Methods for sampling free-living Symbiodinium (zooxanthellae) and their distribution and abundance at Lizard Island (Great Barrier Reef)

Journal of Experimental Marine Biology and Ecology 364 (2008) 48–53

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Journal of Experimental Marine Biology and Ecology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / j e m b e

Methods for sampling free-living Symbiodinium (zooxanthellae) and their distribution and abundance at Lizard Island (Great Barrier Reef) Raechel A. Littman a,b,⁎, Madeleine J.H. van Oppen a,b, Bette L. Willis a a b

ARC Centre of Excellence for Coral Reef Studies and School of Marine and Tropical Biology, James Cook University, Townsville, 4811, Queensland, Australia Australian Institute of Marine Science, PMB 3, Townsville MC, Townsville 4810, Queensland, Australia

A R T I C L E

I N F O

Article history: Received 17 March 2008 Received in revised form 29 June 2008 Accepted 30 June 2008 Keywords: Free-living Symbiodinium Symbiosis Zooxanthellae

A B S T R A C T The abundance and distribution of free-living dinoflagellates in the genus Symbiodinium have important implications for the ecology of coral reefs, determining both the symbionts available to newly recruited corals and symbiont types available for uptake by adult corals during environmental stress. However, little is known about where symbiotic dinoflagellates reside outside the host, due to the difficulty of capturing and detecting unicellular organisms in the marine environment. This study presents a successful protocol for sampling Symbiodinium from both the benthos and the water column. Comparisons of two detection methods for enumerating Symbiodinium indicated that conventional microscope analysis is accurate and more efficient when estimating Symbiodinium densities in sediment samples, while an automated particle counter (FlowCAM) was more efficient in detecting cells in the water column where densities are low. Symbiodinium densities were found to be relatively high (1000–4000 cells/mL) in sediment samples and much lower (up to 80 cells/mL) in the water column, indicating that the free-living form resides mainly in the benthos. Symbiodinium densities were found to be highly variable spatially, differing significantly between two reef locations. Within sites, elevated densities of Symbiodinium along reef margins combined with significant decreases in densities one meter away from the reef, suggest that cells aggregate within the reef habitat. © 2008 Elsevier B.V. All rights reserved.

1. Introduction The high productivity of coral reef ecosystems is largely attributed Symbiodinium, endosymbiotic dinoflagellates commonly referred to as zooxanthellae. These photosynthetic symbionts are important for vital nutrient cycling within their hosts (Hoegh-Guldberg, 1999), and a prolonged loss can lead to coral mortality (Brown, 1997; Hughes et al., 2003). Presently, several genera of dinoflagellates have been identified as endosymbionts of various marine invertebrates and protists, including Symbiodinium, Amphidinium, Aureodinium, Gymnodinium, Gyrodinium, Prorocentrum, Pyrocystis, Scrippsiella and Gloeodinium (Rowan, 1998; Wakefield et al., 2000). Symbiodinium is the most studied genus within this group (Baker, 2003). There are currently eleven named species of Symbiodinium and phylogenetic analyses of nuclear ribosomal DNA [including the small subunit (SSU) or 18S rDNA (Rowan and Powers, 1991a,b; McNally et al., 1994) and the large subunit (LSU) or 28S rDNA (Wilcox, 1998; Pochon et al., 2001)] and the chloroplast rDNA (23S) (Santos et al., 2002) have established that there are several lineages, designated as clades A through H.

⁎ Corresponding author. Department of Marine and Tropical Biology, James Cook University, Townsville, 4811, Queensland, Australia. Tel.: +61 07 4781 4801; fax: +61 07 4772 5852. E-mail address: [email protected] (R.A. Littman). 0022-0981/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.jembe.2008.06.034

Some zooxanthellae have been known to exist outside of the host and are commonly referred to as ‘free-living.’ Of the eight symbiotic dinoflagellate genera, Symbiodinium is the only group not known to have exclusively free-living species (Rowan, 1998). Genetic studies have recently found that several free-living strains belong to Symbiodinium, suggesting that some or perhaps all species can exist outside of a host organism. Through analysis of the LSU rDNA sequence data, two free-living species, Gymnodinium varians and G. simplex were reclassified within the Symbiodinium genus (Wilcox, 1998). Phylogenetic analysis of the SSU rDNA sequence revealed two other free-living strains belonging to Symbiodinium (Carlos et al., 1999; Gou et al., 2003). In addition, 5 free-living Symbiodinium-like isolates from the benthos were found to belong to clades A and B by analysing the ITS region of rDNA (Coffroth et al., 2006). Free-living zooxanthellae may serve as a potential reservoir of various genetic types from which corals can select. Hosts can initially acquire dinoflagellate symbionts in two ways: vertically in a “closed system” where coral eggs maternally inherit the symbionts or horizontally in an “open system” in which symbionts are obtained from the environment (Douglas, 1994). Approximately 85 percent of cnidarian host species acquire zooxanthellae from the environment (Harrison and Wallace, 1990; Richmond, 1997), indicating that sea water is a prevalent source of Symbiodinium. One study has verified that adult Briareum colonies can be infected by exogenous sources of zooxanthellae, suggesting that adult coral hosts can take up zooxanthellae

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externally and perhaps change their initial symbionts (Lewis and Coffroth, 2004). Asymbiotic octocoral recruits placed 20 m above the reef were also able to attain algal symbionts from the water column (Coffroth et al., 2006). Understanding the population biology and genetic diversity of the free-living phase of Symbiodinium requires the development of sampling protocols that account for variability in the distribution and abundance of Symbiodinium in the marine environment. Although free-living Symbiodinium cells have been isolated from the benthic environment, to date, no method has been devised to detect the genus in reef waters (Carlos et al., 1999; Gou et al., 2003; Coffroth et al., 2006). The existence of zooxanthellae in the water column has been demonstrated by using asymbiotic coral polyps as “symbiont samplers” (Coffroth et al., 2006). However, this did not allow for isolating the cells directly from the environment to determine their abundance. The purpose of this study was to find an effective sampling method that allows for future comprehensive analysis of the diversity of freeliving Symbiodinium in the reef environment. The methodology for sampling zooxanthellae must account for precision and efficiency of sampling and detection methods as well as spatial variability in abundance on small and large scales. Therefore, the objectives were: (a) to develop a sampling protocol for free-living zooxanthellae in the water column and sediments; (b) to compare the accuracy and efficiency of microscopy and the FlowCAM in detecting Symbiodinium; (c) to estimate the most effective sampling volume and size that will maximize sampling precision; (d) to determine if Symbiodinium abundance varies between separate reefs; and (e) to find the area of highest zooxanthellae abundance within the reef habitat. 2. Materials and methods 2.1. Study area and sampling design Lizard Island is located in the northern section of Australia's Great Barrier Reef (14°40′S 145°28′E). The Island is 7 square kilometers in size, and surrounded by three smaller islands (Palfrey, South and Bird), and a fringing reef that encompasses the 10 meter deep Blue Lagoon. The Lizard Island Group is considered a mid-shelf reef, positioned 30 km from the Australian shore and 19 km from the outer barrier reefs. Symbiodinium density was compared at 4 different locations on fringing reefs surrounding Lizard Island to determine the level of spatial variability among reefs. Two sites were located east of Lizard Is within the Blue Lagoon area, where reefs are subject to northwesterly winds during summer wind reversals and partially sheltered from the strong Southeast Trade Winds. The Lagoon reef extends approximately one kilometer and consists of shallow, patchy reefs to the South (ranging 0.5–2 m) and a somewhat deeper (1–3 m) section to the North. Western Lizard Is sites were more exposed to Southeasterly winds and may experience stronger currents during severe weather conditions. These sites tended to be slightly deeper (1.5–4 m) with dense coral cover. To account for probable differences in zooxanthellae advection and dispersal as well as reef effects, sediment was collected from random locations at two sites west (Horseshoe and Vicky's Reefs) and two sites within the Blue Lagoon. All samples were collected within a week span (November 9–15, 2006) during calm weather and high tide to minimize temporal effects that may have arisen from variation in oceanographic conditions. Variability in Symbiodinium abundance was also examined within a reef site due to the possibility of Symbiodinium aggregating near potential hosts (Carlos et al., 1999; Pasternak et al., 2006) and/or high micro-scale variability from reef effects (Martin et al., 2003). To determine if zooxanthellae abundance is greater within certain areas of the reef environment, sediment samples were collected in random locations at three regions of the Blue Lagoon Reef: the reef flat (0.5 m), bottom of reef slope (3 m) and 1 m from the reef's edge (3 m). Sampling was carried out on the same day to eliminate temporal variability.

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2.2. Sampling and processing Sediment sampling was performed on SCUBA and involved collecting 30 replicate cores per site. Coring was carried out by pressing a PVC tube of known volume (1.0 L water plus 500 mL sediment) into the sea floor, trapping the sediment and surface water. The tube was capped at the top to form a vacuum while pulling the container from the ground. The end was then rapidly closed to recover the sample (Summerhayes and Thorpe, 1996). Sediment samples were immediately washed, in turn, over filter trays with 102 µm and then 63 µm mesh size to remove larger particles. Replicate 10 mL aliquots were removed from the solution and preserved with formalin (10% final concentration) for abundance counts, or instantly frozen in liquid nitrogen for genetic analysis. Thirty replicate water samples were collected at random locations within the Lagoon Site directly above the boundary layer of the coral canopy. A 5 L Niskin Bottle sampler was used to entrap water at the desired distance from the coral. Samples were immediately filtered using a vacuum pump and millepore apparatus to concentrate the water constituents on 2 µm micro-filter paper. The concentrate was re-suspended in a known volume and fixed with formalin (10% final concentration). The sampling methods were further refined to maximize efficiency and accuracy. To establish the optimal volume of sediment that would minimize standard error, sediment samples of varying volume were collected in the same location within Blue Lagoon where zooxanthellae were known to occur. Sediment was targeted for this comparison because higher densities of Symbiodinium were detected in sediments than in water samples. Samples were collected and processed as described above and Symbiodinium cells were counted using a haemocytometer. Precision values were calculated by dividing the percent variation (standard error) by the mean Symbiodinum densities. The effective volume size was determined by comparing the precision of each sampling volume. Similarly, the most efficient sampling size was estimated by evaluating the precision for each number of replicates, indicated by a decrease in standard error. 2.3. Detection of Symbiodinium The conventional method for identifying plankton is microscopy; however, this is time intensive (Davis et al., 2004). Furthermore, zooxanthellae may be rare in the environment or patchy in distribution, necessitating high resolution detection methods (Fasham, 1978; Haury et al., 1978; Mackas et al., 1985). The flow cytometer and microscope (FlowCAM) measures light scatter and fluorescence of particles (Dubelaar and Jonker, 2000) while storing their digital images. This allows for counting, imaging and identification of plankton based on several parameters and has been found to produce similar results as microscopy in real-time (See et al., 2005; Sieracki et al., 1998). Therefore, the FlowCam may be a viable option for detecting Symbiodinium in water samples in low concentrations and in less time. As such, two detection methods were used to analyze sediment and water samples: haemocytometer counts using a light microscope and an imaging flow cytometer and microscope system (FlowCAM). Symbiodinium were identified in both methods by comparing cells to known Symbiodinium isolates from fresh coral tissue. The main characteristics used for identification were their coccoid form, cell plate structure, ~ 10 µm size and gold colour. Abundance of zooxanthellae was enumerated in suspended samples of known volume using a conventional high-powered microscope and haemocytometer (0.001 mL of concentrate). Four replicate 15 mL aliquots were concentrated by centrifugation at 10,000 ×g for one minute. The concentrated pellets were then resuspended to 0.5 mL in filter-sterilised seawater and placed on a haemocytometer with a Pasteur pipette. In addition, zooxanthellae were counted from replicate subsamples via a FlowCAM. In the flow cytometer and microscope system,

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samples are pumped into a flow cell, passing in front of an optical element that has been adjusted to focus on particles in its field of view. The video camera/framegrabber images the sample and counts particles as they travel past the optical element. The digitized images are then collected and stored by computer where they can be analyzed with FlowCAM software (Sieracki et al., 1998). A 20x objective lens was used to enable detection of the ~10 µm sized Symbiodinium. A 50 µm flow chamber was installed, which has a shallow width that matches the depth of focus of the lens. All samples were pre-filtered with a 30 µm nylon mesh to remove larger particles that could block the flow chamber. A drop of sodium dodecyl sulphate (SDS) was added to each sample to prevent cells from flocculating. Samples were constantly agitated to maintain suspension of particles and prevent clogging of the flow cell. Due to the low concentration of Symbiodinium, each sample was run for 10 min at 1 mL/min to increase the level of detection. A total of 0.02 mL of suspended sample passed through the 50 µm flow cell and was analyzed within the 10 min time period. A “library” file of Symbiodinium images was developed with the FlowCAM software by running a sample of cells removed from coral tissue and storing the images. After sample particles were analyzed by the FlowCAM, the particle images were filtered and sorted according to the parameters set by the zooxanthellae library. 2.4. Statistical analysis Variability in detection methods for water and sediment samples was compared using an orthogonal two-factor (“detection method” and “sample type”) ANOVA. The F-test for a Model 1 ANOVA was calculated since the levels of both factors were fixed. Due to low counts of Symbiodinium in water samples, many of the observed values contained zero counts. Furthermore, data tended to be positively skewed with standard deviations proportional to the mean concentrations. To account for heteroscedasticity in the data and low observed values, concentration values were log transformed using the formula from Zar (1999):

using oligonucleotide primers that amplify a wide range of dinoflagellates, including Symbiodinium and Gymnodinium (Lin et al., 2006). The PCR mixtures (50 ul) contained 10 pmol of the forward primer Dino18SF2 (5′-ATTAATAGGGATAGTTGGGGGC) and reverse primer 18ScomR1 (5′-CACCTACGGAAACCTTGTTACGAC), 2.50 mM each dNTP, 1X PCR buffer (Scientifix), and 0.5 U of Taq polymerase (Scientifix). PCR was performed with Applied Biosystems 2720 thermocycler and programmed with an initial 3 min step at 94 °C and thirty cycles consisting of 94 °C for 1 min, 53 °C for 1 min and 72 °C for 1.5 min and a final extension for 10 min at 72 °C. PCR products were visualized by agarose gel electrophoresis. PCR fragments were inserted into TOPO vectors and the ligations sent to the Australian Genome Research Facility for transformation and sequencing. A total of four clone libraries consisting of 96 clones each were analyzed using a BLAST search (http://www.ncbi.nlm.nih.gov/) to determine the proportion of Symbiodinium and Gymnodinium sequences present in the sediment samples. 3. Results 3.1. Detection of Symbiodinium The total mean density of Symbiodinium detected using both methods was more than 15-fold greater in sediment samples (810.01 ± 923.50 cells/mL) than in water column samples (47.11 ± 86.33 cells/mL). In microscopic analysis, cells were detected in only 14% of seawater samples, with an estimated mean of 14.48± 33.80 cells/mL, whereas the mean density detected by the FlowCAM was three times greater at 79.93 ± 108.58 cells/mL. Conversely, microscope counts for the sediment samples yielded approximately 1176.15 ± 946.66 cells/mL, whereas the FlowCAM estimate of mean density was less than half this at 443.878 ± 749.38 cells/ml (Fig. 1).

X0 ¼ logðX þ 1Þ: A nested ANOVA was carried out to test for differences in Symbiodinium density on eastern versus western Lizard Island reefs. Reef sites were randomly chosen and a hierarchical design was used to test for differences among the two “Island locations”, a fixed effect factor (Zar, 1999). Group density mean values tended to be proportional to the group variances. To normalize data distributions for Symbiodinium density, values were square root transformed using the formula from Zar (1999): X0 ¼

pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi X þ :05:

Spatial variability within reef was determined with a one-way ANOVA. Because Symbiodinium density was measured at three chosen locations (flat, margin and outside), the model was considered fixed and therefore interpreted as a Model 1 ANOVA. A Tukey multiple comparisons test was used to identify reef areas containing significantly different mean densities of Symbiodinium. 2.5. Genetic analysis Using light microscopy, Symbiodinium may be visually indistinguishable from its close relative, Gymnodinium, which may confound abundance estimates. A genetic analysis was therefore carried out on sediment collected from Blue Lagoon and Horseshoe Reefs to estimate the proportion of Symbiodinium versus Gymnodinium recovered from samples. Due to the low concentration of Symbiodinium cells in water samples, only sediment was used for genetic analysis. Genomic DNA was extracted as described in Wilson et al. (2002) from 0.5 ml of filtered sediment samples. The 18S rRNA gene was PCR-amplified

Fig. 1. Comparison of methods for detecting Symbiodinium densities. (a.) Concentration of Symbiodinium as measured by FlowCAM and Microscope. Vertical lines denote standard deviation of the mean, (b.) Precision of Symbiodinium density estimates as measured by microscope and FlowCAM.

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Analysis of variance confirmed that Symbiodinium densities differed significantly between sediment and water samples (F = 57.04, p b .001). However, a significant interaction between sample type and detection method (F = 0.337, p b .001) may have masked differences in counts between the microscope and FlowCAM. Furthermore, there was a large discrepancy in estimated densities between water and sediment samples using the microscope, whereas FlowCAM estimates remained relatively close for both water and sediment samples. The precision of each detection method was found to depend upon the type of sample analyzed (Fig. 1b). The microscope method was more precise than the FlowCAM for estimating Symbiodinium densities in sediment samples. In contrast, microscope counts were dramatically less precise than FlowCAM counts for water samples. For both sample types, however, the FlowCAM only reached a precision level between 0.25 and 0.3. The most precise method for detecting Symbiodinium in the ambient environment was through microscopic analysis of sediment, which estimated densities of Symbiodinium with twice the level of precision (precision level of 0.14) of the FlowCAM. The efficiency of the FlowCAM system was lower compared to the microscope for identifying and enumerating Symbiodinium. On average, four replicate counts of each sample (two 10 mL aliquots) took approximately 20 min with the microscope. In contrast, it took more than 30 minutes to analyze one sample with the FlowCAM. Each sample was run for 10 min to ensure detection of Symbiodinium and run twice to achieve a higher level of accuracy. Subsequently, data manipulation required 5 min for each sample and water was allowed to pass through the system for another 5 min between samples. Furthermore, frequent clogging by flocculating particles required many flow reversals and disassembling the apparatus to clean out the flow cell. This resulted in a much longer analysis process when compared to the microscope.

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Fig. 3. Comparison of mean Symbiodinium densities in sediment from eastern (Lagoon 1&2) and western (Horseshoe & Vicky's) reef sites. Vertical lines denote standard deviation of the mean.

3.2. Effective volume and sample size To find the optimal sample volume, the mean density of Symbiodinium was compared for a variety of sample volumes. Results indicated that mean density, as well as standard deviation around the mean, both decreased and stabilized when sample volume increased beyond 750 mL. Similarly, Symbiodinium density estimates were more precise and formed an asymptote at a sample volume of one liter (Fig. 2a). The optimal sample volume with regard to the best precision (0.05) was achieved at a sample size of 1750 mL. Comparison of Symbiodinium densities measured for sample sizes ranging from 10 to 80 samples indicated that mean density and standard deviation both remained relatively stable across this range of sample sizes. However, density estimates became more precise, reaching a level of 0.1 at a sample size of 30 and beyond (Fig. 2b). The optimal sample size with regard to the greatest level of precision (0.087) was attained at 70 samples. 3.3. Spatial variability between reefs The abundance of Symbiodinium differed significantly among reefs on Lizard Island (F = 28.39, p b .001). The mean density of Symbiodinium in the sediment was more than two-fold greater on western (2700.281 ± 2439.729 cells/mL) compared to eastern Lizard Island reefs (1073.709 ± 810.420 cells/mL). There was also a significant difference in Symbiodinium densities between sites within the island locations (F = 31.55, p b .001), indicating high variability between reefs. The greatest density of Symbiodinium was found within Horseshoe

Fig. 2. Determining optimal sample volumes and sample sizes for measuring Symbiodinium density (a.) Symbiodinium density estimates become more precise (standard error decreases) with increase in sediment sample volume, (b.) Symbiodinium density estimates become more precise with increase in sediment sample number.

Fig. 4. The density of Symbiodinium within the sediment at different locations of Blue Lagoon. A and B denote homogenous subsets as indicated by Tukey HSD.

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Table 1 Proportion of sequences that closely aligned (N97% identity) with Gymnodinium and Symbiodinium from four clone libraries constructed from sediment samples Clone Library Horseshoe Reef Blue Lagoon Total

1 2 3 4

Gymnodinium sp.

Symbiodinium sp.

0.06% 0.03% 0.03% 0.02% 0.03%

0.01% 0% 0% 0.03% 0.01%

Reef (4186.493 ± 2231.273 cells/ml) while the Lagoon Reefs and Vicky's Reef contained similar densities (Lagoon 1: 971.273 ± 646.781 cells/ml; Lagoon 2: 1176.145 ± 946.664 cells/ml; Vicky's Reef: 1214.069 ± 1600.390 cells/ml) (Fig. 3). 3.4. Spatial Variability within the Reef Environment The abundance of Symbiodinium varied among micro-environments of the Blue Lagoon reef ranging from the reef flat to one meter beyond the reef margin (Fig. 4; F = 3.573, p b 0.05). The sediment of the reef margin, approximately 3 m deep, contained the greatest density of Symbiodinium (2265.536 ± 1540.889 cells/ml). The reef flat, approximately 0.5 m deep or partially exposed, contained approximately twothirds the mean density of reef margin Symbiodinium (1356.944 ± 808.6705 cells/ml). The mean density of cells decreased to less than one-half the density of reef margin Symbiodinium at one meter from the reef margin (1033.036 ± 632.2204 cells/ml). However, mean cell densities were only significantly different between the reef margin and 1 m outside the reef (Tukey's HSD, p b 0.05). 3.5. Genetic analysis Using dinoflagellate-specific primers, over 100 eukaryotic genera were amplified from the sediment samples (electronic supplement). Both Gymnodinium and Symbiodinium were isolated from samples collected from Blue Lagoon and Horseshoe Reef (Table 1). Out of 319 sequences, derived from four clone libraries, ten sequences aligned with Gymnodinium sequences (0.03%) and four aligned with Symbiodinium sequences (0.01%). 4. Discussion This study shows that free-living Symbiodinium are mainly benthic, and that Symbiodinium cells occur in the water column, albeit it at low densities. Coffroth et al. (2006) proposed that zooxanthellae reside in the benthos and possibly migrate into the water column to seek a competent host. Our results are consistent with this hypothesis. High densities of cells were found at the base of corals while a few were detected within the water above the coral canopy, suggesting that cells were somehow suspended in the water column near potential hosts. However, it is possible that Symbiodinium sampled from the environment may be exclusively free-living types that are incapable of infecting corals. Coffroth et al. (2006) found that some isolates from the benthos were unable to establish symbiosis with corals. Therefore, cell aggregations near coral may not necessarily mean these dinoflagellates are “seeking” hosts. 4.1. Detecting Symbiodinium in the environment This study clearly demonstrates that Symbiodinium cells are detectable outside the host, and can sometimes be found in high densities. Further, the level of detection depends upon the sample type and the detection method. Microscopic analysis is precise when measuring sediment samples, with a standard error value only 15% of the mean density. Conversely, as a tool, the microscope was virtually incapable of detecting Symbiodinium in the water column as the

density of cells was too low to be found on a haemocytometer that only holds 0.001 mL of concentrated sample. As a higher through-put method, the FlowCAM was able to detect Symbiodinium in samples from the water column. The flow system allowed nearly 0.02 mL of suspended sample to be imaged within the 10 min observation period. However, regardless of its speed in analyzing samples, the FlowCAM was an imprecise and inefficient tool for both sample types, but particularly for sediments. Symbiodinium abundance estimated in sediment samples using the FlowCAM was dramatically lower than estimates from microscope counts. Under-estimation by the FlowCAM may have been caused by flocculating cells obstructing flow. Clogging was experienced when analyzing both sample types but particularly with sediment samples, which undoubtedly explained how density measurements became less precise. Particles could not pass through the cell and therefore, were unable to be counted. In addition, Symbiodinium occasionally attached to other cells, changing their measured parameters and causing them to be misidentified as larger particles. The efficiency of the FlowCAM was also negatively affected by the additional time necessary to clear the flow cell (see also Sieracki et al., 1998). As such, both accuracy and efficiency of this detection method could be dramatically improved by addressing the clogging issue. Although Sieracki et al. (1998) found that FlowCAM counts generally were more precise than microscope counts, underestimation of cell counts occurred when there was a rapid sink rate and cells could not reach the counting area of the flow chamber. In addition, large counting errors were attributed to cells clumping and being misinterpreted by the system. Observations obtained through both detection methods were compared to zooxanthellae cells freshly isolated from coral tissue to ensure correct identification of Symbiodinium. Nonetheless, identification errors may have existed in estimating Symbiodinium densities. Wilcox (1998) found that the use of cell structural morphology is limited in its capacity for classifying dinoflagellates since morphological plasticity and complex life cycles can obscure distinctions among groups. Using genetic analysis of the large ribosomal DNA sequence, Wilcox discovered that a dinoflagellate species from the Symbiodinium genus had been misclassified as Gymnodinium varians due to discrepancies in morphology. Therefore other dinoflagellates could have been mistaken for Symbiodinium. Genetic analysis of sediment samples confirmed that both Symbiodinium and Gymnodinium could be isolated from sediment samples in similar proportions (Table 1, electronic supplement). Since these dinoflagellates are indistinguishable by eye, estimates of Symbiodinium abundance may have been overestimated. 4.2. Accuracy of sampling methods In this study, estimated mean densities had large standard error values, indicating high levels of variability in the natural environment. This is consistent with most phytoplankton distributions which tend to have high spatial and temporal heterogeneity (Mehner et al., 2005). Nevertheless, a one liter sample yielded densities with a precision value of 0.15 and a sample size of 30 and above yielded a precision value of 0.1. Therefore, a one litre sample combined with a sample size of 30 captured a representative density of Symbiodinium with appropriate precision and is recommended for studies of the distribution and abundance of Symbiodinium. 4.3. Symbiodinium distribution patterns among and within reefs There is high spatial variability in Symbiodinium abundance between reef sites, as well as within reefs. In the free-living stage, it is assumed that Symbiodinium cells are subject to the same environmental conditions as other microalgae. As such, the distribution of zooxanthellae would be influenced by complex interactions between

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several physical and biological factors (Mouritsen and Richardson, 2003; Post et al., 2002). For instance, variation in the substrate and coral dimensions can cause differential rates in turbulence and mixing throughout the water column (Gross and Werner, 1994), which may have led to different degrees of Symbiodinium suspension and dispersal from the benthic environment. Phytoplankton retention is also possible within the reef environment through cell motility. Several studies have noted that Symbiodinium is mobile and responds to host chemical attractants, possibly enabling dinoflagellates to aggregate near potential hosts. (Fitt, 1984; Pasternak et al., 2006; Yacobovitch et al., 2004). Furthermore, disparities in host abundance, as a source of zooxanthellae may contribute to the differences in freeliving Symbiodium among and within reef sites. Jones and Yellowlees (1997) proposed that corals can continuously release zooxanthellae as a post-mitotic control for regulating density within coral tissue. This has been supported by Baghdasarian and Muscatine (2000) who have shown that a variety of healthy cnidarians expel dividing algal cells as a means to maintain a constant algal population density within the host. Therefore, corals may serve as a source of environmental Symbiodinium and may be the cause of elevated abundance within a reef location. 4.4. Conclusions and recommendations To confirm if Symbiodinium cells present in the environment are capable of forming an endosymbiosis, future studies must establish whether Symbiodinium found in the benthos can indeed colonize hosts. Secondly, benthic sampling can be used to genetically characterize free-living zooxanthellae strains to establish what types of Symbiodinium exist in the environment. This may further elucidate what potential symbionts are available in a particular environment for host uptake. Acknowledgements We thank R. Berkelmans of the Australian Institute of Marine Science for access to and aid in using the FlowCAM, J. Madams and M. McCormick for support in the field, and the Australian Research Council for funding. [SS] Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.jembe.2008.06.034. References Baghdasarian, G., Muscatine, L., 2000. Preferential expulsion of dividing algal cells as a mechanism for regulating algal-cnidarian symbiosis. Biol. Bull. 199 (3), 278–286. Baker, A.C., 2003. Flexibility and specificity in coral-algal symbiosis: Diversity, ecology and biogeography of Symbiodinium. Annu. Rev. Ecol. Syst. 34, 661–689. Brown, 1997. Coral bleaching: causes and consequences. Coral Reefs 16, 129–138. Carlos, 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 free-living strain. J. Phycol. 35, 1054–1062. Coffroth, M.A., Lewis, C.F., Santos, S.R., Weaver, J.L., 2006. Environmental populations of symbiotic dinoflagellates in the genus Symbiodinium can initiate symbioses with reef cnidarians. Curr. Biol. 16, 985–987. Davis, C.S., Hu, Q., Gallager, S.M., Tang, X., Ashjian, C.J., 2004. Real-time observation of taxa-specific plankton distributions: an optical sampling method. Mar. Ecol. Prog. Ser. 284, 77–96. Douglas, A.E., 1994. Symbiotic Interactions. Oxford University Press, Oxford. Dubelaar, G.B.R., Jonker, R.R., 2000. Flow cytometry as a tool for the study of phytoplankton. Sci. Mar. 64, 135–156. Fasham, M.J.R., 1978. The statistical and mathematical analysis of plankton patchiness. Oceanogr. Mar. Biol. Annu. Rev. 16, 43–79.

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