain the "yellow" in YBD, but they do not address the cause of the pathology. Most colonies we encountered were in two clusters that were surrounded widely by unaffected corals, which suggested an infectious agent with limited dispersal. YBD may have more than one etiology: our experimental results imply that it would arise when anything spread through a coral and disturbed stress-sensitive communities of zooxanthellae (Symbiodinium B or C) without actually killing the host immediately. One agent might even be a "rogue" (parasitic) genotype of Symbiodinium A that prospered at the expense of its coral host.
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