Characterization of Symbiotic Algae, Genus Symbiodinium, in Corals at St

CHARACTERIZATION OF SYMBIOTIC ALGAE, GENUS SYMBIODINIUM, IN CORALS AT

ST. LUCIE REEF, FLORIDA

Courtney Klepac

A Thesis Submitted to the Faculty of

The Charles E. Schmidt College of Science

in Partial Fulfillment of the Requirements for the Degree of

Master of Science

Florida Atlantic University

Boca Raton, Florida

August 2014

ACKNOWLEDGEMENTS

I am extremely grateful to Dr. Joshua Voss for his resolute dedication towards mentorship and entrusting me to be his first graduate student; members of the Voss Lab for their constructive criticism and assistance during crunch time; Dr.

Clay Cook for his continued involvement in developing scientific minds and willingness to edit all my thesis drafts; Dr. Randy Brooks; Jeff Beal for his invaluable expertise and field support at St. Lucie Reef; Dr. Misha Matz and Dr. Carly Kenkel for molecular and bioinformatics training and sequencing preparation materials; Scott

Hunicke-Smith at the University of Texas’s Genomic Sequencing and Analysis

Facility; Dr. Greg O’Corry-Crowe and Dr. Dennis Hanisak for lab equipment usage.

This research could not have been completed without funding support from the following: a Florida Fish and Wildlife Conservation (FWC) Commission State

Wildlife Grant (USFWS T-19-1); a grant from the Harbor Branch Oceanographic

Institute Foundation’s “Save Our Seas” Specialty License Plate Program; FWC for match contribution for boat time and personnel; Dr. and Mrs. Alex Marsh’s

Scholarship in Marine Science; a FAU President’s Challenge Scholarship, Graduate

Fellowship of Academic Excellence, and Graduate Grant; Research Assistantship support from the NOAA Cooperative Institute for Ocean Exploration, Research, and

Technology (CIOERT). A collection permit provided by the St. Lucie Inlet State Park also allowed coral sampling at St. Lucie Reef.

iii ABSTRACT

Author: Courtney Klepac

Title: Characterization of Symbiotic Algae, Genus Symbiodinium, in Corals at St. Lucie Reef, FL

Institution: Florida Atlantic University

Thesis Advisor: Dr. Joshua D. Voss

Degree: Master of Science

Year: 2014

The unique coral reef at St. Lucie Reef (Stuart, FL) persists despite environmental variability from extensive freshwater discharges, summer upwelling, and thermal instability. By examining the symbiotic zooxanthellae, or

Symbiodinium, that reside in corals, we can gain insight to coral physiology impacted by local stressors. Two scleractinian corals, Montastraea cavernosa and

Pseudodiploria clivosa were sampled over 1.5 years, including both wet and dry seasons. Zooxanthellae were isolated and quantitatively characterized using standard measurements and molecular techniques. Both coral species varied in zooxanthellae biomass, where Pseudodiploria clivosa had higher cell densities and chlorophyll concentrations than Montastraea cavernosa. Over time, these

iv parameters varied, but were not significantly altered by freshwater discharge events. Symbiodinium diversity and abundances were identified by ITS2 region amplification and next-generation sequencing. Novel associations between

Symbiodinium and each coral explained the observed physiological differences. The symbioses remained stable throughout and could indicate local adaptation for St.

Lucie Reef corals.

v CHARACTERIZATION OF SYMBIOTIC ALGAE, GENUS SYMBIODINIUM, IN CORALS AT

ST. LUCIE REEF, FL

LIST OF TABLES ...... viii

LIST OF FIGURES ...... ix

CHAPTER 1 ...... 1

BACKGROUND ...... 1

CHAPTER 2 ...... 6

INTRODUCTION ...... 6

Study site and coral collection ...... 9

Zooxanthellae density and chlorophyll analysis ...... 11

Statistical analyses ...... 12

RESULTS ...... 13

Zooxanthellae density ...... 13

Chlorophyll content ...... 14

Environmental variables ...... 16

DISCUSSION ...... 17

CHAPTER 3 ...... 23

INTRODUCTION ...... 23

METHODS ...... 28

Study site and coral collection ...... 28

DNA extraction ...... 28

Symbiodinium genotyping ...... 29

Statistical analyses ...... 31

RESULTS ...... 32

Symbiont genotyping ...... 32

DISCUSSION ...... 35

APPENDICES ...... 42

REFERENCES ...... 63

vii TABLES

TABLE 1 OTU sequence NCBI BLASTn matches ...... 58

viii FIGURES

FIGURE 1 Map of St. Lucie Estuary and Reef with study sites indicated by stars ...... 48

FIGURE 2 Density of zooxanthellae in millions of cells per square centimeter in

relation to site and sampling event season for P. clivosa (a) and M. cavernosa (b)

...... 49

FIGURE 3 Density of zooxanthellae in millions of cells per square centimeter in

relation to sampling event season ...... 50

FIGURE 4 Chl a (a and b) and c (c and d) densities among P. clivosa and M. cavernosa

over time ...... 51

FIGURE 5 Correlation of cellular chlorophyll a (pg cell-1) and zooxanthellae density

(cells cm-2)...... 52

FIGURE 6 Correlation of chlorophyll a scaled to coral tissue (μg cm-1) against

zooxanthellae density (cells cm-2) ...... 52

FIGURE 7 Cell volume (μm3) of zooxanthellae cells from P. clivosa and M. cavernosa

collected Summer 2013 ...... 53

FIGURE 8 Chlorophyll a content relative to zooxanthellae cell volume from Summer

2013 samples ...... 54

FIGURE 9 Discharge flow rates of canals in Lake Okeechobee and the St. Lucie Estuary

Watershed ...... 55

FIGURE 10 Environmental variables at St. Lucie Inlet ...... 56 ix

FIGURE 11 Proportion of Symbiodinium OTUs present in more than half of coral hosts ...... 57

FIGURE 12 Proportion of significant Symbiodinium OTUs in M. cavernosa by site across

five sampling events ...... 59

FIGURE 13 Proportion of significant Symbiodinium OTUs in P. clivosa by site across five

sampling events ...... 60

FIGURE 14 Principal coordinate analysis of Symbiodinium in M. cavernosa in relation to

season (a) and site (b) ...... 61

FIGURE 15 Principal coordinate analysis of Symbiodinium in P. clivosa in relation to

season (a) and site (b) ...... 61

FIGURE 16 Symbiodinium diversity in the Wider Caribbean ...... 62

x CHAPTER 1

BACKGROUND

In the past 3 decades coastal coral reefs in the Caribbean and southern

Florida have been increasingly affected by disease outbreaks, storm damage, freshwater runoff, and anthropogenic practices (Szmant 2002, Fabricius 2005,

D’Angelo & Wiedenmann 2014). Episodes of runoff are characterized by lowered salinity, high amounts of suspended solids, increased dissolved inorganic nutrients, and potential phytoplankton blooms, subjecting coastal coral reefs to osmotic and metabolic stress (Coles & Jokiel 1992, Mayfield & Gates 2007). Possible negative indirect effects include competitive inhibition by macroalgae and light limitation imposed by high densities of phytoplankton (D’Angelo & Wiedenmann 2014), and has been attributed to terrestrial inputs (Lapointe 1997, Lapointe et al. 2007).

However, many coral reef ecosystems along the Florida reef tract sustain a high diversity of commercially important species and support local economies despite this suite of stressors (Beal et al. 2012). Assessing potential physical and chemical stressors and the associated health of Florida’s coral reefs has been defined top priority in reef management strategies such as the Comprehensive Everglades

Restoration Plan (2012) and NOAA’s Coral Reef Conservation Program (2009).

The dominance of highly productive reef-building corals in benthic communities can be attributed to symbiotic associations between corals and

1 unicellular zooxanthellae from the genus Symbiodinium. Considered the most prolific phototrophic algae in tropical ecosystems (Trench 1993), photosynthate produced by Symbiodinium provides large quantities of organic carbon and energy to the coral host, thus enhancing calcification and contributing to reef structure accretion (Muscatine & Porter 1977, Trench 1993). Performance is dependent on light and thermal thresholds, and growth is regulated by nutrient availability (D’Elia and Cook 1988, Toller et al. 2001, Hoegh-Guldberg 1994, Rowan & Knowlton 1995,

LaJeunesse et al. 2010). Therefore, local environmental factors may impose beneficial or detrimental influences that ultimately dictate coral-Symbiodinium functioning.

In addition to Symbiodinium physiology, the identity and abundances of symbiont genotypes can affect overall coral growth and performance (Baker 2003,

Berkelmans & van Oppen 2006, Bellantuono & Baker 2008). Symbiodinium is a diverse genus, containing 9 sub-generic clades (A-I) (Pochon & Gates 2010), which also diverge into a multitude of genetic variants, or types. Distribution of variants is primarily correlated with temperature and light regimes (Warner et al. 1996,

Rowan et al. 1997, Baker 2003, Goulet et al. 2005, Berkelmans & van Oppen 2006,

Abrego et al. 2008, Sampayo et al. 2007). The physiology of diverse symbiotic associations contributes to differential growth rates in coral hosts (Muller-Parker &

D’Elia 1997) and allows corals to exist over a variety of depth, temperature, light, and latitudinal gradients (Rowan & Knowlton 1995, Rowan et al. 1997, Baker 2003,

Rodriguez-Lanetty et al. 2001, Sampayo et al. 2007). Furthermore, coral host life history and symbiont maintenance contributes to the local dominance and

2 prevalence of specific Symbiodinium clade types in some reef habitats (Buddemeier

& Fautin 1993, LaJeunesse 2005, LaJeunesse et al. 2010, Putnam et al. 2012).

However, fluctuating water conditions exert external stressors on the coral-algal association and have important implications for resulting coral reef health.

St. Lucie Reef (SLR; Stuart, Florida) is a shallow reef habitat within the St.

Lucie Inlet State Park. This reef lies 2.3 km south of St. Lucie Inlet and is subject to regulated freshwater discharge events from Lake Okeechobee and unregulated freshwater runoff from the broader St. Lucie Estuary watershed (Beal et al. 2012).

During the wet season (June to November), discharges and runoff can result in nutrient over-enrichment and harmful algal blooms in the St. Lucie Estuary, as well as turbid, low salinity water migrating over SLR (Graves et al. 2004, Lapointe et al.

2012). In addition, summer upwelling events can lower temperatures from an average of 23 - 31°C down to 17 °C (Beal et al. 2012). Light limitation, variable water temperatures and salinity synergistically reduce the growing season at reef habitats experiencing upwelling (Glynn 1977) in addition to terrestrial runoff.

Corals at SLR encrust the substrate, likely to maximize light attenuation and avoid displacement (Beal et al. 2012). Although subject to variable water conditions, SLR coral diversity is substantial in contrast to more northern reef habitats along the

Florida coast, which contain only a fourth of the 25 scleractinian species present at

SLR (Beal et al. 2012). This suggests that the corals at SLR are at the northernmost limit for many Caribbean corals along the Florida coastline (Reed 1982).

SLR is a unique reef based on coral presence at the edge of most coral species latitudinal limits and persistence under an array of environmental conditions, and

3 this reef has been relatively unstudied until recently. To better understand this coral community, detailed characterization is needed to determine what makes these species different than conspecifics further south and how such coral-algal holobionts are able to survive in such challenging conditions. One approach to describing the unique corals here is determination of algal symbiont assemblages and whether the association is different between coral host species, perhaps exhibiting a gradient extending southward from the inlet. The possible change in these associations over time is an important aspect of reef ecology. Are novel

Symbiodinium species found at SLR when compared to those detected in the

Caribbean? Do these associations change following disturbances imposed by natural or anthropogenic sources of environmental variation? Will there also be differences over time and along the reef tract? To establish baseline observations of the coral- algal symbiosis at SLR and assess the potential influences of environmental variability present, the research presented here examined possible variation in

Symbiodinium functioning, diversity, and abundance over time. If freshwater runoff during the wet season correlates to changes in these parameters, it can be inferred that external stressors modify the performance of the symbiotic association and may have implications for long-term functioning under stressful conditions.

Conversely, if the coral-Symbiodinium association remains relatively stable through time, then perhaps this relationship has been conditioned to tolerate the environmental conditions present.

To elucidate potential differences in Symbiodinium between the two dominant host coral species (Montastraea cavernosa and Pseudodiploria clivosa) at

4 SLR, cellular density (cells per unit area) and chlorophyll concentrations were assessed. To complement these measures, next-generation sequencing of the internal transcribed spacer region 2 (ITS2) in ribosomal DNA was used to detect

Symbiodinium genetic variants and determine if proportions change over time.

Recent advances in molecular ecology have allowed determination of symbiont genotypes, even at abundances < 1% of the entire assemblage. Detecting fine-scale changes in Symbiodinium dynamics could further our understanding of symbiosis ecology in the absence or presence of a variety of stressors. Any findings will contribute to assessing coral health under current watershed management of

Central Florida. Upcoming changes to this system via proposed improvements to the

St. Lucie Watershed set by the Comprehensive Everglades Restoration Plan (CERP) could affect the dynamics of coral-algal physiology and ecology. Therefore, gathering baseline information about the underlying association can provide diagnostic tools for effective management for this ecosystem and its valuable resources.

5 CHAPTER 2

Differential Fluctuations in Zooxanthellae Density and Chlorophyll

Content from Two Coral Hosts

INTRODUCTION

The health of coastal ecosystems is of growing concern worldwide as effects of climate change and anthropogenic activities are increasing (Mumby & Stenek

2008). Many economically and ecologically valuable coastal ecosystems are impacted by agricultural and residential runoff (Lapointe et al. 2004, 2012,

Fabricius 2005). Runoff components such as siltation, dissolved inorganic nutrients, and lowered water clarity aggregate during large-scale stormwater discharges concurrent with seasonality, providing fluctuating conditions that may not be beneficial for such prolific habitats (Fabricius 2005, Wooldridge & Done 2009,

Lapointe et al. 2012). Therefore, coastal coral reef communities are influenced by a suite of anthropogenic and natural environmental influences. Community structure on coral reefs can change as a result of chronic eutrophication, exemplified by a switch to increased macroalgal biomass (Lapointe et al. 2012, Szmant 2012,

Fabricius 2005, Mumby & Stenek 2008). In southeast Florida, coral diversity and depth distribution are known to generally decrease with persistent, increased turbidity (Kautz et al. 2007). However, these coastal reef tracts still foster diverse fish and benthic communities. Southeast Florida reefs substantially contribute to

6 local economies, yielding an annual value of approximately $5.7 billion in sales and income (Southeast Florida Coral Reef Initiative). Characterizing coral health on

Southeast Florida reefs and understanding the impacts of various stressors, including land-based sources such as runoff and freshwater discharge, is critical for conservation of these important ecosystems.

High productivity of coral ecosystems in shallow waters is primarily attributed to the symbiotic associations between corals, endosymbiotic bacteria, and single-celled dinoflagellate algae, or zooxanthellae. These algal symbionts are classified in the genus Symbiodinium (Muscatine & Porter 1997) and subsequently subdivided into nine clades (designated A through I; Pochon & Gates 2010). Of these nine, only four, clades A - D, are known to occur in scleractinian corals (Baker 2003).

Distributions of Symbiodinium variants are driven by host geography (LaJeunesse

2005, Finney et al. 2010, LaJeunesse et al. 2010, Davies et al. 2014, Thornhill et al.

2014) as well as temperature and light regimes (Iglesias-Prieto & Trench 1997,

Muller-Parker & D’Elia 1997, Rowan et al. 1997, LaJeunesse & Trench 2000,

LaJeunesse 2002, Ulstrup & van Oppen 2003, Finney et al. 2010). Physiological differences among zooxanthellae types promote coral host subsistence in a plethora of environmental conditions and habitats (Rowan & Knowlton 1995, Rowan et al.

1997, Baker 2001, LaJeunesse 2002, Rodriguez-Lanetty et al. 2003, Thornhill et al.

2008, Sampayo et al. 2007). Numerous studies have documented coral-algal responses to temperature variations from bleaching events and seasonality (Gates et al. 1992, Jones 1997b, Brown et al. 1999). Following episodes of thermal stress, corals demonstrate osmoregulatory stress, growth reduction, and symbiont

7 modifications (Gates et al. 1992, Fitt et al. 2000, Baker 2003, Cunning & Baker 2012,

Wooldridge 2013), which can lead to mortality if there is not sufficient recovery or acclimation (Buddemeier & Fautin 1993, Rowan & Knowlton 1995, Baker 2003,

LaJeunesse et al. 2007). However the mechanisms that allow corals to persist in subtropical habitats influenced by both coastal runoff stressors (fluctuating light, salinity, nutrients, and water clarity) and periods of relative environmental stability are less understood.

Zooxanthellae density and photosynthetic capability in corals serve as a proxy for measuring host health and performance, predominantly assessed after coral bleaching events (Jones 1997a, Lesser & Shick 1989, Fitt et al. 2000, Cunning &

Baker 2012). Loss of zooxanthellae also occurs under non-bleaching conditions as a regulatory mechanism when light and temperature regimes fluctuate (Jones 1997a

& 1997b, Titlyanov et al. 1996, Brown et al. 1999, Fagoonee 1999, Fitt et al. 2000,

D’Croz et al. 2001, Chen et al. 2005, Vargas-Ángel et al. 2007, Dubinsky & Stambler

2011, Tremblay et al. 2014). Seasonal patterns can significantly affect symbiont physiology, where density and chlorophyll concentration are negatively correlated with summer maxima (Brown et al. 1999, Fitt et al. 2000, but see also Chen et al.

2005, Cunning & Baker 2012). Chronic and dynamic photoinhibition of zooxanthellae (Brown et al. 1999) is concomitant with depleted coral energy reserves via increased carbon demand (Warner et al. 1999). Moreover, densities and pigment can increase when sea surface temperatures (SST) and photosynthetically active radiation (PAR) lowers during wet seasons (Fagoonee

1999, Fitt et al. 2000, Warner et al. 2002, Chen et al. 2005), presumably in response

8 to increased photosynthetic capacity. During the wet season, coastal coral reef environments may also be exposed to increased turbidity, nutrients, and light attenuation (Costa et al. 2004). Eutrophic and upwelled environments supplement nutrients and can result in reduced carbon translocation to the coral host under proliferating algal densities (Wooldridge 2013), thus reducing calcification rates

(Glynn 1977, McGuire & Szmant 1997, Fagoonee 1999, Tremblay et al. 2014).

Location of coral colonies also influences algal dynamics such that deeper corals or those situated under ledges have increased chlorophyll content but reduced densities (Falkowski and Dubinsky 1981, Dubinsky et al. 1984, Fitt et al. 2000). It is apparent that many factors influence zooxanthellae physiology, however the suite of influences imposed by freshwater runoff needs to be examined.

Coral-algal symbioses were examined over time in two dominant coral species, Montastraea cavernosa and Pseudodiploria clivosa, by monitoring two zooxanthellae parameters, cell numbers and chlorophyll concentrations. These zooxanthellae response variables were tested for potential effects of 1) host coral species, 2) site location along a distance gradient from the inlet, and 3) time

(including wet versus dry season). In addition, potential influences of disturbances, involving upwelling and discharge events, were considered and addressed through targeted sampling events.

METHODS

Study site and coral collection

Corals were collected at SLR (~2 – 8 m in depth) within St. Lucie Inlet State

Park from four sites of increasing distance from the St. Lucie Inlet, Stuart, FL (Figure 9 1). Individually mapped and monitored Montastraea cavernosa and Pseudodiploria clivosa colonies were sampled during the dry (June 2012, January and February

2013, June 2013) and wet (August, October, and November 2013) season. Sampling months correspond to the following seasons: January and February = Winter, June =

Spring, August = Summer, and October and November = Fall. Five to six host colonies of each species at each site were GPS marked to ensure subsequent collection (n= 33-37). However, following Hurricane Sandy (October 2012), two colonies were dislodged and not relocated. In addition, one M. cavernosa colony at the Ledge site could no longer be sampled after October 2012 due to partial mortality and small overall size. Coral fragments (2 – 3 cm2) were chiseled from the perimeter each host colony and transported to the surface in individual plastic bags.

Each coral sample was separated into two subsamples: Trizol for preservation of nucleic acids and 500 mL bags of seawater for zooxanthellae collection. The samples were stored on ice during transport to the laboratory in insulated coolers (<6 hours). Samples were stored at -20 °C (fragments in seawater) or -80 °C (Trizol subsamples) until further processing.

Temperature, pH, and conductivity were measured both at the surface and at depth of each site during sampling using a YSI Pro Plus (Yellow Springs, OH).

Additionally, salinity, temperature, and discharge rate were assessed from June

(Spring) 2012 to November (Fall) 2013 using the South Florida Water Management

District (SFWMD) DBHYDRO database, which collected daily in situ data from a canal station (Fig 1) within the watershed

(http://my.sfwmd.gov/dbhydroplsql/show_dbkey_info.main_menu). Weekly

10 salinity and temperature measurements from Florida Oceanographic Society (FOS) were provided by the citizen science program’s public data

(www.floridaocean.org/p/21/water-quality#.U484DF5jods).

Zooxanthellae density and chlorophyll analysis

Tissue was removed from the previously frozen, seawater-stored coral fragments with a recirculating Waterpik™ containing 35 psu Instant Ocean seawater®. The resulting homogenized slurry was concentrated twice through centrifugation. First, samples in 500 mL bottles were centrifuged at 5,031 g for 25 min, the seawater supernatant discarded. The pelleted material was resuspended and transferred to 15 mL centrifuge tubes and spun at 3,219 g for an additional 10 min. Remaining seawater supernatant was decanted and coral animal tissue was removed and frozen at -80°C for long-term storage. The zooxanthellae were resuspended in 35 psu Instant Ocean to yield a final 10mL cell suspension. For each sample, three individual 2mL aliquots of zooxanthellae suspensions were used for subsequent analysis. One aliquot was preserved in 10% formalin for zooxanthellae density. The second was used to determine chlorophyll a (chl a) and c concentration. The final aliquot was preserved in 20% DMSO-EDTA for later molecular phylotyping.

Formalin-preserved zooxanthellae placed on a haemocytometer were enumerated from scaled replicate photographs (n=5). A Canon G12 digital camera attached to a microscope was used to photograph cells within a 1 mm2 square of the haemocytometer under 40 x magnification, and cells were counted using the feature counter option within the digital program Coral Point Count in Excel (Nova

11 Southeastern Univ., FL). Surface area of coral fragments was determined using the foil method (Marsh 1970), where each fragment was analyzed in triplicate and mean surface area was used in subsequent analyses. Cell numbers from Summer 2013 were also normalized to zooxanthellae cell volume (μm3 = 1/6πd3), calculated by measuring diameters (d) of 50 haphazardly selected cells for each coral sample using CPCe. Mean cell volume among 50 cells per sample was used to estimate total cellular volume of zooxanthellae within each sample.

Chlorophyll samples were filtered through a 25mm GF/F filter with 35 psu

Instant Ocean seawater prior to freezing at -80° C overnight. Filters were homogenized in 90% acetone on ice using an OMNI GLH 7 mm x 95 mm Saw Tooth

Generator Probe, after which chlorophyll was extracted with acetone in the dark for

20 – 24 hrs at -20°C. Absorbances at 750 nm, 663 nm, and 630 nm were determined on a Spectronic 601 (Milton Roy, Wilmington, DE) and chlorophyll content was calculated using the Jeffrey & Humphrey equation (1975). Zooxanthellae density as well as chlorophyll a and c were standardized to coral fragment surface area.

Chlorophyll content from Summer 2013 samples were also standardized to zooxanthellae cell volume.

Statistical analyses

Statistical analyses were conducted with R 2.13.2 (R Developmental Core

Team 2013). Mean zooxanthellae densities, cellular chlorophyll, and chlorophyll concentrations were calculated for each host species at each site. A MANOVA collectively assessed the five response variables (zooxanthellae cell density, chl a per cell, chl c per cell, chl a per unit coral area, chl c per unit coral area) to account

12 for correlations among these responses against the factors species, site, and time

(season). Additionally, individual ANOVA tests were run with season, species, site, and the interaction of species*site, species*season, site*season, and species*site*season as factors for cell density and chlorophyll a and c concentrations. Response variables were tested against each other for correlations, and environmental variables such as temperature and salinity were also examined for correlations to density and/or chlorophyll. Mean zooxanthellae cell volumes and chlorophyll a and c content per cell volume from Summer 2013 were compared between M. cavernosa and P. clivosa using a Welch’s two-sample t-test (Table 1).

RESULTS

Zooxanthellae density

There were fewer zooxanthellae cells per cm2 in M. cavernosa than in P. clivosa (F1,165 = 63.93, p < 0.001) with densities in P. clivosa twice those observed in

M. cavernosa (Fig 2 and 3). On average, zooxanthellae densities in P. clivosa were 5.6

± 2.0 x 106 cells cm-2 while M. cavernosa had densities of 2.8 ± 1.1 x 106 cells cm-2.

Zooxanthellae densities did not vary significantly among sampling locations (F3,164 =

2.22, p = 0.09; Fig 2). Densities changed significantly over time (F4,163 = 9.65, p <

0.001) for both P. clivosa (F4, 96 = 5.81, p < 0.001) and M. cavernosa (F4,63 = 3.65, p <

0.05; Fig 2 and 3); and were highest in Summer 2013 (7.5 ± 3.9 x 106 cells cm-2, P. clivosa Central site) and lowest in Spring 2013 (1.6 ± 4.9 x 105 cells cm-2, M. cavernosa Central site). Tukey post hoc multiple comparison of sampling season showed that Spring 2013 had significantly lower densities than other sampling events (p < 0.001) except for Winter 2013 (p = 0.07). No two-way interaction 13 between species*site (F2,165 = 0.62, p = 0.54), species*season (F4,163 = 0.85, p = 0.49), and site*season (F12,155 = 0.30, p = 0.99) had an effect on zooxanthellae density, nor did the interactions among the three factors (F8,159 = 0.77, p = 0.63). Cell density was not correlated with salinities recorded in situ at the time of sampling (r = -0.12, p =

0.19) or temperature at the reef (r = 0.09, p = 0.34).

Chlorophyll content

Zooxanthellae in M. cavernosa had significantly higher cellular chlorophyll a and c content compared to P. clivosa (F1,166 = 34.20, p < 0.001 and F1,166 = 36.39, p <

0.001, respectively for chl a and c), where mean concentrations of chl a were 4.68 ±

-1 -1 1.25 pg cell and chl c were 1.85 ± 0.56 pg cell in M. cavernosa. In contrast, mean chl a and c for zooxanthellae in P. clivosa were 3.35 ± 0.87 pg cell-1 and 1.28 ± 0.33 pg cell-1, respectively (Fig 4a and c). Cellular chlorophyll content was negatively correlated to zooxanthellae density (chl a: r = -0.43, p = < 0.001; Chl c: r = -0.45, p <

0.001). Figure 5 displays separate correlations for chlorophyll a and cell density in

P. clivosa (Fig 5a) and M. cavernosa (Fig 5b). No significant site effects were observed among chlorophyll data (chl a: F 3,164 = 2.01, p = 0.11 and chl c: F3,164 =

2.12, p = 0.10). Chl a (F4,163 = 2.07, p = 0.09) and c (F4,163 = 1.60, p = 0.18) content per zooxanthellae cell did not vary by sampling event season. In addition, none of the tested interactions among factors had significant effects on chlorophyll levels. There was no correlation of cellular chl content with salinity (chl a: r = -0.04, p = 0.67 and chl c: r = -0.13, p = 0.19) or temperature (chl a: r = 0.04, p = 0.67 and chl c: r = -0.02, p = 0.86).

14 Expressing cellular chlorophyll content as chlorophyll content per coral tissue area provides a better representation of total pigment content from zooxanthellae in each coral species. Colonies of P. clivosa had higher amounts of both pigments than those of M. cavernosa (chl a: F1,166 = 15.81, p < 0.001 and chl c:

F1,166 = 12.84, p < 0.001). On average, zooxanthellae in P. clivosa contained 17.51 ±

6.07 x 106 μg cm-2 coral of chl a and 6.96 ± 2.31 x 106 cm-2 coral of chl c, while M. cavernosa zooxanthellae chl a and chl c was 12.73 ± 5.99 x 106 μg and 4.95 ± 2.45 x

106 μg cm-2 coral, respectively (Fig 4b and d). Chlorophyll content per unit coral surface area was positively correlated with zooxanthellae density (chl a: r = 0.77, p <

0.001; chl c: r = 0.77, p < 0.001). Figure 6 displays separate correlations for chlorophyll a per coral surface area and cell density in P. clivosa (Fig 6a) and M. cavernosa (Fig 6b). Chl a and c also changed over time (F4,163 = 10.32, p < 0.001 and

F4,163 = 9.47, p = < 0.001, respectively; Fig. 4b and d), where Spring 2013 had the lowest chlorophyll levels compared to all other sampling event seasons, except during Summer 2013 for chl a. Neither site nor interaction terms between all factors demonstrated significant effects. Chlorophyll content factored into coral tissue area was not correlated with salinity and temperature.

M. cavernosa zooxanthellae cells are larger than P. clivosa (Fig 7), where mean cell volumes were 752.17 ± 97.10 μm3 versus 422.81 ± 68.70 μm3. Total mean cell volume per coral tissue area in M. cavernosa was 2.07 x 105 ± 0.13 x 105 μm3 cm-2 and 2.29 x 105 ± 0.09 x 105 μm3 cm-2. Total zooxanthellae surface area did not differ between the two hosts (T = 1.40, p = 0.16), but did change over time (F4,163 =

10.54, p < 0.001). When chlorophyll content is normalized to zooxanthellae cell

15 volume, zooxanthellae cells from M. cavernosa contained less chlorophyll (chl a =

0.0066 ± 0.0028 pg μm-3 and chl c = 0.0026 ± 0.0011 pg μm-3) than those from P. clivosa (chl a = 0.0096 ± 0.0049 pg μm-3 and chl c = 0.0037 ± 0.0020 pg μm-3)(chl a:

T = -2.30, p < 0.05; and chl c: T = -2.20,p < 0.05). The difference in chlorophyll a per cell volume in both coral hosts is displayed in Figure 8.

Environmental variables

Salinity and temperature measured at the surface and at depth for the four reef sites did not differ among depth (T-test, p > 0.05), or by site (F3,3 = 0.50, p > 0.05 and F3,3 = 1.24, p > 0.05, respectively). Salinities at SLR ranged from 37.9-38.6 PSU, where the mean salinity was 38 PSU. The temperature average at SLR was 27°C and ranged from 24-28°C. During the freshwater discharge in Summer 2013, flow rate leaving Lake Okeechobee increased from 10 m3 s-1 on July 18, 2013 to 84 m3 s-1 by

July 25 (Fig 9), and lasted 4 weeks (DBHYDRO, SFWMD). Salinity, temperature, and depth during high tide in the St. Lucie Inlet at the time of coral sampling (August 8,

2013) were 19.5 PSU (high tide, 8.5 PSU low tide), 30°C, and 0.6 m (high tide), respectively (Fig 10 for salinity and temperature). However, salinity and temperature measured during high tide at depth at all four sites had a mean of 38

PSU and 27°C. A second planned discharge to further lower residual floodwater levels from July occurred on October 5, 2013 where flow increased to 43 m3 s-1 and lasted 2 weeks. Salinity, temperature and Secchi depth during high tide in the inlet were 33 PSU (high tide), 28°C, and 1.2 m, respectively, on October 18, 2013 where mean salinity and temperature at SLR was 28°C and 38 PSU.

16 DISCUSSION

The abundance and physiological functioning of symbiotic zooxanthellae are important for overall coral holobiont fitness. The potential effects on coral performance by changes in local water quality are substantiated by direct phototropic responses of zooxanthellae (Brown et al. 1999). The results of 5 sampling events spanning 1.5 years show variation in zooxanthellae density and photosynthetic pigment over time. Cell density was slightly higher in samples collected after discharge events (Summer and Fall 2013; Fig 9), which are generally characterized by increased surface water temperature, reduced salinity, increased dissolved inorganic nutrients, and lowered light availability (see also Costa et al.

2004). However, the zooxanthellae cell densities observed were not significantly higher than those recorded during non-discharge sampling in 2012. Previous long- term (Fagoonee 1999, Fitt et al. 2000) and short-term (Chen et al. 2005, Kemp et al.

2014) observations reported higher zooxanthellae densities in the winter and spring under reduced temperature and irradiance, which also applies at SLR (Fig 2 and 3). Costa et al. 2004 observed an increase in cell density during northeast

Brazil’s wet season (fall/winter), while chlorophyll increased during the dry season

(summer). In contrast to the observations presented here, zooxanthellae densities and chlorophyll in Brazilian corals were drastically reduced following heavy summer rains (Costa et al. 2001). The increases in photosynthetic pigments following extensive freshwater discharges in Summer and Fall 2013 could be a photoacclimatory response to reduced water clarity.

17 Turbid freshwater runoff travelling over SLR alters light attenuation, thereby reducing down-welling light to resident coral colonies (Beal et al. 2012).

Zooxanthellae are known to acclimatize to reduced light availability by either increasing their chlorophyll content or by changing the size of photosynthetic pigment molecules (Falkowski & Dubinsky 1981, Iglesias-Prieto & Trench 1994).

Increases in chlorophyll could be a compensatory mechanism to changes in water clarity following discharge events. It is also possible that the positive correlation between zooxanthellae density and chl content per unit coral skeletal area explains the higher amount of chl a and c measured; or self-shading from increased densities promoted an acclimatory increase in pigment. During future samplings, Secchi depth and PAR readings should be incorporated to provide further insight on correlations between water clarity and zooxanthellae responses.

There are several differences observed in zooxanthellae density and chlorophyll concentration between M. cavernosa and P. clivosa. Many studies report a range of average cell densities of 1 to 4 million cells cm-2 (Brown et al. 1999,

Fagoonee 1999, Fitt et al. 2000, Chen et al. 2005), and zooxanthellae in the coral M. cavernosa fall within this range. P. clivosa however, contains significantly more zooxanthellae cells year-round. M. cavernosa has larger corallites and thicker tissue associated with heterotrophy (Porter 1976, Lesser et al. 2000), which could reduce reliance on autotrophy by zooxanthellae (Houlbréque & Ferrier-Pagés 2009) or provide less interstitial space for zooxanthellae cells. It is hypothesized an influx of phytoplankton concurrent with freshwater discharge could support polyp feeding

(Beal et al. 2012), thereby providing more nutrients to associated zooxanthellae

18 (D’Elia & Cook 1988). Chlorophyll content per cell was higher in zooxanthellae from

M. cavernosa than P. clivosa. Examining scaled images of zooxanthellae cells in both coral hosts from Summer 2013, cells in M. cavernosa are 40% larger than those in P. clivosa (Fig 7). Wilkerson et al. 1988 also observed larger zooxanthellae cells from

M. cavernosa in comparison to other coral species. Larger cells could provide more space for either more or larger photosynthetic units (Iglesias-Prieto & Trench

1994), and the potential explanation is supported by the chlorophyll per volume results. When chlorophyll content is normalized to coral tissue area, P. clivosa contains more pigments molecules than M. cavernosa, a net result of having higher zooxanthellae cell densities albeit comparable total cellular volume. Larger cells require more host cellular space, which could in part explain the lower cell densities within M. cavernosa. In addition, functional differences in holobiont performance may be a result of variations in symbiont assemblages.

Interestingly, neither zooxanthellae cell density nor chlorophyll content not differed for both coral species among the four sites at SLR. These sites are of increasing depth and distance from the St. Lucie Inlet. The most southerly site supports a higher diversity of reef species (Beal et al. 2012), suggestive of mitigated or reduced impacts from freshwater discharge. The North site is approximately 2.3 km from the inlet, so effects of environmental fluctuations synergistic with coastal runoff were expected to elicit a response in zooxanthellae response variables. In addition, this site is only 2 m deep whereas southern sites are 5 – 7 m, where impacts from sedimentation and tidal surge could be alleviated (Beal et al. 2012).

Despite high turbidity and surge at the north site, yearly mean light availability for

19 corals is similar to the deeper southern sites (2012 North PAR = 1000 mol m-2 sec-1,

South PAR = 1000 mol m-2 sec-1, Beal unpub. data). Lowered cell density with increases in pigment at depth has been reported previously (Dubinsky et al. 1984,

Muscatine et al. 1989), but similar measurements have not been made at SLR.

Therefore, SLR corals and their associated zooxanthellae may be adapting differently to reduced light regimes via cellular density increases, as seen in P. clivosa, or by containing more cellular chlorophyll, as seen in M. cavernosa.

Increases in land use and anthropogenic practices combined with continued freshwater discharges could exacerbate local water quality conditions and introduce potential stressors to coastal corals (Kautz et al. 2007). Freshwater intrusion is associated with increased sediment and nutrient loading, lowered light availability, phytoplankton production, and reduced salinity (Szmant 2002, Fabricius 2005,

Lapointe et al. 2012). Corals and symbiotic zooxanthellae are directly and indirectly affected by physical environmental factors occurring at SLR. Excess sedimentation resulting from runoff can trigger increased mucus production in M. cavernosa

(Vargas-Ángel et al. 2007), interfering with feeding, and can also negatively affect calcification (Carricart-Ganivet & Merino 2001, Cook et al. 2002). This is one potential explanation for the absence of M. cavernosa colonies at the North site.

Direct effects of freshwater runoff can also be linked to modifications in coral calcification (Fabricius 2005). As a result of lowered light availability as well as wave surge, coral colonies at SLR exhibit encrusting growth forms and have reduced, fragile skeletons (Beal et al. 2012). Lowered calcification rates from reduced carbon translocation (McGuire & Szmant 1997) to the coral host occur

20 despite increases in zooxanthellae densities after discharge events. Zooxanthellae that regularly experience lowered irradiances may be shade-adapted, able to photoacclimatize by increasing chl content (Falkowski & Dubinsky 1981, Tremblay et al. 2014). In addition to photoacclimation, Tremblay et al. (2014) showed increased heterotrophy with reduced carbon translocated under low irradiances.

Together, these mechanisms sustain calcification rates and may explain how the corals at SLR are able to grow in otherwise unfavorable conditions.

The variations in zooxanthellae density content observed over time in both

M. cavernosa and P. clivosa has important implications for the complex underpinnings of zooxanthellae responses to continued environmental fluctuations.

Since algae at high densities are nutrient limited and carbon translocation to the coral host is reduced (Muscatine et al. 1989, Fitt et al. 1993), SLR corals containing higher than normal zooxanthellae densities concurrent with fragile skeletons could be an impairment if discharges continue. In addition, it has been demonstrated that corals with higher zooxanthellae densities augmented by terrestrial runoff are more susceptible to thermal stress (Cunning & Baker 2012). Under stress, a disruption of autotrophic processes affects growth, reproduction, and tissue biomass of the coral host (Fitt et al. 2000, Szmant 2002, Fabricius 2005).

The results of this study reveal dynamic changes in zooxanthellae density and photosynthetic pigment under environmental variability at Florida’s most northern reef for many Caribbean coral species. Since zooxanthellae within

Montastraea cavernosa and Pseudodiploria clivosa respond differently over time, effective management shouldn’t focus on a particular coral species as an indicator

21 but include the entire reef system for holistic strategies. Moreover, zooxanthellae responses and environmental variables were similar at all sites, suggesting that any potential effects from discharge events could extend the entire reef tract. The differential responses observed in cell number and chlorophyll concentration may represent tradeoffs between the two host coral species and can indicate diversity in coral persistence from challenging environments. It is apparent that corals here are surviving under intense conditions and could reveal future mechanisms of symbiont performance at coastal reefs if anthropogenic practices and freshwater runoff increase in Southeastern Florida.

22 CHAPTER 3

Symbiodinium Sequencing of ITS2 Region Reveals Stable Symbiont

Assemblages and Novel Symbiotic Associations

INTRODUCTION

Recognized for thriving in tropical, oligotrophic environments, many coral reef habitats are increasingly exposed to thermally variable and eutrophic conditions, especially near populated and/or upwelled coasts (Coles 1975, Fabricius

2005, Wooldridge & Done 2009, Szmant 2002, Lapointe et al. 2012). Some of these reefs are able to persist despite the presence of abiotic stressors, and elucidating the mechanisms for coral resilience has become a priority of coral reef ecology

(International Society for Reef Studies 2004, NOAA Coral Reef Conservation

Program 2009). The differential responses of reef-building corals to environmental change have primarily been attributed to the underlying ecology and physiology of coral hosts and their symbiotic dinoflagellates from the genus Symbiodinium (Baker

2003, Berkelmans & van Oppen 2006, Hoegh-Guldberg et al. 2007). This mutualistic association contributes to the ecological dominance of coral species in reef ecosystems over evolutionary time scales (LaJeunesse 2010, Finney et al. 2010).

However, past environmental changes have influenced the functional response of coral-algal symbioses, and resilience through time may be affected by endosymbiotic community structure.

23 Over the past two decades, much research has focused on determining the diversity and community structure of these algal symbionts that reside within coral hosts (Trench 1993, LaJeunesse 2002, 2004, 2010, Finney et al. 2010, Thornhill et al.

2006, Savage et al. 2002). Taxonomic understanding of symbiotic zooxanthellae is limited; to date, all types have been grouped into a single genus, Symbiodinium. The current extent of elucidating Symbiodinium diversity has primarily been achieved using molecular genotyping with the ribosomal internal transcribed spacer region 2

(ITS2; Litaker et al. 2007, Sampayo et al. 2009, Stat et al. 2011). This work has revealed high diversity within the genus (Rowan & Knowlton 1995, Baker and

Rowan 1997, LaJeunesse & Trench 2000, LaJeunesse 2001, Pochon et al. 2012, van

Oppen et al. 2001), and variations in intragenomic sequences indicate distinct lineages (LaJeunesse 2002, Thornhill et al. 2007, Sampayo et al. 2009), even within the same sample (LaJeunesse 2002, Thornhill et al. 2006). Currently, nine sub- generic clades (identified as A through I) (Pochon & Gates 2010) are recognized, and types (or sub-clades) have been identified using molecular marker sequence differences and DGGE profiles (Litaker et al. 2007, Thornhill et al. 2007, Pochon et al. 2012, Bellantuono & Baker 2008, Sampayo et al. 2009, LaJeunesse 2002, 2003,

2004a & b, 2005).

Prevalence of certain Symbiodinium types within a reef locale are believed to be primarily determined by biogeography and environmental conditions that affect photophysiology, most notably light and temperature (Rowan et al. 1997, Baker

2003, Warner et al. 1996, Berkelmans & van Oppen 2006, Sampayo et al. 2007,

Thornhill et al. 2008). It is likely that each type has particular environmental

24 thresholds (LaJeunesse & Trench 2000, LaJeunesse 2002), and genetic differentiation can arise under niche integration. One reef locale or region can contain an ecologically dominant type or an assortment of coexisting variants

(LaJeunesse 2002, LaJeunesse et al. 2003, Baker 2003, Goulet 2006). The particular coral-Symbiodinium combination can influence how the symbiotic entity, or holobiont, functions as an ecological unit and responds to environmental changes

(Berkelmans & van Oppen 2006).

Maintenance of Symbiodinium community assemblages under fluctuating environments has important implications for the persistence of reef-building corals.

The symbiosis can persist, breakdown (as observed during coral bleaching), undergo advantageous modifications, or evolve adaptive traits. Unless resident

Symbiodinium are broadly tolerant to the local environment, it is critical that acclimatization occurs within monthly time scales for a coral-algal holobiont to withstand changing environmental conditions (Hoegh-Guldberg et al. 2007,

McGinley et al. 2012, Putnam et al. 2012). Generally, stable, less diverse

Symbiodinium assemblages are maintained in the absence of environmental variability and in brooding corals (LaJeunesse 2002, 2004, 2005, Thornhill et al.

2006, Bellantuono et al. 2011). In broadcast-spawning coral populations that have been studied, modifications made during bleaching events are gradually reverted back to the dominant Symbiodinium assemblage (Toller et al. 2001, Thornhill et al.

2005, LaJeunesse 2005). In a few of these studies, the opportunistic stress-tolerant

Clade D1a (Symbiodinium trenchi) appeared during bleaching and remained in coral hosts several months afterwards. Baker (2003) defined symbiont switching or

25 shuffling as a means to acquire symbionts physiologically tolerant to thermal stress, which could be an adaptive mechanism to overcome or recover during bleaching events (Baker et al. 2004). The occurrence and strength of disturbance promotes symbiont turnover that persists and confers increased host fitness (Baker et al.

2004, Berkelmans & van Oppen 2006, Jones et al. 2008, Silverstein et al. 2012). This mode of flexibility is challenged (Mieog 2009, Mieog et al. 2009, Putnam et al. 2012) as a short-term benefit with ecological consequences, such as reduced growth or increased competition among resident symbionts. If coral hosts can indeed remain stable in their associations under environmental fluctuations, associating with a single dominant clade may be a preconditioned trait (Thornhill et al. 2005,

Bellantuono et al. 2011, McGinley et al. 2012, Kenkel et al. 2013) or physiological flexibility of the coral-algal holobiont (Bellantuono et al. 2011).

The contention in delineating whether a coral species hosts a single

Symbiodinium genotype or several could be a bias of the molecular technique employed. Users of methodologies such as denaturing gradient gel electrophoresis

(DGGE) have often reported that coral hosts may harbor only one or a few dominant types (LaJeunesse et al. 2002, 2003, 2004, 2005, Thornhill et al. 2005). Thornhill et al. (2006) estimated that potential background types constituting <10% of the symbiont composition may not be detected using these traditional methods. Recent high-resolution techniques including real time PCR and next-generation sequencing have a much higher sensitivity to detect target genes, entire genomes, or transcriptomes (Mieog et al. 2007 & 2009, Putnam et al. 2012, Bellantuono et al.

2011, Quigley et al. 2014). A recent study using such approaches (Silverstein et al.

26 2012) has indicated that most, if not all, coral species have the ability to associate with many clades, even in species previously determined to be specific to only one type of Symbiodinium. Examining 39 reef coral species, of which 26 were deemed specialists, all were found to associate with more than one type (Silverstein et al.

2012). It was concluded that a gradient of specificity exists over which dominant and background strains are associated, and this can vary in frequencies within the same coral population. It is clear that high-resolution techniques such as next- generation sequencing can detect extremely low abundances of any Symbiodinium type associated (Quigley et al. 2014). Discerning the incidence of assemblage modifications or stability is critical for examining coral-algal ecology and evolutionary persistence under environmental fluctuations.

If variation in environmental conditions, such as water clarity and salinity levels from seasonality and freshwater runoff, prompt changes in symbiotic associations, this should be evident in coral species at St. Lucie Reef (SLR).

Alternatively, stability in Symbiodinium assemblages throughout seasonal fluctuations could imply adaptation by the resident symbiont types to local abiotic stressors. Two dominant coral species at SLR, Montastraea cavernosa and

Pseudodiploria clivosa, exhibit distinctive morphologies as well as differences in

Symbiodinium abundance and chlorophyll content. As broadcast spawners, these two coral species acquire symbionts from the environment (cf. Szmant 1986) and the Florida Current may connect larval dispersal to south Florida conspecifics. Many

Caribbean coral species at SLR are at their northern limits along coastal Florida and the genetic identities of Symbiodinium populations are unknown. This study

27 assesses the diversity and abundance of Symbiodinium spp. over time, between two coral species, and among sites by amplifying and sequencing the ITS2 region of ribosomal DNA.

METHODS

Study site and coral collection

The samples used in this study were those collected for the zooxanthellae studies in Chapter 2. There were 100 samples from P. clivosa and only 67 samples from M. cavernosa since this species is absent at the North site. Table 2 displays which colonies from each site were sampled at each collection date; the final aliquot was preserved in 20% DMSO-EDTA buffer for DNA extraction and sequencing and stored at -80° C.

DNA extraction

Zooxanthellae DNA was extracted from M. cavernosa and P. clivosa using modified Promega Wizard Genomic DNA Extraction Protocol (Madison, WI;

LaJeunesse et al. 2010). An aliquot of 20-40 mg isolated zooxanthellae cells was placed in a 1.5 mL microcentrifuge tube and washed with 10X Tris-EDTA buffer. For cell lysation, 500 µL of 0.5mm glass beads and 600 µL Nuclei Lysis Buffer (Promega) were added and bead beaten for 140 s at 6.5 m/s in a FastPrep®-24 (MoBio, Santa

Anna, CA). Lysates were incubated with 0.1 mg/mL proteinase K and mixed at 1200 rpm every 15 min at 65°C for a total of 90 min. After cooling, 250 µL of Protein

Precipitation Buffer (Promega) was added and samples were chilled on ice for 20 min. Following centrifugation at 12,000 rpm for 5 min, 400 µL of supernatant was removed and transferred to a new 1.5 mL microcentrifuge tube containing 700 µL 28 chilled 100% isopropanol and 25 µL of sodium acetate (3M, pH 5.6). Precipitated

DNA extracts were stored overnight at -20°C and afterwards centrifuged to pellet nucleic acids. The isopropanol-sodium acetate supernatant was removed and the pellet was washed with chilled 70% ethanol twice where it was then dried and rehydrated in 80 µL of Resuspension Solution (Promega) at 65°C for 60 min. DNA extracts were stored at -80°C until PCR.

Symbiodinium genotyping

To elucidate Symbiodinium diversity for each of the 167 samples, the ITS2 region was amplified via touch-down PCR (Don et al. 1991) using the forward primer its-dino (GTGAATTGCAGAACTCCGTG) and the reverse primer its2rev2

(CCTCCGCTTACTTATATGCTT; Pochon et al. 2001). Each 30 µl reaction consisted of

2 µl (=20 ng) of template DNA, 3 µl of ExTaq HS 10x ExTaq Buffer (Takara

Biotechnology), 0.7 µl of 10mM dNTPs, 1µl of an ITS2 10 µM forward and reverse primer mix, 0.15 µl ExTaq HS Polymerase (Takara Biotechnology), 0.15 µl Pfu polymerase (Agilent Technologies), and milli-Q water was added to achieve final reaction volume. “Cycle-check” PCR amplifications were ran on a Tetrad 2 Peltier

Thermal Cycler (Bio-Rad, Hercules, CA) using reaction parameters successfully employed by LaJeunesse and Trench (2000). An initial melting temperature of 94°C was set for 5 min, followed by 19 cycles of 94°C for 40 sec, 59°C for 2 min, and 72°C for 1 min, then a final extension of 72°C for 10 min. Any sample that did not represent equal band intensity on a 1% agarose gel following 19 cycles had 1 – 12 cycles added until a faint band was visible on the gel. This cycle-checking approach

29 avoids over/underrepresentation of product amplicons (Kenkel et al. 2013). Total qPCR cycle numbers for each sample are reported in Appendix A.

Each PCR product was cleaned using a PCR clean-up kit (Fermentas Life

Sciences), quantified, and diluted to 10 ng/µL. These templates were subject to a second PCR reaction to ligate unique dual barcoded primers and adaptors to each sample. Since the identity of each sample is preserved through the barcode sequence, samples can be pooled for sequencing. The adaptor designs were as follows: Rapid primer + unique barcode + its-dino and its2rev2 primer, respectively

(TCGTCGGCAGCGTC+AGATGTGTATAAGAGACAG+GTGAATTGCAGAACTCCGTG,

GTCTCGTGGGCTCGG+AGATGTGTATAAGAGACAG+CCTCCGCTTACTTATATGCTT, see Green et al. 2014, Quigley et al. 2014). The reaction profile was identical to ITS2 amplification but only four cycles were needed to incorporate barcode primers.

Once all samples are verified on a 1% agarose gel, relative band intensity was visually inspected as an indicator to pool PCR products into a final sample. For example, if the brightest bands indicate 5 µL of the sample should be pooled and if a lane is half the brightness, then 10 µL of that sample should be added to the final pooled sample. These final pooled samples were cleaned using the PCR clean-up kit

(Fermentas Life Sciences). Samples were eluted to 20 µL and run on a 1 % agarose gel stained with SYBR green to achieve a single product band at ~500 bp, ensuring that all pooled PCR products were of appropriate length. Bands were excised and products were extracted using a commercial gel extraction kit. The resulting samples were submitted for Illumina MiSeq sequencing at the University of Texas

30 (Austin, TX) sequencing facility. Barcode assignment of pooled samples were preformed, and based on the barcode results samples were diluted to 500 pM, followed by prepatory qPCR prior to loading the MiSeq.

Following trimming and quality filtering of individual reads to remove adaptors, barcodes, and primers 2,757,308 reads remained. The program CD-HIT-

OTU (Weizhong 2011, UCSD, CA) was used to cluster concatenated reads from all samples into identical groups at 97% identity to identify true operational taxonomic units (OTUs). An OTU is a commonly used term in molecular biology to delineate between taxonomic groups, superseding phylotype designation. After mapping sequence reads from all samples to each OTU and calculating the total number of mapped reads per sample, the proportion of reads mapped to each cluster relative to the total reads mapped per sample were calculated. The 19 OTU sequences were then aligned in the publicly available Cyberinfrastructure for Phylogenetic Research

(CIPRES) gateway using two multiple sequence alignment programs for nucleotide sequences, Multiple Alignment using Fast Fourier Transform (MAFFT; Katoh et al.

2002) and ClustalX (Larkin et al. 2007); these sequences were then blast against the

GenBenk (NCBI) nucleotide reference collection.

Statistical analyses

All analyses were performed using R v 3.0.2 (R Development Core Team

2013). Samples and OTUs containing low read counts (less than 0.1% of all samples) were removed from the dataset. The difference between representation of raw OTU counts within each species and among site and season were estimated using the

MCMC.OTU package (Matz 2014, UT, TX). Pairwise comparisons were carried out

31 using Poisson-lognormal generalized linear mixed models using fixed effects of species, site, season, species*site, species*season, and site*season. Downstream analysis adjusted the model to retain those OTUs that were modeled reliably, delimited by autocorrelations and meeting confidence parameter estimates.

A principal coordinate analysis using the library ‘vegan’ (Oksanen et. al 2013) was also used to assess Symbiodinium relative abundance dissimilarities relative to season, species, and site. Low-count (under-sequenced) samples were removed from the original dataset if these samples and/or OTUs represented less than 10% of all samples. Bray-Curtis similarities were computed for all log-transformed samples, and the PCoA derived relationships were displayed as multi-dimensional scaling plots.

Multiple stepwise, paired correlations determined the relationship among the dominant OTUs from each coral host against each other. A positive relationship indicates an interaction between two OTUs, and negative correlations represent dominance or replacement of one OTU read abundance over another.

RESULTS

Symbiont genotyping

Of the 167 samples used for ITS2 sequencing, 2,757,308 sequence reads across all samples were produced for clustering in the CD-HIT-OTU pipeline. Cluster analysis yielded 945,730 (72%) unique sequences that were mapped to 19 reference OTUs. Of the 19 OTUs, average base pair length was 321 and ranged from

304 (OTU 15) to 361 bases (OTU 1). Mapped number of sequences per sample to each OTU ranged from 0 to 17787 sequences (mean: 5663). The most frequent OTU 32 (OTU 1) represented 69% of all mapped reads, where the least assigned OTU (OTU

16) represented only 1.06 x 10-6 % of all sample reads. Within M. cavernosa and P. clivosa, 6 of the 19 OTUs (M. cav: OTU 1, 2, 4, 6, 9, 11 and P. cli: OTU 1, 2, 3, 5, 7, 8) accounted more than half of all samples sequenced (median count exceeding one) in each species, and were analyzed further. The most dominant OTU sequenced from

M. cavernosa was OTU 2 (91%; Fig 11), followed by OTU 4 (4%), 1 (2%), and 6 (2%).

The dominant OTU detected in P. clivosa was OTU 1 (96%), followed by small proportions of OTU 3 (3%).

OTU sequence query blast against the NCBI GenBank nucleotide library

(Table 1) matched the 19 OTUs to the following Symbiodinium types: B1 and B224

A4 (Gymnodinium linucheae), uncultured clade A, and Symbiodinium clade C isolates from Turbinaria peltata (hereafter designated type CX.1) and Stylocoeniella guentheri (designated type CX.4) collected in Japan, and from a parasitic flatworm,

Amakusaplana acroporae, collected from coral aquarists in the United Kingdom

(called type CX.3). All OTUs aligned to reference sequences at ≥ 96% except OTU 6

(CX.2), where percent identity was only 88%, indicating a high amount of dissimilarity among any documented Symbiodinium spp. sequence.

Generalized linear mixed modeling analysis of the 6 retained OTUs from M. cavernosa were modeled reliably and pairwise comparisons between all pairs of factor combinations revealed that OTUs 1, 2, 4, 6, and 9 (B1, CX.1, CX.3, CX.2, and

CX.4, respectively) were present in >99% of all samples (Fig 12). After adjusting p- values for multiple comparisons, only OTU 1_B1 and OTU 9_CX.4 differed by site over time. The proportion of OTU 1_B1 increases during Winter 2013, and becomes

33 less frequent during the subsequent sampling seasons. OTU 9_CX.4 appears to fluctuate across all reef sites. Within P. clivosa, OTUs 1, 3, 5, 7, 8 (all B1) fit the

MCMC.OTU model reliably. After calculating differences between all pairs of factor combinations, OTUs 1, 3, 5, and 7 (all B1) retained significant abundances (>99%) within P. clivosa by site over time (Fig 13). However, none of these OTUs changed over time or by reef site, and this consistency is evident in Figure 11 and 13.

The first principal coordinate analysis (PC1) of relative OTU abundance in M. cavernosa samples by colony and season (Fig 14a) or site (Fig 14b) explained 52.5% of the variation in OTUs present in ≤90% of all samples. The second coordinate

(PC2) explained 23.3% of the variation. Symbiodinium in corals at the South site grouped together slightly and differed from other reef sites, indicating OTU assemblage similarity. Samples from Winter 2013 had a slight negative loading relative to other seasons, possibly from the increased proportion of OTU 1_B1. PC1 from the P. clivosa PCA (Fig 15a and b) explained 51.3% of the variation in OTU representation, and PC2 explained 20% of the variation. A positive loading by North site samples is mostly explained by PC1. Similar to M. cavernosa, OTU representation in samples are grouped at the South site, as well as the Central reef site, suggesting assemblage similarities within each site. In addition, there is no visible partitioning of season, which could be explained by the increased spread of

North site samples.

Stepwise-paired correlation matrices between phylotypes portrays the relationship between two OTUs (Appendix B and C). Within M. cavernosa, OTU

2_CX.1 and 6_CX.2, 2_CX.1 and 11_CX.1, and 6_CX.2 and 11_CX.1 have slight positive

34 correlations (r = 0.11, 0.34, 0.29 respectively) with p-values of < 0.05 (Appendix B) indicating that as OTU 2_CX.1 increases, OTU 6 CX.2and 11CX.1 increase as well.

OTU 4_CX.3 compared against OTU 11_CX.1, has a correlation coefficient of 0.37, and could indicate that OTU 11_CX.1 increases in proportion alongside increases in OTU

4_CX.3. In addition, all OTUs designated to clade C showed negative relationships against OTU 1_B1, supporting competing representation between the two clades.

Relationships between OTUs 1_B1 and 3_B1, and 1_B1 and 8_B1 from P. clivosa show significant negative correlations (r = -0.92, -0.88 and p ≤ 0.001, respectively;

Appendix C), where OTU 3_B1 and 8_B1 representations decrease with an increase in OTU 1_B1. Additional pairwise comparisons including OTUs 3_B1 and 5_B1, 5_B1 and 8_B1, and 7_B1 and 8_B1 (r = -0.16, -0.37, -0.17 and p > 0.05) also show negative relationships. Although OTU 3_B1 and 8_B1 are different than OTU 1_B1, they are positively correlated to each other (r = 0.81 and p < 0.001), indicating similar assemblage representation in all samples and a positive interaction between the two may support coexistence.

DISCUSSION

M. cavernosa predominantly hosts Symbiodinium types CX.1, B1, and CX.3, which account for 91%, 2%, and 4% of all mapped ITS2 sequence reads, respectively. Conversely, P. clivosa was found to contain Symbiodinium phylotype B1 at 97% of mapped reads, followed by a genetic variant of B1 at 3%, and OTUs assigned to clade C types account for only 0.07%. We did not detect any significant changes in dominant symbiont proportion among the four reef sites, suggesting the types discovered at SLR are adapted to local regimes at the reef (Savage et al. 2002, 35 Rodriguez-Lanetty et al. 2003, LaJeunesse et al. 2010). Clade B1 in M. cavernosa did significantly increase in abundance at the Central and Ledge site during Winter

2013. Assigned proportions that generally accounted for 0.5 - 3% of total mapped reads from other sampling events increased to 8%, where abundance at the Ledge was 5% and 24% at the Central site. Symbiodinium B1 has been shown to be a host- generalist (LaJeunesse 2002, Finney et al. 2010), and could have the potential to become abundant from background populations in M. cavernosa if the local environment remains unperturbed. However, the abundance of B1 did decline thereafter, and still within the dry season (November – May) as other resident

Symbiodinium may have increased to diminish B1 representation. Quantifying relative abundances of Symbiodinium types cannot conclude absolute assemblage composition, and shifts in representation can only be inferred for this study.

In addition to the lack of change in of Symbiodinium abundance over space,

Symbiodinium assemblages within both coral species remained relatively constant over this study period, despite environmental variability occurring near SLR. Only type B1 and CX.4 within M. cavernosa significantly changed during Winter 2013 (Fig.

3), but these low frequency strains account for 7 and 0.3% of mapped reads and may not be functionally significant for overall holobiont performance. Determining fine-scale genetic responses in symbiont composition following disturbance events is one potential advantage of using highly sensitive molecular techniques such as deep sequencing. Jones and Yellowlees (1997) report that complete replacement of symbiont assemblages (‘shuffling’ or ‘switching’, Baker 2003) following disturbance can take at least a month (see also Dimond et al. 2013). Unfortunately, coral

36 sampling after the two major freshwater discharge releases in Summer and Fall

2013 occurred 3 – 4 weeks after flow rate increased exponentially. If genetic composition were to change following discharge events, sampling may have occurred too early. In addition, the time frame for Symbiodinium assemblage modification following disturbances is unknown, as well as whether symbiont populations within these two host species actually ‘switch’ or ‘shuffle’ (Thornhill et al. 2006, LaJeunesse et al. 2009). The results of this study suggest it is unlikely that the dominant Symbiodinium in each coral changes, but future samplings should aim to collect closer to the discharge release and repetitively during subsequent months to potentially detect any short-term trends.

Next-generation sequencing detected novel Symbiodinium types and associations within M. cavernosa and P. clivosa, suggesting that evolutionary genetic differentiation occurred in Caribbean populations to produce locally adapted

Symbiodinium in corals found at SLR. Although Symbiodinium phylotype B1 has been reported in P. clivosa within the Caribbean (Finney et al. 2010; Fig 16), this is the first report of its occurrence in M. cavernosa. Additionally, the clade C variants detected at SLR represent divergent lineages, as previous studies have documented variants of clade C3, and only within M. cavernosa not P. clivosa (LaJeunesse 2002,

Savage et al. 2002, Finney et al. 2010). Haplotype network analysis between these variants can illustrate whether the C types from SLR indeed have differentiated from type C3. Since both M. cavernosa and P. clivosa can associate with multiple types not previously recognized for either coral host, it is possible that the Symbiodinium types discovered at SLR are more suited to a particular host, the hosts exhibit a level

37 of selectivity on dominant symbionts (LaJeunesse et al. 2002, LaJeunesse 2003,

Goulet 2006), or selection of types capable of surviving at SLR occurred. Since these two coral species acquire their Symbiodinium horizontally (Szmant 1986), they have the propensity to build assemblages with types adapted to the local environmental conditions.

Although M. cavernosa and P. clivosa contain different Symbiodinium assemblages, both corals are the dominant scleractinian species in a coastal reef that experiences frequent bouts of environmental variability. These assemblages do not significantly change following freshwater discharge events, at least not 2 – 4 weeks afterwards, which corresponds to the time frame for coral sampling. Implications for coral performance under fluctuating water quality conditions without consequent changes in symbiont types could be attributed to many factors, including modifications in Symbiodinium structure. Genetic allopatry of some

Symbiodinium clade C types found further south in the Caribbean may restrict connectivity (Finney et al. 2010, Thornhill et al. 2014) and precludes flexible symbiont responses, although this remains to be examined. Moreover, symbiont physiological functioning rather than assemblage modifications under poor environmental conditions could substantiate continued but lowered growth rates of coral hosts at SLR. Though the dominant associated types, CX.1 in M. cavernosa and

B1 in P. clivosa, are morphologically distinct, it is likely both symbionts respond similarly under changing environments and contribute to overall holobiont fitness.

Another possible explanation for community-level similarities in holobiont fitness under different Symbiodinium types is adaptation through the formation of

38 stable symbioses (Thornhill et al. 2005, Putnam et al. 2012). The tendency for type

B1 to dominate P. clivosa and M. cavernosa to associate with mainly CX.1 indicates some level of host specificity (LaJeunesse et al. 2010) or coral host mediated exclusion or inclusion (Davy et al. 2012, Kemp et al. 2014). Symbiodinium types discovered can coexist in hospite, and the stable symbioses at SLR could have coevolved to withstand environmental disequilibrium characterized by stable dry seasons and hypervariable wet seasons. This is analogous to the ‘Intermediate

Disturbance Hypothesis’ (Connell 1978) that diversity and species composition is maximized under frequent episodes of disturbance. Dynamic water quality conditions can incite low levels of interspecific competition between locally adapted

Symbiodinium and could allow coexistence of genetically diverse symbiont types among different coral hosts (Putnam et al. 2012, Silverstein et al. 2012). Had uniform environmental conditions occurred at SLR, we may have seen significant increases in clade B1 to competitively exclude resident symbionts and potentially reduce local symbiont diversity. Alternatively, too much environmental variability could corroborate stable Symbiodinium associations including those adapted to local stressors and eliminating species incapable of colonizing (McGinley et al. 2012).

Stable patterns in symbiotic associations have been observed in several coral species when exposed to environmental variability (Thornhill et al. 2005, Putnam et al. 2012, McGinley et al. 2012, Dimond et al. 2013, Kenkel et al. 2013). Acclimatory responses to stressors may be suppressed in holobionts located in stable environments when exposed to infrequent stress events (Thornhill et al. 2005), and the results of this study and other research (McGinley et al. 2012, Putnam et al.

39 2012) indicates historical exposure to environmental stressors could modulate adapted associations. Moreover, flexible symbioses are suggested to be maladaptive and may be metabolically costly for coral hosts through increased interspecific competition among Symbiodinium types following stress events (Putnam et al.

2012). Therefore, preconditioned symbioses may be able to outperform conspecific corals from homogeneous environments (Kenkel et al. 2013), and the associations observed at SLR may be representative of future coastal, subtropical reef composition if reefs are increasingly subject to terrestrial influences.

Many corals were previously thought to preferentially host specific

Symbiodinium types (LaJeunesse 2002, LaJeunesse et al. 2003, 2005, Goulet 2006), but it is now suggested a gradient of specificity exists over which dominant and background populations vary in frequency (Silverstein et al. 2012). Although SLR corals contain mixed, background populations of Symbiodinium, it is uncertain whether these background populations offer any functional advantage to the coral host attributing to stress tolerance (Silverstein et al. 2012, Barshis et al. 2010).

Analyzing Symbiodinium genetic structure from other coral species at SLR may reveal additional symbiotic associations inclusive of these background symbiont and could aid in explaining their prevalence at this reef. While fine-scale diversity can be measured via deep sequencing (Quigley et al. 2014), the proportion of symbionts detected cannot yet elucidate functionality (Green et al. 2014). In addition, sequence diversity and abundance does not directly correlate to cellular quantity of ITS2 symbiont types, so care should be taken when assigning overall holobiont fitness to Symbiodinium type abundances.

40 The examination of Symbiodinium assemblage diversity and abundances over time within two different corals species, M. cavernosa and P. clivosa, provides evidence that stable, adapted associations occur in coastal reefs influenced by environmental perturbations. Next-generation sequencing of Symbiodinium revealed types B1, CX.1, CX.4, and CX.3 that could represent locally adapted ecotypes that differentially associate between the two coral species at SLR. Additionally, the low identity coverage of type CX.2 to any documented Symbiodinium type suggests this variant could be a new species, and further analyses utilizing haplotype networks can elucidate relatedness or novelty of this type. Assessment of coral-algal associations in conspecifics at coastal reefs down the Florida tract could test the hypothesis that adaptive radiations and coevolved symbioses have indeed formed.

Local cultivated associations have important implications for management strategies as these could be reflective of future reefs adjacent to coastal communities, especially if the rate of selection in a more sensitive coral holobiont is outpaced by expanding anthropogenic practices and climate change.

41 APPENDICES

42 PCR Mapped Mapping Species Season Site Colony Read # % B1 %CX.1 %CX.3 %CX.2 cycles Reads Efficiency M. cavernosa Spring 2012 Central 2 31 780 471 0.60 0.03 0.96 0.01 0.00 6 31 3416 2057 0.60 0.01 0.95 0.04 0.00 Ledge 2 31 12486 7635 0.61 0.01 0.97 0.02 0.00 5 31 5526 3276 0.59 0.01 0.95 0.03 0.01 6 24 5612 2436 0.43 0.00 0.97 0.03 0.00 8 30 3655 1460 0.40 0.00 0.96 0.03 0.00 South 1 31 1453 595 0.41 0.02 0.93 0.04 0.00 2 31 4310 1757 0.41 0.01 0.94 0.05 0.00 3 31 2098 939 0.45 0.01 0.90 0.09 0.00 4 31 928 389 0.42 0.05 0.91 0.04 0.00 5 31 5100 2612 0.51 0.01 0.96 0.03 0.00 Winter 2013 Central 2 28 4449 2548 0.57 0.08 0.88 0.04 0.00 4 31 6966 4077 0.59 0.39 0.58 0.03 0.00 6 31 1154 637 0.55 0.05 0.92 0.03 0.00 Ledge 3 27 13091 7572 0.58 0.01 0.92 0.06 0.01 4 25 8333 4889 0.59 0.04 0.91 0.05 0.00 5 27 4596 2648 0.58 0.08 0.87 0.05 0.01 6 27 5584 3284 0.59 0.04 0.91 0.05 0.00 8 31 9883 5747 0.58 0.10 0.86 0.04 0.00 South 1 27 1331 756 0.57 0.06 0.88 0.05 0.00 2 30 2583 1453 0.56 0.08 0.88 0.04 0.00 3 30 1162 690 0.59 0.06 0.89 0.05 0.00 4 30 2653 1613 0.61 0.01 0.93 0.06 0.00 5 30 2833 1600 0.56 0.02 0.93 0.05 0.00 Spring 2013 Central 2 29 7358 4232 0.58 0.03 0.92 0.04 0.01 4 30 14234 8494 0.60 0.01 0.96 0.03 0.00 6 30 19048 11216 0.59 0.01 0.94 0.05 0.01 7 25 8318 4846 0.58 0.00 0.93 0.06 0.01 Ledge 1 31 8206 4802 0.59 0.00 0.94 0.06 0.00 2 31 5449 3200 0.59 0.01 0.95 0.03 0.01 3 31 8795 5304 0.60 0.00 0.94 0.05 0.01 4 31 14494 8603 0.59 0.01 0.95 0.03 0.01 5 31 9781 5926 0.61 0.00 0.96 0.04 0.00 6 27 11635 6870 0.59 0.27 0.69 0.03 0.00 8 31 12896 7680 0.60 0.01 0.95 0.04 0.01 South 1 31 6258 3523 0.56 0.03 0.93 0.04 0.00 2 31 3961 2247 0.57 0.04 0.93 0.03 0.00 3 31 3713 2089 0.56 0.02 0.93 0.05 0.00 4 31 7671 4369 0.57 0.01 0.95 0.04 0.00 5 31 4096 2345 0.57 0.02 0.92 0.06 0.00 Summer 2013 Central 2 29 11328 5961 0.53 0.01 0.96 0.03 0.00 4 31 6023 3110 0.52 0.02 0.93 0.05 0.00 7 31 3999 2077 0.52 0.01 0.94 0.04 0.01 Ledge 1 25 11138 6194 0.56 0.00 0.93 0.06 0.00 3 23 12398 6940 0.56 0.00 0.95 0.05 0.00 4 23 11050 7676 0.69 0.01 0.91 0.03 0.05 5 23 10713 5905 0.55 0.00 0.95 0.04 0.01 6 27 6276 3553 0.57 0.01 0.95 0.04 0.00 8 27 6506 3307 0.51 0.01 0.95 0.04 0.01 South 1 27 11678 6521 0.56 0.01 0.94 0.05 0.00 2 27 7889 4442 0.56 0.01 0.95 0.04 0.00 4 20 10467 6001 0.57 0.00 0.97 0.03 0.00 5 29 12847 7643 0.59 0.00 0.97 0.03 0.00

43 PCR Mapped Mapping Species Season Site Colony Read # % B1 %CX.1 %CX.3 %CX.2 cycles Reads Efficiency M. cavernosa Fall 2013 Central 2 31 10567 5470 0.52 0.00 0.96 0.03 0.01 4 31 5488 2960 0.54 0.01 0.97 0.02 0.00 7 31 1640 735 0.45 0.09 0.86 0.05 0.00 Ledge 1 21 11897 7182 0.60 0.01 0.93 0.03 0.03 3 20 8703 4745 0.55 0.01 0.94 0.05 0.00 4 21 7375 3986 0.54 0.00 0.94 0.04 0.01 5 27 8853 4661 0.53 0.01 0.94 0.05 0.01 6 21 12660 6516 0.51 0.01 0.93 0.06 0.00 8 27 9156 5043 0.55 0.00 0.95 0.04 0.01 South 1 31 4260 2194 0.52 0.01 0.95 0.04 0.00 2 29 7647 3848 0.50 0.01 0.95 0.04 0.00 3 31 9323 4588 0.49 0.02 0.96 0.02 0.00 4 31 13127 7208 0.55 0.00 0.97 0.03 0.00 5 31 4591 2475 0.54 0.01 0.97 0.02 0.00 P. clivosa Spring 2012 North 1 30 3487 2094 0.60 1.00 0.00 0.00 0.00 2 30 9203 5188 0.56 1.00 0.00 0.00 0.00 4 26 17809 10646 0.60 1.00 0.00 0.00 0.00 6 22 76 32 0.42 1.00 0.00 0.00 0.00 Central 1 25 26757 16038 0.60 1.00 0.00 0.00 0.00 3 23 9216 5515 0.60 1.00 0.00 0.00 0.00 4 25 8307 5157 0.62 1.00 0.00 0.00 0.00 5 31 10907 6697 0.61 1.00 0.00 0.00 0.00 6 31 5908 3541 0.60 1.00 0.00 0.00 0.00 Ledge 1 31 12921 7915 0.61 1.00 0.00 0.00 0.00 2 31 4859 2883 0.59 1.00 0.00 0.00 0.00 4 30 506 292 0.58 0.99 0.01 0.00 0.00 5 30 5327 3078 0.58 1.00 0.00 0.00 0.00 South _ 30 7437 5417 0.73 1.00 0.00 0.00 0.00 1 22 8376 4506 0.54 1.00 0.00 0.00 0.00 2 31 16484 4939 0.30 1.00 0.00 0.00 0.00 3 31 6110 9742 1.59 1.00 0.00 0.00 0.00 4 31 7716 3564 0.46 1.00 0.00 0.00 0.00 5 25 9083 4985 0.55 1.00 0.00 0.00 0.00 Winter 2013 North 2 19 13589 7087 0.52 1.00 0.00 0.00 0.00 4 19 13594 7181 0.53 1.00 0.00 0.00 0.00 Central 1 24 13506 8277 0.61 1.00 0.00 0.00 0.00 3 24 7620 4579 0.60 1.00 0.00 0.00 0.00 4 24 14169 8760 0.62 1.00 0.00 0.00 0.00 5 24 19354 11937 0.62 1.00 0.00 0.00 0.00 6 21 14477 8935 0.62 1.00 0.00 0.00 0.00 Ledge 1 19 10287 6262 0.61 1.00 0.00 0.00 0.00 2 19 12590 7831 0.62 1.00 0.00 0.00 0.00 4 19 9371 5985 0.64 1.00 0.00 0.00 0.00 6 19 15516 9533 0.61 1.00 0.00 0.00 0.00 South 1 21 10691 6779 0.63 1.00 0.00 0.00 0.00 2 31 8719 5454 0.63 1.00 0.00 0.00 0.00 3 22 13733 8229 0.60 1.00 0.00 0.00 0.00 4 24 9709 5722 0.59 1.00 0.00 0.00 0.00 5 28 11287 6845 0.61 1.00 0.00 0.00 0.00 Spring 2013 North 1 19 12324 10012 0.81 1.00 0.00 0.00 0.00 17 19 20128 7668 0.38 1.00 0.00 0.00 0.00 18 19 16060 12452 0.78 1.00 0.00 0.00 0.00 2 19 11156 6767 0.61 1.00 0.00 0.00 0.00 4 19 7368 4548 0.62 1.00 0.00 0.00 0.00 6 19 29921 19130 0.64 1.00 0.00 0.00 0.00 Central 1 19 12838 8181 0.64 1.00 0.00 0.00 0.00 3 19 8140 5088 0.63 1.00 0.00 0.00 0.00 4 19 13143 5366 0.41 1.00 0.00 0.00 0.00 5 19 4706 1953 0.42 1.00 0.00 0.00 0.00 6 19 5386 2257 0.42 1.00 0.00 0.00 0.00

44 PCR Mapped Mapping Species Season Site Colony Read # % B1 %CX.1 %CX.3 %CX.2 cycles Reads Efficiency P. clivosa Ledge 1 25 3158 1134 0.36 1.00 0.00 0.00 0.00 2 19 3166 1291 0.41 1.00 0.00 0.00 0.00 4 19 3606 1469 0.41 1.00 0.00 0.00 0.00 5 19 5233 2524 0.48 1.00 0.00 0.00 0.00 6 19 4918 2398 0.49 1.00 0.00 0.00 0.00 South 1 19 6134 2852 0.46 1.00 0.00 0.00 0.00 2 19 5574 2587 0.46 1.00 0.00 0.00 0.00 3 19 4435 2051 0.46 1.00 0.00 0.00 0.00 4 19 5952 2585 0.43 1.00 0.00 0.00 0.00 5 20 13630 8516 0.62 1.00 0.00 0.00 0.00 Summer 2013 North 1 21 12855 8069 0.63 1.00 0.00 0.00 0.00 17 19 13081 7177 0.55 1.00 0.00 0.00 0.00 18 19 13866 6561 0.47 1.00 0.00 0.00 0.00 2 20 13033 7611 0.58 1.00 0.00 0.00 0.00 4 20 9903 5619 0.57 1.00 0.00 0.00 0.00 6 19 16119 9917 0.62 1.00 0.00 0.00 0.00 Central 1 19 10361 6064 0.59 1.00 0.00 0.00 0.00 3 23 18481 10986 0.59 1.00 0.00 0.00 0.00 4 20 15772 9221 0.58 1.00 0.00 0.00 0.00 5 20 26251 15618 0.59 1.00 0.00 0.00 0.00 6 20 11028 6303 0.57 1.00 0.00 0.00 0.00 Ledge 1 19 20859 12214 0.59 1.00 0.00 0.00 0.00 2 21 14447 8579 0.59 1.00 0.00 0.00 0.00 4 20 13511 8170 0.60 1.00 0.00 0.00 0.00 5 19 17056 10237 0.60 1.00 0.00 0.00 0.00 6 19 20337 12380 0.61 1.00 0.00 0.00 0.00 South 1 19 14087 8446 0.60 1.00 0.00 0.00 0.00 2 19 13934 8486 0.61 1.00 0.00 0.00 0.00 3 19 17597 10772 0.61 1.00 0.00 0.00 0.00 4 19 12746 6625 0.52 1.00 0.00 0.00 0.00 5 20 6977 3492 0.50 1.00 0.00 0.00 0.00 6 20 12367 6379 0.52 1.00 0.00 0.00 0.00 Fall 2013 North 1 31 11576 2976 0.26 0.99 0.01 0.00 0.00 17 19 18446 7098 0.38 1.00 0.00 0.00 0.00 18 19 4812 11009 2.29 1.00 0.00 0.00 0.00 2 19 15578 9433 0.61 1.00 0.00 0.00 0.00 4 21 18321 11003 0.60 1.00 0.00 0.00 0.00 6 19 11956 7458 0.62 1.00 0.00 0.00 0.00 Central 1 19 16183 9902 0.61 1.00 0.00 0.00 0.00 3 19 8311 5009 0.60 1.00 0.00 0.00 0.00 4 21 9102 5616 0.62 1.00 0.00 0.00 0.00 5 19 10033 6032 0.60 1.00 0.00 0.00 0.00 6 21 9069 5634 0.62 1.00 0.00 0.00 0.00 Ledge 1 21 6927 4127 0.60 1.00 0.00 0.00 0.00 2 21 14654 8828 0.60 1.00 0.00 0.00 0.00 4 20 9498 5122 0.54 1.00 0.00 0.00 0.00 5 20 10854 6668 0.61 1.00 0.00 0.00 0.00 6 23 6447 3882 0.60 1.00 0.00 0.00 0.00 South 1 19 11893 7549 0.63 1.00 0.00 0.00 0.00 2 23 8009 5005 0.62 1.00 0.00 0.00 0.00 3 19 7428 4737 0.64 1.00 0.00 0.00 0.00 4 19 9624 5900 0.61 1.00 0.00 0.00 0.00 5 21 16574 8710 0.53 1.00 0.00 0.00 0.00 6 21 9286 4824 0.52 0.99 0.01 0.00 0.00 Appendix A. Summary of ITS2 amplicon sequencing cycle number, coverage, and mapping efficiency at St. Lucie Reef. Samples are sorted by species, season, reef site, and colony.

45 7.8 8.0 8.2 3.5 4.5 0 1 2 3 7 5 1I_B1 -0.65 -0.041 -0.29 -0.029 -0.25 3 1 8.2 II_CJ -0.14 0.11 -0.089 0.34

8.0 2_CX.1 7.8 5.5 4_CX.3IV_CX 0.062 -0.043 0.37 4.5 3.5

0.051 4.5 6_CX.2VI_CJ 0.29 3.5 3

IX_CSJ -0.38 2 9_CX.4 1 0 3 2

1 11_CX.1XI_CJ 0

1 3 5 7 3.5 4.5 5.5 0 1 2 3

Appendix B. Multiple pairwise correlation matrix between dominant OTU frequencies among M. cavernosa colonies across all sites over all sampling events.

Scatterplots of the correlation relationship between two OTUs are displayed in cross- sections. Correlation coefficients are displayed in the upper right section of the matrix.

46 4.6 5.0 5.4 5.8 1.0 2.0 3.0 8.80

1I_B1 0.11 0.031 -0.92 -0.88 8.77 8.74 5.8

-0.16 -0.095 5.2 III_B13 0.81 4.6 4.0 V_B15 0.13 -0.37 3.0 3.0 VII_B17 -0.17 2.0 1.0 4

VIII_B18 2 0

8.74 8.76 8.78 8.80 3.0 3.5 4.0 4.5 0 1 2 3 4 5 Appendix C. Multiple pairwise correlation matrix between dominant OTU frequencies among P. clivosa colonies across all sites over all sampling events. Scatterplots of the correlation relationship between two OTUs are displayed in cross-sections. Correlation coefficients are displayed in the upper right section of the matrix.

47 FIGURES

Project Sites

North

10 Mile Central

C-24 Ledge

C-23 South

Stuart Inlet

C-44

Lake Okeechobee

Figure 1. Map of St. Lucie Estuary and Reef with study sites indicated by stars. The triangle shows where additional water quality loggers were installed. Left panel indicates water routes of canals entering St. Lucie estuary and stations where the SFWMD DBHYDRO continuously logged environmental variables.

48 a) b)

Figure 2. Density of zooxanthellae in millions of cells per square centimeter in relation to site and sampling event season for P. clivosa (a) and M. cavernosa (b). M. cavernosa colonies are not present at the North site. Samples were averaged and grouped by site and coral species. Error bars are expressed as standard error of the grouped mean.

Figure 3. Density of zooxanthellae in millions of cells per square centimeter in relation to sampling event season. Samples from the four sites are averaged for each coral species. Error bars are expressed as standard error of the grouped mean.

50 Figure 4. Chl a (a and b) and c (c and d) densities among P. clivosa and M. cavernosa over time. Cellular amount (a and c) is based on zooxanthellae density data. Concentrations of chl a (b) and c (d) is based on chlorophyll amount scaled up to coral fragment tissue area.

Samples from the four sites are averaged for each coral species. Error bars are expressed as standard error of the grouped mean.

51 a) b)

10 10

8 8

6 6 r = -0.096 Cellular Chlorophyll a Cellular Chlorophyll a 4 r = -0.345

2.0e+06 4.0e+06 6.0e+06 8.0e+06 1.0e+07 1.2e+07 1.4e+07+07 2e+06 4e+06 6e+06 8e+06 Zooxanthellae Density Zooxanthellae Density Figure 5. Correlation of cellular chlorophyll a (pg cell-1) and zooxanthellae density

(cells cm-2). a) Relationship between P. clivosa, and b) relationship between chl a and cell density in M. cavernosa. Pearson’s correlation coefficients are indicated above the regression line.

a)b b)) 35 r = 0.765 r = 0.859

40 30

25 30 20

15 20 Chlorophyll a Area Chlorophyll a Area

10 5

2.0e+06 4.0e+06 6.0e+06 8.0e+06 1.0e+07 1.2e+07 1.4e+07+07 2e+06 4e+06 6e+06 8e+06 Zooxanthellae Density Zooxanthellae Density Figure 6. Correlation of chlorophyll a scaled to coral tissue (μg cm-1) and zooxanthellae density (cells cm-2). a) Relationship between between chl a per coral surface area and cell density in P. clivosa, and b) relationship between chl a per coral surface area and cell density in M. cavernosa. Pearson’s correlation coefficients are indicated above the regression line.

52 Figure 7. Cell volume (μm3) of zooxanthellae cells from P. clivosa and M. cavernosa collected Summer 2013. Mean cell sizes are displayed by species and pooled by site.

53 Figure 8. Chlorophyll a content relative to zooxanthellae cell volume from Summer

2013 samples. Mean cellular chlorophyll a content (pg) in zooxanthellae from P. clivosa and M. cavernosa scaled to mean cell volume.

54 Figure 9. Discharge flow rates of canals in Lake Okeechobee and the St. Lucie Estuary

Watershed. Each line represents continuous flow data in cubic meters per second, which was obtained from five stations (canals) of the South Florida Water Management District’s

DBHYDRO database. Solid stars represent the five coral sampling events at SLR.

Figure 10. Environmental variables at St. Lucie Inlet. Salinity (a) and temperature (b) recorded within the St. Lucie Inlet by the SFWMD DBHYDRO database during five sampling events.

CX.2, CX.4

B1, CX.1

Figure 11. Proportion of Symbiodinium OTUs present in more than half of coral hosts. Pie charts are split by coral host and separated by sampling event season. OTUs representing < 0.005 of an assemblage from each coral host,

OTU4_CX.3, OTU5_B1, OTU7_B1, OTU9_CX.4, and OTU10_B1 are not visible.

OTU Season Species Clade0 Accession0ID %0Identity0 E0Value Identity Reference Source OTU1 Summer*2013 P.#clivosa B1 JN558059.1 100 4.00E787 B1*v*704*c*3*5.8S Pochon#et#al.*2012 Plexaura#kuna,*Panama Lien*YT,*Fukami*H,*Yamashita* Turbinaria#peltata,*Okinawa,* OTU2 Winter*2013 M.#cavernosa Cx1 JX434382.1 100 4.00E787 C*isolate*Tub06*c1 (2012,*Direct*Submission) Japan OTU3 Winter*2013 M.#cavernosa B1 JN558059.1 100 4.00E787 B1*v*704*c*3*5.8S Pochon#et#al.*2012 Plexaura#kuna,*Panama* Hume*BCC,*Wiedenmann*J* Amakusaplana#acroporae* OTU4 Winter*2013 M.#cavernosa Cx3 JN711493.1 100 4.00E787 C*isolate*BH34 (2011,*Direct*Submission) (flatworm),*UK OTU5 Winter*2013 M.#cavernosa B1 JN558059.1 98.88 1.00E783 B1*v*704*c*3*5.8S Pochon*et#al.*2012 Plexaura#kuna,*Panama Lien*YT,*Fukami*H,*Yamashita* Turbinaria#peltata,*Okinawa,* OTU6 Winter*2013 M.#cavernosa Cx2 JX434382.1 87.68 5.00E752 C*isolate*Tub06*c1 (2012,*Direct*Submission) Japan OTU7 Winter*2013 M.#cavernosa B1 JN558059.1 99.44 2.00E785 B1*v*704*c*3*5.8S Pochon*et#al.*2012 Plexaura#kuna,*Panama OTU8 Winter*2013 M.#cavernosa B1 JN558059.1 96.97 6.00E771 B1*v*704*c*3*5.8S Pochon#et#al.*2012 Plexaura#kuna,*Panama Lien*YT,*Fukami*H,*Yamashita* Stylocoeniella#guentheri,* OTU9 Winter*2013 M.#cavernosa Cx4 JX434262.1 97.75 2.00E780 C*isolate*Sty.gue_TNS31*c*3 (2012,*Direct*Submission) Tanegashima,*Japan OTU10 Winter*2013 P.#clivosa B24 GU907643.1 99.37 3.00E774 B24*isolate*BEL02_46 Finney*et#al.*2010 Diploria,*Belize Lien*YT,*Fukami*H,*Yamashita* Turbinaria#peltata,*Okinawa,* OTU11 Winter*2013 M.#cavernosa Cx1 JX434382.1 100 4.00E787 C*isolate*Tub06*c1 (2012,*Direct*Submission) Japan OTU12 Winter*2013 M.#cavernosa B1 FJ811916.1 96.57 3.00E774 B1*c*C4_7*5.8S DeSalvo#et#al.#2010 Obicella#faveolata OTU13 Winter*2013 M.#cavernosa B1 JN558059.1 100 4.00E787 B1*v*704*c*3*5.8S Pochon*et#al.*2012 Plexaura#kuna,*Panama

58 Lien*YT,*Fukami*H,*Yamashita* Turbinaria#peltata,*Okinawa,* OTU14 Winter*2013 M.#cavernosa Cx1 JX434382.1 96.22 4.00E777 C*isolate*Tub06*c1

(2012,*Direct*Submission) Japan OTU15 Winter*2013 P.#clivosa B1 JN558059.1 100 4.00E787 B1*v*704*c*3*5.8S Pochon*et#al.*2012 Plexaura#kuna,*Panama OTU16 Winter*2013 M.#cavernosa B2 JN558062.1 100 4.00E787 B2*v*Pflex*c*3*5.8S Pochon#et#al.*2012 Eunicea#flexuosa,*HI OTU17 Spring*2013 M.#cavernosa A4 AF333509.1 100 4.00E787 Gymnodinium#linucheae#(A4)*18S LaJeunesse*2001 Linuche#unguiculata# OTU18 Spring*2013 M.#cavernosa A AB849697.1 100 4.00E787 Uncultured*Symbiodinium*clade*A Yamashita*H,*Suzuki*G*2013 Sea*water,*Okinawa,*Japan Santos*SR,*Taylor*DJ,*Coffroth* OTU19 Winter*2013 P.#clivosa B AF360569.1 96.09 7.00E775 Symbiodinium*clade*B* Plexaura#homomalla,*Panama MA*2001 Table 1. OTU sequence NCBI BLASTn matches. Each OTU was queried against the GenBank nucleotide reference library, and the closest

matched strain’s Accession number, % identity, E value, reference, and host organism are provided. The sample from which the OTU was

identified is delimited by sampling event, site, species and colony number. Clade column corresponds to the designated Symbiodinium

type for each OTU based on BLASTn results. Source column refers to zooxanthellae extracted from host organism.

C L S

-2 M.cavernosa OTU OTU1_B1 OTU2_CJOTU2_CX.1 OTU4_CXOTU4_CX.3 -4 OTU6_CJOTU6_CX.2

log10(proportion) OTU9_CSJOTU9_CX.4

-6

-8

Fall2013 Fall2013 Fall2013

Spring2012Winter2013Spring2013 Spring2012Winter2013Spring2013 Spring2012Winter2013Spring2013 Summer2013 Summer2013 Summer2013 Season

Figure 12. Proportion of significant Symbiodinium OTUs in M. cavernosa by site across five sampling events. Distribution in relative abundance (log10 transformed) of five OTUs that MCMC.OTU modeled by calculating differences and p-values between all pairs of factor combinations. OTU1_B1, OTU2_ CX.1, OTU4_CX.3, OTU6_CX.2, and OTU9_CX.4 autocorrelate among sample parameter values. Sites are C: Central, L: Ledge, and S: South.

C L

-2

-4

-6 P.clivosa OTU -8 OTU1_B1 N S OTU3_B1 OTU5_B1 0 OTU7_B1 log10(proportion)

-2

-4

-6

-8

Fall2013 Fall2013

Spring2012 Winter2013 Spring2013 Spring2012 Winter2013 Spring2013 Summer2013 Summer2013 Season

Figure 13. Proportion of significant Symbiodinium OTUs in P. clivosa by site across five sampling events. Distribution in relative abundance (log10 transformed) of five OTUs that MCMC.OTU modeled by calculating differences and p-values between all pairs of factor combinations. OTU1_B1, OTU3_B1, OTU5_B1, and OTU7_B1 autocorrelate among sample parameter values. Sites are C: Central, L: Ledge, N: North, and S: South.

Figure 14. Principal coordinate analysis of Symbiodinium communities in M. cavernosa in relation to season (a) and site (b). Points in both plots represent log- transformed OTUs that are present in more than 10% of samples. Partitioning of samples is the same in both plots, but each are color-coded and shaped based on the different factor levels. a) Differences with respect to season. b) Differences with respect to reef site.

Figure 15. Principal coordinate analysis of Symbiodinium communities in P. clivosa in relation to season (a) and site (b). Points in all plots represent log-transformed OTUs that are present in more than 10% of samples. Partitioning of samples is the same in both plots,

but each are color-coded and shaped based on the different factor levels. a) Differences in relation to season. b) Differences with respect to reef site.

CX.2, CX.3, CX.4 Montastraea cavernosa B24 Pseudodiploria clivosa B1, CX.1 Both C3e C3

C3d, C3e

C3d, C3e, C3g B1

C3o, C3p B1 LaJeunesse 2002, Savage et al. 2002, Finney et al. 2010, LaJeunesse et al. 2010, this D1a study Figure 16. Symbiodinium diversity in the Wider Caribbean. Previous studies determining Symbiodinium types in corals M. cavernosa and P. clivosa are used for comparison with this study. Diagonally hashed boxes indicate Symbiodinium types found in both coral host species. (Map credit: http://en.wikipedia.org/wiki/Caribbean#mediaviewer/File:CIA_map_of_the_Caribb ean.png)

REFERENCES

Abrego D, Ulstrup KE, Willis BL, van Oppen MJ (2008) Species–specific interactions

between algal endosymbionts and coral hosts define their bleaching response

to heat and light stress. ProcR Soc B 275:2273-2282.

Baker AC, Rowan R (1997) Diversity of symbiotic dinoflagellates (zooxanthellae) in

scleractinian corals of the Caribbean and eastern Pacific. In Proc 8th Int Coral

Reef Symp 2:1301-1306.

Baker AC (2001) Ecosystems: reef corals bleach to survive change. Nature 411:765-

766.

Baker AC (2003) Flexibility and specificity in coral-algal symbiosis: diversity,

ecology, and biogeography of Symbiodinium. Annu Rev Ecol Evol Syst 34:661–

689.

Baker AC, Starger CJ, McClanahan TR, Glynn PW (2004) Coral reefs: corals' adaptive

response to climate change. Nature 430:741-741.

Barshis DJ, Stillman JH, Gates RD, Toonen RJ, Smith LW, Birkeland C (2010) Protein

expression and genetic structure of the coral Porites lobata in an

environmentally extreme Samoan back reef: does host genotype limit

phenotypic plasticity? Molec Ecol 19: 1705-1720.

Beal J, Voss JD, Edge S, Cohen L (2012) Assessment of coral stressors on SLR:

Florida’s northernmost coral reef. Final Report for State Wildlife Grant. USFWS

T-19-1.

Bellantuono AJ, Hoegh-Guldberg O, Rodriguez-Lanetty M (2011) Resistance to

thermal stress in corals without changes in symbiont composition. Proc R Soc B

279:1100–1107.

Bellantuono AMS, Baker AC (2008) Understanding diversity in coral-algal

symbiosis: a cluster-based approach to interpreting fine-scale genetic variation

in the genus Symbiodinium. Coral Reefs 28:81–93.

Berkelman R, van Oppen MJ (2006) The role of zooxanthellae in the thermal

tolerance of corals: a ‘nugget of hope’ for coral reefs in an era of climate change.

Proc R Soc B 273:2305-2312.

Brown BE, Dunne RP, Ambarsari I, Le Tissier MDA, Satapoomin U (1999) Seasonal

fluctuations in environmental factors and variations in symbiotic algae and

chlorophyll pigments in four Indo-Pacific coral species. Mar Ecol Prog Ser

191:53–69.

Buddemeier RW, Fautin DG (1993) Coral bleaching as an adaptive mechanism.

Bioscience 43:320-326.

Carricart-Ganivet JP, Merino M (2001) Growth responses of the reef-building coral

Montastraea annularis along a gradient of continental influence in the southern

Gulf of Mexico. Bull Mar Sci 68:133-146.

Chen CA, Wang J-T, Fang L-S, Yang Y-W (2005) Fluctuating algal symbiont

communities in Acropora palifera (Scleractinia: Acroporidae) from Taiwan. Mar

Ecol Prog Ser 295:113-121.

Coles SL, Jokiel PL (1992) Effects of salinity on coral reefs In Connell DW, Hawker

DW (Eds) Pollution in tropical aquatic systems. CRC Press, the University of

California, 147-166.

Coles SL (1975) A comparison of effects of elevated temperature versus

temperature fluctuations on reef corals at Kahe Point, Oahu. Pac Sci 29:15-18.

Connell JH (1978) Diversity in tropical rain forests and coral reefs. Science 199:

1302-1310.

Costa CF, Sassi R, Amaral FD (2004) Population density and photosyntheitic pigment

content in symbiotic dinoflagellates in the Brazilian scleractinian coral

Montastraea cavernosa (Linneaus, 1767). Brazil Jour Oceanogr 52:93–99.

Cunning R, Baker AC (2012) Excess algal symbionts increase the susceptibility of

reef corals to bleaching. Nature Climate Change 2:1–4.

D’Angelo C, Wiedenmann J (2014) Impacts of nutrient enrichment on coral reefs:

new perspectives and implications for coastal management and reef survival.

Curr Opin Environ Sustainability 7:82-93.

Davies SW, Meyer E, Guermond SM, Matz MV (2014) A cross-ocean comparison of

responses to settlement cues in reef-building corals. PeerJ 2: e333.

Davy SK, Allemand D, Weis VM (2012) Cell biology of cnidarian-dinoflagellate

symbiosis. MMBR 76:229-261.

D’Croz L, Maté JL, Oke JE (2001) Responses to elevated sea water temperature and

UV radiation in the coral Porites lobata from upwelling and non-upwelling

environments on the Pacific coast of Panama. Bull Mar Sci 69:203–214.

D’Elia CF, Cook CB (1988) Methylamine uptake by zooxanthellae-invertebrate

symbioses: insights into host ammonium environment and nutrition. Limnol

Oceanogr 33:1153-1165.

Dimond JL, Bingham BL, Muller-Parker G, Oakley CA (2013) Symbiont physiology

and population dynamics before and during symbiont shifts in a flexible algal-

cnidarian symbiosis. Jour Phycol 49:1074-1083.

Dubinsky Z, Falkowski PG, Porter JW, Muscatine L (1984) Absorption and utilization

of radiant energy by light-and shade-adapted colonies of the hermatypic coral

Stylophora pistillata. Proc R Soc B 222: 203-214.

Dubinsky Z, Stambler N (2011) Coral Reefs: An Ecosystem in Transition. Springer,

New York.

Fabricius KE (2005) Effects of terrestrial runoff on the ecology of corals and coral

reefs: review and synthesis. Mar Poll Bull 50: 125–46.

Fagoonee I (1999) The dynamics of zooxanthellae populations: a long-term study in

the field. Science 283: 843–845.

Falkowski PG, Dubinsky Z (1981) Light-shade adaptation of Stylophora pistillata, a

hermatypic coral from the Gulf of Eilat. Nature 289:172–174.

Finney J, Pettay DT, Sampayo EM, Warner ME, Oxenford H, LaJeunesse TC (2010)

The relative significance of host-habitat, depth, and geography on the ecology,

endemism, and speciation of coral endosymbionts in the genus Symbiodinium.

Microb Ecol 60:250–263.

Fitt WK, Spero HJ, Halas J, White MW, Porter JW (1993) Recovery of the coral

Montastraea annularis in the Florida Keys after the 1987 Caribbean bleaching

event. Coral Reefs 12:57-64.

Fitt WK, Mcfarland FK, Warner ME, Chilcoat GC (2000) Seasonal patterns of tissue

biomass and densities of symbiotic dinoflagellates in reef corals and relation to

coral bleaching, Limnol Oceanogr 45: 677–685.

Gates RD, Baghdasarian G, Muscatine L (1992) Temperature stress causes host cell

detachment in symbiotic cnidarians: implications for coral bleaching. Biol Bull

182:324-332.

Glynn PW (1977) Coral growth in upwelling and non-upwelling areas off Pacific

coast of Panama. Jour Mar Research 35:567-585.

Goulet TL, Cook CB, Goulet D (2005) Effect of short-term exposure to elevated

temperatures and light levels on photosynthesis of different host-symbiont

combinations in the Aiptasia pallida/Symbiodinium symbiosis. Limnol Oceanog

50:1490-1498.

Goulet TL (2006) Most corals may not change their symbionts. Mar Ecol Prog Ser

321:1-7.

Graves GA, Wan Y, Fike DL (2004) Water quality characteristics of storm water from

major land uses in south Florida. JAWRA 40:1405-1419.

Green E, Davies SW, Matz MV, Medina M (2014) Next-generation sequencing reveals

cryptic Symbiodinium diversity within Orbicella faveolata and Orbicella franksi

at the Flower Garden Banks, Gulf of Mexico. PeerJ PrePrints No. e246v1.

Hoegh-Guldberg O (1994) The population dynamics of symbiotic zooxanthellae in

the coral Pocillopora damicornis exposed to elevated ammonium (NH4Cl)

concentrations. Pac Sci: 263-272.

Hoegh-Guldberg O, Mumby PJ, Hooten AJ, Steneck RS, Greenfield P, Gomez E,

Hatziolos ME (2007) Coral reefs under rapid climate change and ocean

acidification. Science 318:1737-1742.

Houlbreque F, Ferrier-Pagès C (2009) Heterotrophy in tropical scleractinian corals.

Biological Reviews 84:1-17.

Iglesias-Prieto R, Trench RK (1994) Acclimation and adaptation to irradiance in

symbiotic dinoflagellates. I. Responses of the photosynthetic unit to changes in

photon flux density. Mar Ecol Prog Ser 113:163-175.

Iglesias-Prieto R, Trench RK (1997) Acclimation and adaptation to irradiance in

symbiotic dinoflagellates. II. Response of chlorophyll–protein complexes to

different photon-flux densities. Mar Biol 130:23-33.

ISRS (2004) The effects of terrestrial runoff of sediments, nutrients, and other

pollutants on coral reefs. Briefing Paper 3, International Society for Reef

Studies, pp:18.

Jeffrey SW, Humphrey GF (1975) New spectrophotometric equations for

determining chlorophylls a, b, c1, and c2 in higher plants, algae, and natural

phytoplankton. Biochem Physiol Pflanzen 167:191-194.

Jones RJ (1997a) Changes in zooxanthellar densities and chlorophyll concentrations

in corals during and after a bleaching event. Mar Ecol Prog Ser 158:51–59.

Jones RJ (1997b) Zooxanthellae loss as a bioassay for assessing stress in corals. Mar

Ecol Prog Ser 149:163–171.

Jones AM, Berkelmans R, van Oppen MJ, Mieog JC, Sinclair W (2008) A community

change in the algal endosymbionts of a scleractinian coral following a natural

bleaching event: field evidence of acclimatization. Proc R Soc B 275:1359-1365.

Jones RJ, Yellowlees D (1997) Regulation and control of intracellular algae (=

zooxanthellae) in hard corals. Phil Trans R Soc B 352:457-468.

Kautz R, Stys B, Kawula R (2007) Florida vegetation and land use change between

1985-1989 and 2003. Florida Scientist 70:12-23.

Katoh K, Misawa K, Kuma K, Miyata T (2002) MAFFT: a novel method for rapid

multiple sequence alignment based on fast Fourier transform. Nucleic Acids

Res. 30:3059-3066.

Kemp DW, Hernandez-Pech X, Iglesias-Prieto R, Fitt WK, Schmidt GW (2014)

Community dynamics and physiology of Symbiodinium spp. before, during, and

after a coral bleaching event. Limnol. Oceanogr 59:788-797.

Kenkel CD, Goodbody-Gringley G, Caillaud D, Davies SW, Bartels E, Matz MV (2013)

Evidence for a host role in thermotolerance divergence between populations of

the mustard hill coral (Porites astreoides) from different reef environments.

Molec Ecol 22: 4335–4348.

LaJeunesse TC, Trench RK (2000) Biogeography of two species of Symbiodinium

(Freudenthal) inhabiting the intertidal sea anemone Anthopleura elegantissima

(Brandt). Biol Bull 199:126–134.

LaJeunesse TC (2002) Diversity and community structure of symbiotic

dinoflagellates from Caribbean coral reefs. Mar Biol 141:387–400.

LaJeunesse TC, Bhaaooli R, Hidaka M, DeVantier L, Done T, Schmidt GW, Fitt WKW,

Hoegh-Guldberg O (2004) Close related Symbiodinium spp. differ in relative

dominance in coral reef host communities across environmental, latitudinal

and biogeographic gradients. Mar Ecol Prog Ser 284:147-161.

LaJeunesse TC (2005) “Species” radiations of symbiotic dinoflagellates in the

Atlantic and Indo-Pacific since the Miocene-Pliocene transition. Mol Biol Evol

22:570-581.

LaJeunesse TC, Smith R, Finney J, Oxenford H (2009) Outbreak and persistence of

opportunistic symbiotic dinoflagellates during the 2005 Caribbean mass coral

“bleaching” event. Proc R Soc B 276:4139–4148.

LaJeunesse TC, Pettay DT, Sampayo EM, Phongsuwan N, Brown BE, Obura DO, Fitt

WK (2010) Long-standing environmental conditions, geographic isolation and

host-symbiont specificity influence the relative ecological dominance and

genetic diversification of coral endosymbionts in the genus Symbiodinium. Jour

Biogeogr 37:785–800.

LaJeunesse TC, Smith R, Walther M, Pinzón J, Pettay DT, McGinley M, Warner ME

(2010) Host-symbiont recombination versus natural selection in the response

of coral-dinoflagellate symbioses to environmental disturbance. Proc R Soc B

277:2925–2934.

LaJeunesse TC, Thornhill DJ (2011) Improved resolution of reef-coral endosymbiont

(Symbiodinium) species diversity, ecology, and evolution through psbA non-

coding region genotyping. PloS one 6, e29013:1-11.

Lapointe BE (1997) Comparative ecology of harmful macroalgal blooms in south

Florida’s coastal waters (CEHAB). Final Report Center for Coastal Research,

Harbor Branch Oceanographic Institution.

Lapointe BE, Barile PJ, Matzie WR (2004) Anthropogenic nutrient enrichment of

seagrass and coral reef communities in the Lower Florida Keys: discrimination

of local versus regional nitrogen sources. Jour Exp Mar Biol Ecol 308:23–58.

Lapointe BE (2007) Comparative Ecology of Harmful Macroalgal Blooms in South

Florida’s Coastal Waters (CEHAB). Final Report: Center for Coastal Research,

Harbor Branch Oceanographic Institute 1-63.

Lapointe BE, Herren LW, Bedford BJ (2012) Effects of hurricanes, land use, and

water management on nutrient and microbial pollution: St. Lucie Estuary,

Southeast Florida. Jour Coast Res 28:1345–1361.

Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H,

Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG

(2007) Clustal W and Clustal X version 2.0. Bioinformatics 23:2947-2948.

Lesser MP, Shick JM (1989) Effects of irradiance and ultraviolet radiation on

photoadaptation in the zooxanthellae of Aiptasia pallida: primary production,

photoinhibition, and enzymic defenses against oxygen toxicity. Mar Biol

102:243-255.

Lesser MP, Mazel C, Phinney D, Yentsch CS (2000) Light absorption and utilization

by colonies of the congeneric hermatypic corals Montastraea faveolata and

Montastraea cavernosa. Limnol Oceanogr 45:76–86.

Litaker RW, Vandersea MW, Kibler SR, Reece KS, Stokes NA, Lutzoni FM, Yonish BA,

West MA, Black MND, Tester PA (2007) Recognizing dinoflagellate species

using ITS rDNA sequences. Jour Phycol 43:344–355.

Loram JE, Boonham N, O’Toole P, Trapido-Rosenthal HG, Douglas AE (2007)

Molecular quantification of symbiotic dinoflagellate algae of the genus

Symbiodinium. Biol Bull 212:259–68.

Marsh JA (1970) Primary productivity of reef-building calcareous red algae. Ecology

51:255-263.

Mayfield AB, Gates RD (2007) Osmoregulation in anthozoan–dinoflagellate

symbiosis. Comp Biochem Phys A 147:1–10.

McGinley M, Aschaffenburg M, Pettay DT, Smith R, LaJeunesse TC, Warner ME

(2012) Symbiodinium spp. in colonies of eastern Pacific Pocillopora spp. are

highly stable despite the prevalence of low-abundance background populations.

Mar Ecol Prog Ser 462:1–7.

McGuire MP, Szmant AM (1997) Time course of physiological responses to NH4

enrichment by a coral-zooxanthellae symbiosis. Proc 8th Int Coral Reef Sym

1:909-914.

Mieog, J. C., van Oppen, M. J. H., Cantin, N. E., Stam, W. T., and Olsen, J. L. (2007). Real-

time PCR reveals a high incidence of Symbiodinium clade D at low levels in four

scleractinian corals across the Great Barrier Reef: implications for symbiont

shuffling. Coral Reefs 26:449–457.

Mieog JC (2009) Flexibility of the coral-algal symbiosis in the face of climate change:

investigating the adaptive bleaching hypothesis. Center for Ecological and

Evolutionary Studies, University of Groningen, The Netherlands. 1-193.

Mieog JC, van Oppen MJH, Berkelmans R, Stam WT, Olsen JL (2009) Quantification of

algal endosymbionts (Symbiodinium) in coral tissue using real-time PCR. Molec

Ecol Res 9:74–82.

Muller-Parker G, D’Elia CF (1997) Interactions between corals and their symbiotic

algae. Life and death of coral reefs 96-113.

Mumby PJ, Steneck RS (2008) Coral reef management and conservation in light of

rapidly evolving ecological paradigms. TREE 23:555–563.

Muscatine L, Porter JW (1977) Reef corals: mutualistic symbioses adapted to

nutrient-poor environments. Bioscience 27:454-460.

Muscatine L, Falkowski PG, Dubinsky Z, Cook PA, McCloskey LR (1989) The effect of

external nutrient resources on the population dynamics of zooxanthellae in a

reef coral. Proc R Soc B 236:311-324.

NOAA Coral Reef Conservation Program (2009) NOAA Coral Reef Conservation

Program Goals and Objectives 2010-2015. Silver Spring, MD: NOAA.

Oksanen J, Blanchet FG, Kindt R, Oksanen MJ, Suggests MASS (2013) Package ‘vegan’.

Community ecology package Version 2:0-0.

Pochon X, Pawlowski J, Zaninetti L, Rowan R (2001) High genetic diversity and

relative specificity among Symbiodinium-like endosymbiotic dinoflagellates in

soritid foraminiferans. Mar Biol 139: 1069-1078.

Pochon X, Gates, RD (2010) A new Symbiodinium clade (Dinophyceae) from soritid

foraminifera in Hawai’i. Molec Phylo Evol 56:492-497

Pochon X, Putnam HM, Burki F, Gates RD (2012) Identifying and characterizing

alternative molecular markers for the symbiotic and free-living dinoflagellate

genus Symbiodinium. PloS one 7, e29816:1-13.

Porter JW (1976) Autotrophy, heterotrophy, and resource partitioning in Caribbean

reef-building corals. Am Nat 110:731–742.

Putnam HM, Stat M, Pochon X, Gates RD (2012) Endosymbiotic flexibility associates

with environmental sensitivity in scleractinian corals. Proc R Soc B 279:4352–

4361.

Quigley KM, Davies SW, Kenkel CD, Willis BL, Matz MV, Bay LK (2014) Deep-

sequencing method for quantifying background abundances of Symbiodinium

types: exploring the rare Symbiodinium biosphere in reef-building corals. PloS

one 9, e94297.

Reed J (1982) Letter to Dr. Harold Wanless. Univ of Miami RSMAS.

Rodriguez-Lanetty M, Loh W, Carter D, Hoegh-Guldberg O (2001) Latitudinal

variability in symbiont specificity within the widespread scleractinian coral

Plesiastrea versipora. Mar Biol 138:1175-1181.

Rodriguez-Lanetty M, Chang SJ, Song JI (2003) Specificity of two temperate

dinoflagellate-anthozoan associations from the north-western Pacific Ocean.

Mar Biol 143:1193–1199.

Rowan R, Knowlton N (1995) Intraspecific diversity and ecological zonation in

coral-algal symbiosis. PNAS 92:2850-2853.

Rowan R, Knowlton N, Baker A, Jara J (1997) Landscape ecology of algal symbionts

creates variation in episodes of coral bleaching. Nature 388:265-269.

Sampayo EM, Franceschinis L, Hoegh-Guldberg O, Dove S (2007) Niche partitioning

of closely related symbiotic dinoflagellates. Molec Ecol 16:3721-3733.

Sampayo EM, Dove S, LaJeunesse TC (2009) Cohesive molecular genetic data

delineate species diversity in the dinoflagellate genus Symbiodinium. Molec Ecol

18:500–519.

Savage AM, Goodson MS, Visram S, Wiedenmann J, Douglas AE (2002) Molecular

diversity of symbiotic algae at the latitudinal margins of their distribution:

dinoflagellates of the genus Symbiodinium in corals and sea anemones. Mar Ecol

Prog Ser 244:17–26.

Silverstein RN, Correa AM, Baker AC (2012) Specificity is rarely absolute in coral–

algal symbiosis: implications for coral response to climate change. Proc R Soc B

279:2609-2618.

Stat M, Bird CE, Pochon X, Chasqui L, Chauka LJ, Concepcion GT, Gates RD (2011)

Variation in Symbiodinium ITS2 sequence assemblages among coral colonies.

PloS one 6, e15854:1-13.

Szmant AM (1986) Reproductive ecology of Caribbean reef corals. Coral reefs 5:43-

53.

Szmant AM (2002) Nutrient enrichment on coral reefs: is it a major cause of coral

reef decline? Estuaries 25:743-766.

Szmant AM (2012) Nutrient enrichment on coral reefs: is it a major cause of coral

reef decline? Estuaries 25:743-766.

Thornhill DJ, LaJeunesse TC, Kemp DW, Fitt WK, Schmidt GW (2005) Multi-year,

seasonal genotypic surveys of coral-algal symbioses reveal prevalent stability

or post-bleaching reversion. Mar Biol 148:711–722.

Thornhill DJ, Fitt WK, Schmidt GW (2006) Highly stable symbioses among western

Atlantic brooding corals. Coral Reefs 25:515–519.

Thornhill DJ, LaJeunesse TC, Santos SR (2007) Measuring rDNA diversity in

eukaryotic microbial systems: how intragenomic variation, pseudogenes, and

PCR artifacts confound biodiversity estimates. Molec Ecol 16:5326–5340.

Thornhill DJ, Kemp DW, Bruns BU, Fitt WK, Schmidt GW (2008) Correspondence

between cold tolerance and temperate biogeography in a western Atlantic

Symbiodinium (Dinophyta) lineage. Jour Phycol 44:1126-1135.

Thornhill DJ, Lewis AM, Wham DC, LaJeunesse TC (2014) Host-specialist lineages

dominate the adaptive radiation of reef coral endosymbionts. Evolution,

68:352-367.

Titlyanov EA, Titlyanova TV, Leletkin VA, Tsukahara J, Van Woesik R, Yamazato K

(1996) Digestion and regulation of zooxanthellae density in hermatypic corals.

Mar Ecol Prog Ser 134:167-178.

Toller WW, Rowan R, Knowlton N (2001) Zooxanthellae of the Montastraea

annularis species complex: patterns of distribution of four taxa of Symbiodinium

on different reefs and across depths. Biol Bull 201:348-359.

Tremblay P, Grover R, Maguer JF, Hoogenboom M, Ferrier-Pagès C (2014) Carbon

translocation from symbiont to host depends on irradiance and food

availability in the tropical coral Stylophora pistillata. Coral Reefs 33:1-13.

Trench RK (1993) Microalgal-invertebrate symbioses: a review. Endocytobiosis Cell

Res 9:135-175.

Ulstrup KE, van Oppen MJH (2003) Geographic and habitat partitioning of

genetically distinct zooxanthellae (Symbiodinium) in Acropora corals on the

Great Barrier Reef. Molec Ecol 12:3477–3484. van Oppen MJH, Palstra FP, Piquet AMT, Miller DJ (2001) Patterns of coral–

dinoflagellate associations in Acropora: significance of local availability and

physiology of Symbiodinium strains and host–symbiont selectivity. Proc R Soc

B 268: 1759-1767.

Vargas-Ángel B, Peters EC, Kramarsky-Winter E, Gilliam DS, Dodge RE (2007)

Cellular reactions to sedimentation and temperature stress in the Caribbean

coral Montastraea cavernosa. Jour Invert Path 95:140-145.

Warner ME, Fitt WK, Schmidt GW (1996) The effects of elevated temperature on the

photosynthetic efficiency of zooxanthellae in hospite from four different species

of reef coral: a novel approach. Plant Cell Enviro 19:291-299.

Warner ME, Fitt WK, Schmidt GW (1999) Damage to photosystem II in symbiotic

dinoflagellates: a determinant of coral bleaching. PNAS 96:8007-8012.

Warner ME, Chilcoat GC, McFarland FK, Fitt WK (2002) Seasonal fluctuations in the

photosynthetic capacity of photosystem II in symbiotic dinoflagellates in the

Caribbean reef-building coral Montastraea. Mar Biol 141:31–38.

Wilkerson FP, Kobayashi D, Muscatine L (1988) Mitotic index and size of symbiotic

algae in Caribbean Reef corals. Coral Reefs 7:29–36.

Wooldridge SA, Done TJ (2009) Improved water quality can ameliorate effects of

climate change on corals. Ecol Appl 19:1492–1499.

Wooldridge S (2013) Breakdown of the coral-algae symbiosis: towards formalising a

linkage between warm-water bleaching thresholds and the growth rate of the

intracellular zooxanthellae. Biogeosciences 10:1647–1658.