California State University, Northridge an Ecological

Home , Ajax Reef, Davis Reef, Eastern Dry Rocks, French Reef, Gorgoniidae, Hermatypic coral

CALIFORNIA STATE UNIVERSITY, NORTHRIDGE

AN ECOLOGICAL AND PHYSIOLOGICAL ASSESSMENT OF TROPICAL CORAL

REEF RESPONSES TO PAST AND PROJECTED DISTURBANCES

A thesis submitted in partial fulfillment of the requirements

for the degree of Master of Science in Biology

Elizabeth Ann Lenz

May 2014

The thesis of Elizabeth A. Lenz is approved by:

Robert C. Carpenter, Ph.D. Date:

Eric D. Sanford, Ph.D. Date:

Mark A. Steele, Ph.D. Date:

Peter J. Edmunds, Ph.D., Chair Date:

California State University, Northridge

ACKNOWLEDGEMENTS

I would like to thank Dr. Peter J. Edmunds first and foremost for being my fearless leader and advisor - for the incredible opportunities and invaluable mentorship he has provided to me as a graduate student in the Polyp Lab. I am ever so grateful for his guidance, endless caffeinated energy, constructive critiques, and dry British humor.

I would also like to thank my loyal committee members Drs. Robert Carpenter and Mark Steele at CSUN for their availability and expert advise during this process. Their suggestions have greatly contributed to my thesis. I would not only like to acknowledge Dr. Eric Sanford from UC Davis for serving on my committee, but thank him for his incessant support throughout my career over the last 7 years. I will always admire his contagious enthusiasm for invertebrates, passion for scientific research, and unlimited knowledge about marine ecology.

My research would not have been possible without the technical support and assistance from my colleagues in Moorea, French Polynesia and St. John, USVI. I am grateful to Dr. Lorenzo Bramanti, Dr. Steeve Comeau, Vince Moriarty, Nate Spindel, Emily Rivest, Christopher Wall, Darren Brown, Alexandre Yarid, Nicolas Evensen, Craig Didden, the VIERS staff, and undergraduate assistants: Kristin Privitera-Johnson and Amanda Arnold.

I would like to thank the former and current members of the CSUN Biology Department. Particularly, I am thankful for the MBGSA community, WiS group, Hollie Putnam, Anya Brown, Lianne Jacobson, Stella Swanson, Heather Hillard, Amy Briggs, Camdilla Wirth, Lauren Valentino, Mia Adreani, Justin Hackitt, and of course my fellow female scholars in the Polyp Lab: Sylvia Zamudio, Jennifer Smolenski, and Ananda Ellis.

I would like to thank my constant sources of encouragement - Dr. Thomas Lenz and Debbra Lenz for their unconditional love and patience, my five siblings especially Mary Lenz for her generous time and reliability, and for my dear friends for enhancing my graduate experience: Sarah McVay, Emily Tung, Lauren Burgunder, Megan Boyd, Theresa Dineen, Malia Collins, Melanie Levy, Jackie Sones, DSO Jason Heron, Kristin Aquilino, Christopher Kwan, Bart Bagdasaryan, Casey Scott, Hamilton W. Fennie, Alexander G. Snyder, Kerry Nichols, Evelyne Kuo, Brian Lee, Kirk Sato, Tren Kauzer, Paul Logston, Maggie Sogin, Sean McCann, and Lauren Mickool.

Finally, I am grateful to Drs. Annaliese Hettinger and John J. Stachowicz for recommending I pursue my master’s degree at CSUN in the Polyp Lab.

My graduate thesis was made possibly by the financial support of CSUN’s College of Science and Mathematics Graduate Research Promise Fellowship, the National Science Foundation research support for Moorea Coral Reef Long-Term Ecological Research (OCE 10-26852), Division of Ocean Science (OCE 10-41270), St. John USVI Long- Term Research in Environmental Biology (DEB 03-43570, DEB 08-41441, and OCE 13- 32915), and the Gordon and Betty Moore Foundation.

iii

Signature Page ii

Acknowledgements iii

Abstract v

Chapter 1 General Introduction 1

Chapter 2 Decadal-scale changes in the density of Caribbean gorgonians Introduction 14 Methods 17 Results 22 Discussion 27 Tables 36 Figures 48

Chapter 3 Contrasting morphologies of the Pacific coral Porites rus are not affected by ocean acidification Introduction 56 Methods 60 Results 70 Discussion 75 Tables 83 Figures 96

Chapter 4 Concluding remarks 105

Literature Cited 109

Abstract

An Ecological and Physiological Assessment of

Tropical Coral Reef Responses to Past and Projected Disturbances

Elizabeth A. Lenz

Masters of Science in Biology

Tropical reef structure and topographical complexity is a product of coral morphology, which greatly enhances biodiversity, ecological function, and ecosystem services. However, reef-building corals have declined in cover by 50-80% in response to the accumulated effects of natural and anthropogenic disturbances over the last 3 to 4 decades. This thesis is comprised of two different studies that address potential changes in shallow reef communities and benthic structure in response to natural and anthropogenic disturbances. The first objective was to examine changes in Caribbean benthic communities as scleractinians have declined and the second objective was to determine differential responses of contrasting morphotypes of corals to acidified conditions in Moorea, French Polynesia. In Chapter 2, I assessed the abundance of arborescent gorgonians (AGs) at local and regional scales to test the hypothesis that AGs have increased in abundance on Caribbean reefs over multiple decades. In St. John, mean abundance of AGs increased 42% from 1992-2012, with each of the dominant genera

(Gorgonia, Antillogorgia, and pooled genera: Sea Rods) increasing 11-221% over the same period. Regionally, the compiled data show that AGs have increased in abundance, with mean densities rising from 7.0 to 15.1 colonies m-2 over the last 45 years. This study highlights the apparent success and importance of AGs on contemporary Caribbean

reefs, where scleractinian percent cover has remained at ~10%. In Chapter 3, I evaluated the calcification rates of branches and plates of Porites rus in response to ocean acidification (OA) (~800 µatm and 1000 µatm PCO2) with temperature, light and water flow. Skeletal morphology of reef-building hermatypic corals is influenced by abiotic factors, but calcification of coral skeleton is expected to decline as ocean acidification

(OA) increases. Theory predicts that hermatypic corals with contrasting morphologies will differ in calcification rates due to differences in: 1) light absorption by Symbiodinium spp. for photosynthesis to enhance coral calcification, and 2) differential mass transfer characteristics (e.g., modulating the flux of dissolved inorganic carbon and hydrogen), which may mitigate the negative effects on calcification in high PCO2. In the second study, I determined that branches and plates were both tolerant of elevated temperature and PCO2. However, branches had higher variability in calcification rates than plates in response to light, and flow. Together these results of this research demonstrate two successful taxa (gorgonians and a weedy scleractinian) in tropical reef communities that have relatively high tolerances against natural and anthropogenic disturbances.

Although, gorgonians and P. rus may be alternatives to formerly dominant, slow growing, and diverse communities of hermatypic corals, the functional roles and the ecosystem services they provide will likely differ.

vi Chapter 1

General Introduction

Disturbances on Tropical Coral Reefs

Tropical coral reefs facilitate the highest biodiversity of any ecosystems in the world (Connell 1978) and provide economic value through coastal protection, fisheries, and ecotourism (Moberg and Folke 1999; Cesar et al. 2003). Hermatypic corals (P:

Cnidaria, C: Anthozoa, SC: Hexacorallia, O: Scleractinia) are the ecosystem engineers responsible for the topographically complex framework that many organisms rely on for food and shelter (Idjadi and Edmunds 2006; Wild et al. 2011). The skeletal framework of scleractinians is created from the deposition of calcium carbonate (CaCO3) in the form of aragonite. However, natural and anthropogenic stressors are dramatically changing corals, from the molecular (e.g., gene regulation) to ecological scale (e.g., community structure) (Nyström et al. 2000; Bellwood et al. 2004; Hoegh-Guldberg et al. 2007).

Tropical hermatypic coral communities are restricted geographically and constantly exposed to multiple stressors acting on reef systems. These stressors include both pulse

(e.g., hurricanes, disease outbreaks, and thermal bleaching events) and press (e.g., ocean acidification and increased sea surface temperature [SST]) disturbances (Connell 1997).

These assaults can have several grave consequences, with the effects on calcification and maintenance of reef complexity (Kleypas et al. 1999; Langdon and Atkinson 2005;

Kleypas et al. 2001) at the forefront of concerns over the future of tropical coral reef communities (Knowlton 2001; Hough-Guldberg et al. 2007). As anthropogenic and natural disturbances continue, it is crucial to understand the physiological and ecological

responses of tropical corals to projected environmental conditions in order to determine reef resilience and shifts in coral reef communities.

Changes in Coral Reef Communities

Field surveys, photoquadrats, video recordings, and aerial photographs have been common tools used for long-term monitoring of scleractinian communities with percent cover the most widely used metric of coral abundance (Gardner et al. 2003; De’ath et al.

2013; Edmunds 2013; Ruzicka et al. 2013). In the last 3-5 decades, monitoring efforts have determined coral loss to be ~50% on the Great Barrier Reef from 1965-2012

(De’ath et al. 2013), ~80% throughout the Caribbean from 1977 to 2001 (Gardner et al.

2003), and ~22% in the Florida Keys from 1999-2009 (Ruzicka et al. 2013). In the

Caribbean, the massive decline in scleractinian cover has been attributed to local disturbances such as die-off of the sea urchin Diadema antillarum (Lessios et al. 1984), white and yellow band disease outbreaks (Aronson and Precht 2001; Schutte et al. 2010), hurricanes (Woodley et al. 1981), and global drivers such as El Niño Southern Oscillation

(ENSO) events that cause mass coral bleaching (Glynn 1993; Lesser 2011). Over the decades, monitoring of shallow (≤ 25 m) coral reef communities has focused on ‘phase shifts’ from a “coral-dominated” to a “macroalgae-dominated” state (Hughes 1994).

Macroalgae can successfully outcompete and hinder the success (e.g., recruitment and growth) of scleractinians when herbivores are removed from the reef by overfishing

(Jackson et al. 2001) or Diadema die off (Lessios et al. 1984). Recently, a few studies have included other benthic taxa beyond macroalgae and scleractinians, for example corallimorpharians, sponges, and gorgonians (Diaz and Rützler 2001; Norström et al.

2009; Ruzicka et al. 2013). Accounting for other benthic organisms in monitoring will provide a more accurate description of shifts occurring in tropical reef communities and alterations in ecosystem services as disturbances persist and intensify (Diaz and Rützler

2001; Inoue et al. 2013; Rossi 2013; Ruzicka et al. 2013).

Gorgonians (P: Cnidaria, C: Anthozoa, SC: Octocorallia, O: Alcyonacea, SO:

Holaxonia, F: Gorgoniidae) have been long reported to be common on shallow

Caribbean reefs (Kinzie 1973; Lasker 1995), yet they have received little attention in most monitoring efforts (Norstöm et al. 2009; Ruzicka et al. 2013), with studies of their ecology uncommon when compared to the efforts taken to understand the biology and ecology of scleractinians (Gardner et al. 2003; Schutte et al. 2010; Edmunds 2013).

Despite the limited information, gorgonians have a suite of traits that may contribute to their success under multiple disturbances such as hurricanes (Witman 1992; Lugo et al.

2000), increased SST (Lasker 1995), competition with macroalgae (Pawlik et al. 1987;

Yoshioka and Yoshioka 1989), and disease (Kim et al. 2000; Bruno et al. 2003), which have contributed to the decline of scleractinian cover in the Caribbean (Gardner et al.

2003; Schutte et al. 2010).

Gorgonians have the potential to tolerate the stressors scleractinians are weakened by, such as wave stress generated by hurricanes (Koehl 1984), thermal bleaching (Lasker

1995), and competition with macroalgae (Pawlik et al. 1987). The flexible and soft bodies of gorgonians have spicules, CaCO3 structures in the form of magnesium-calcite, embedded in the mesoglea where high regulation of skeletogenesis occurs within the

gorgonian tissue (Chave 1954; Bond et al. 2005). Spicules are tightly packaged and closely arranged in low flow environments whereas in high wave energy conditions, the spicules are loosely arranged to allow for higher flexibility (West 1997, 1998). This pliability alleviates the impacts of strong currents and reduces high drag forces acting on gorgonians (Koehl 1996). As the intensity and frequency of hurricanes are expected to increase in the Caribbean (Gardner et al. 2005), the capacity for gorgonians to withstand such forces is potentially greater compared to some scleractinian species with delicate, thin branches such as the “weedy” (sensu Knowlton) corals, Poritidae and Agaricidae

(Hughes 1994).

Although less extensively studied relative to scleractians, gorgonians and their association with Symbiodinium spp., the unicellular dinoflagellates embedded in the gastroder, are less negatively impacted by thermal stress (Goulet and Coffroth 1997;

Lasker 2003; Kirk et al. 2005; Lasker 2005). Hermatypic corals can be sensitive to thermal stress, resulting in mass coral bleaching and death (Glynn 1993; Brown 1997).

Symbiodinium spp., which translocate photosynthetic carbon to meet the metabolic demands of the coral host (Muscatine et al. 1984), becomes impaired and are expelled during high SST or ENSO events (Gates et al. 1992; Jones et al. 1998). When bleaching occurs, this tightly coupled relationship dissociates and the source of ≥ 90% of organic carbon for corals is lost (Muscatine et al. 1984). During past high SST events in the

Caribbean, the Symbiodinium spp. in gorgonians, such as Plexaura kuna (Lasker, Kim, and Coffroth 1996) and Gorgonia ventalina (Linnaeus 1758), were unaffected with minor variability in bleaching responses within species. Gorgonian populations appear to have

higher resistance to avoid the bleaching responses to high temperatures often observed in scleractinians (Goulet and Coffroth 1997; Lasker 2003; Kirk et al. 2005). For example, in the San Blas Islands, Panama, densities of Symbiodinium spp. in Plexaura kuna were similar before and after the 1983 ENSO event with higher resistance to bleaching relative to scleractinians in the same area (Lasker 2003). As severe disturbances, such as bleaching and hurricanes, continue in the Caribbean, gorgonian populations may have higher resilience and success compared to scleractinians (Ruzicka et al. 2013).

While disturbances and recovery processes cause changes in tropical reef community structure (Connell 1997; Hughes 1994; Norström et al. 2009), it is important to thoroughly consider the compositional changes of benthic taxa that generate “animal forests”. The “animal forest” is a recently coined ecological term, describing three- dimensional canopies comprised of living and actively feeding fauna such as scleractinians, cnidarians, bryozoans, ascidians, and sponges (Rossi 2012). Descriptive studies of the abundances, biomass, and biodiversity of alternative benthic taxa are needed to assess the interactive effects of anthropogenic and natural disturbances altering coral reef ecosystems (Rossi 2013). In Chapter 2, I describe changes in shallow tropical reef communities by focusing on gorgonian abundances over 2-4 decades. I studied changes in the abundance (colonies m-2) of gorgonians at the local-scale scale (≤ 20 km

[Mittelbach et al. 2001]) in St. John, U.S. Virgin Islands, using archived photoquadrats from 1992 to 2012, and at the regional-scale (i.e., 200-4000 km [Mittelbach et al. 2001]) throughout the Caribbean using data in peer-reviewed articles and unpublished monitoring reports from 1968 to 2013. The purpose of this descriptive study on

Caribbean gorgonians was to test for changes and success of other benthic taxa beyond scleractinians and macroalgae.

Future Disturbances on Tropical Coral Reefs

Ocean Acidification

The accumulation of carbon emissions from increased human use of fossil fuels since the Industrial Revolution has led to increases in the concentration of atmospheric carbon dioxide (CO2) at unparalleled rates (Hönisch et al. 2012; Zeebe et al. 2012). Excess

CO2 dissolving into the oceans alters the carbonate chemistry equilibrium in seawater (Orr eet al. 2005; Doney et al. 2009; Feely et al. 2009). As CO2 dissolves in seawater, carbonic

+ acid (H2CO3) is produced and releases a hydrogen ion (H ), causing a decline ocean pH. In

2- acidic conditions, carbonate ions (CO3 ) from the water column or dissociated from

+ - calcium carbonate (CaCO3) binds to H to form HCO3 and causes saturation states (Ω) of

2- CaCO3 (aragonite, calcite, and magnesium-calcite) to decline. CO3 is essential for the deposition of CaCO3, which produces the skeleton and shells of calcifying marine organisms like corals, sea urchins, and oysters (Orr et al. 2005). The altered carbonate seawater chemistry caused by the absorption of excess atmospheric CO2 that results in the reduction in pH and available carbonate ions with no change in total alkalinity is known as ocean acidification (OA).

As OA is expected to intensify with increases in CO2 emissions, there is a need to understand the biological impacts of OA on calcifying marine taxa in order to identify weak and resistant responses. The objective of these studies is to determine the capacities

for acclimatization and the potential for adaptation to predict the ecological implications for marine organisms exposed to OA. The majority of studies have used calcification rates as the primary parameter in testing for effects of OA (Ries et al. 2009; Kroeker et al. 2010,

2013). From the exponentially growing OA literature on calcification rates, meta-analyses have concluded that OA will have overall detrimental effects on calcifying marine organisms such as echinoderms and cnidarians (Kroeker et al. 2010; 2013). In elevated

PCO2 conditions, calcifying marine organisms experience altered molecular and physiological, such as changes in gene expression and respiration, respectively, which may help sustain biological functions such as calcification and reproduction (Hofmann and

Kelly 2012). Despite negative impacts of OA, variation in calcification responses to more acidified conditions has been detected within and among taxa using experiments conducted in situ (Kline et al. 2012; Wall and Edmunds 2013) and in the laboratory (Comeau et al.

2013). This variability may reflect differences in coping mechanisms and strategies that either enhance or hinder the capacity for marine calcifiers to acclimatize (e.g., metabolic adjustments and gene regulation) (Gates and Edmunds 1999; Hofmann et al. 2010; Pespeni et al. 2013) and their potential for adaptation (Melzner et al. 2009; Kroeker et al. 2010;

Kroeker et al. 2013). Furthermore, there are marine systems with natural pH fluctuations, such as along high upwelling regions (Feely et al., 2009; Hofmann et al. 2011; Pespeni et al. 2012; Shamberger et al., 2014) and CO2 vents (Kroeker et al. 2011; Inoue et al. 2013).

In such acidified conditions, there has been strong selection for calcifying organisms to adapt (Kelly and Hofmann 2013; Reusch 2014). However, little is known about the genetics, physiology, phenotypic traits, and ecological interactions that promote resilient populations (Sunday et al. 2014).

Tropical hermatypic corals have received enormous attention in climate change and

OA studies because the complex reef structures the produce support diverse communities, which may be threatened under acidified conditions (Hoegh-Guldberg et al. 2007).

Tropical scleractinians are particularly vulnerable to OA because they are geographically restricted to regions with oligotrophic and high Ωarag waters (Veron 1986). Favorable conditions for coral calcification are becoming more restricted with warming and acidity increasing (Atkinson and Cuet 2008). In attempts to predict changes in future species composition of coral reef communities and structure, some OA research on scleractinians has focused on categorizing coral species and functional groups as “winners” and “losers”

(sensu Loya et al. 2001). This categorization is designed to identify the biological traits and reveal molecular processes that enable scleractinians to persist under stressful conditions (Anthony et al. 2008, Edmunds et al. 2012; Comeau et al. 2013; Shamberger et al. 2014). Moreover, studies of calcification responses of corals exposed to acidified conditions provide evidence for the next phase of OA research to understand mechanisms contributing to reef resiliency through either increased capacity for acclimatization (e.g. plasticity) or adaptation to local and global stressors (Pespeni et al. 2013; Munday et al.

2013; Reusch 2014). This understanding can then be applied to conservation efforts to preserve hardy corals and mitigate local pressures that drive corals to the edge of their physiological thresholds.

Scleractinian Morphology and Phenotypic Plasticity

Morphology in sessile, modular organisms has been described extensively as a structural product shaped by biological interactions and the physical environment, such as

from competition and light, respectively (Jackson 1979). Scleractinians provide good examples of how morphological diversity within and among species, such as branches, plates, massive, and encrusting, which can be used to overcome resource-limitations such as limited nutrients and space in tropical coral reefs (Jackson 1979; Chappel 1980).

Environmental parameters influencing coral morphologies within and among species include light (Jaubert 1977), water flow (Lesser 1994), sedimentation (Padillo-Gamiño et al. 2012), predation (Jackson 1979), light (Muko et al. 2000), and competition (Jackson

1979). As skeletal morphology is a prevalent trait of coral reef communities, it is relevant to reexamine how the environment and predicted abiotic stressors (OA and climate change) will impact the physiological performance of contrasting morphological groups.

Previous OA studies have compared morphologies at the genus-level to identify

“winners” and “losers”, but they have neglected to address and eliminate the confounding factors of differences in genetics, composition in Symbiodinium spp., and physiology across taxa. OA studies should conduct morphological comparisons within coral species to determine sensitivity related to “morpho-functional” groups such as massive, plating, or encrusting corals to reduce inherent biological differences (Jackson 1979; Patterson

1992; Todd et al. 2008). Comparing morphologies within species is an initial step towards examining the response of a phenotypically plastic coral to OA conditions.

Physiological plasticity remains of interest for studying coral acclimatization strategies to environmental conditions, as it allows for wider geographical distribution and may be a

short-term strategy for corals to acclimatize to OA and climate change conditions

(Hofmann et al. 2010).

Light

In oligotrophic waters, light is the most essential resource for hermatypic corals containing photosynthetic symbiotic dinoflagellates (Symbiodinium spp.) within their gastrodermal tissue. In shallow waters, these symbionts can provide ≥ 90% of the carbon required to meet the metabolic demands of the coral host (Muscatine Muscatine et al. 1984) and are responsible for increased rates of CaCO3 deposition through light-enhanced calcification (LEC) (Pearse and Muscatine 1971). As light intensity and quality change with depth, hermatypic corals exhibit changes in corallum morphology with depth within and among species (Jaubert 1977; Vermeij and Bak 2002). In shallow waters and high light conditions, branching is a common morphology that can scatter high light intensity to reduce photodamage from excess light energy acting on Symbiodinium spp. (Enriquez et al.

2005). In deeper waters, corals can exhibit more plated and encrusting morphologies to acquire attenuated light (Jaubert 1977; Muko et al. 2002).

Skeletal morphology and calcification rates of hermatypic corals are strongly influenced by light intensity (Jaubert 1977; Muko et al. 2000; Enriquez et al. 2005). Both features should be considered in OA and climate change research to determine ecologically relevant responses of coral holobionts. Previous studies have shown scleractinians to be negatively effected by OA (Kroeker et al. 2010). The experimental designs in these studies (e.g., Edmunds et al. 2012), however, involved corals incubated

under low light conditions measured in the field may reduce the capacity for photosynthesis of Symbiodinium spp. and calcification. Recently, studies of OA have started to investigate the interactive effects of elevated PCO2 and light intensities that reflect the natural environment (Edmunds et al. 2012; Dufault et al. 2013; Comeau et al.

2014). Dufault et al. (2013) concluded that the response to PCO2 is light-dependent because calcification rates of scleractinian recruits varied hyperbolically with light intensity in response to elevated PCO2. A species-level comparative study demonstrated that high light intensity fully mitigated the negative effects of OA on the calcification rates for some corals, but not for others (Suggett 2013). Further research is needed to understand the calcification response of corals to OA under a range of light intensities because few studies recognize the important biological role of Symbiodinium spp. in the context of light-enhanced calcification.

Water Flow

Water flow is another environmental parameter that can strongly influence corallum shape and size in scleractinians (Bruno and Edmunds 1998; Lesser et al. 1994). In high flow environments (≥ 25 cm s-1), branching scleractinians are typically thicker while in low flow conditions (5 cm s-1) branches are thinner and more delicate (Lesser et al. 1994;

Kaandorp 1999). As sessile organisms, many aspects of the physiological performance of scleractinians can be improved by flow rates depending on the shape of the organism

(Patterson and Sebens 1989; Patterson 1992; Nakamura et al. 2005; Goldenheim and

Edmunds 2011). Scleractinians rely on the delivery of dissolved solutes such as metabolites and nutrients for active or passive transport across boundary layers into their

tissues (Patterson 1992; Lesser et al. 1994). In response to varying flow rates, many scleractinians have the capacity to alter their skeletal morphology and physiology through phenotypic plasticity (Todd 2008), thereby maintaining a Reynold’s number (Re) that decreases the thickness of diffusion boundary layers and promotes high rates of mass transfer to and from the coral tissue (Patterson 1992; Lesser et al. 1994; Gardella and

Edmunds 2001). As the effects of OA are anticipated to reduce calcification with consequences affecting the morphological structure of the reef, it is important to take into consideration how carbon delivery to the site of calcification will be mediated by flow and

2- skeletal morphology under reduced Ωarag and [CO3 ] to determine the impacts of OA on calcification (Langdon and Atkinson 2005; Jokiel 2013).

Scleractinian Morphology and Ocean Acidification

Porites rus is an abundant and morphologically plastic hermatypic coral found throughout the tropical Pacific Ocean (Veron 1986), and is most common on fringing reefs where it can be distributed along steep vertical gradients from 0-60 m depth (Jaubert

1977; Padillo-Gamiño et al. 2012). P. rus has branching and plating morophologies with respect to high and low light conditions (Jaubert 1977). These morphological differences within a single species provide an ideal model system to test for differences in calcification rates between contrasting morphologies under elevated PCO2 conditions.

Chapter 3 describes a series of experiments that tested for differential effects between branching and plating morphologies of P. rus exposed to elevated PCO2 under ecologically relevant conditions of light, temperature, and flow.

Current State of Tropical Reefs

The objective of this study was to assess how communities may change in tropical reefs as a result of past and future disturbances. First, I investigated the changes in gorgonian abundances that create “animal forests” or “animal canopies”, which may be more resilient to disturbances than scleractinian corals. As percent cover of scleractinians in the Caribbean is expected to continue to decline or remain low, gorgonians may provide an alternative habitat for reef-dwelling organisms. Second, I investigated the response of scleractinians to OA, a press disturbance that is expected to increase as anthropogenic burning of fossil fuels continues. I focused on skeletal morphology derived from plasticity because it is a common coral trait, to determine if branched and plated morphologies of Porites rus differed in susceptibility to the effects of OA. The intention of testing responses based on morphology was to understand how scleractinians might change as OA conditions intensify.

Chapter 2

Decadal-scale changes in the density of Caribbean gorgonians

Introduction

Over the least several decades there have been large-scale declines in cover of scleractinian corals on coral reefs throughout the world (Gardner et al. 2003; De’ath et al.

2012) as a result of multiple natural and anthropogenic disturbances (Nyström et al.

2000). These changes have probably been underway since the late 1700’s (Jackson 1997) but have accelerated in recent times despite efforts promoting effective management of coral reefs (Bellwood et al. 2004). In the Caribbean, large disturbances impacting coral reefs have attracted widespread attention since the early 1960’s (Goreau 1964; Hughes et al. 1999), with these events including hurricanes (Woodley et al., 1981; Gardner et al.,

2003), die-off of the echinoid Diadema antillarum (Lessios et al., 1984), overfishing

(Jackson et al. 2001), disease outbreaks (Goreau et al., 1998; Schutte et al., 2010), and large-scale bleaching due to increased seawater temperature (Glynn 1993; McWilliams et al. 2005). Overall, the percent cover of scleractinians has declined ~80% throughout the

Caribbean from 1977 to present, with the region-wide mean cover now ~10% (Gardner et al. 2003, Schutte et al. 2010, Jackson et al. 2013). The decline in cover of scleractinians is associated with a reduction in topographic complexity of the benthos (Alvarez-Filip et al. 2009), which has negative implications for fish and invertebrates using coral reefs as habitat (Idjadi and Edmunds 2006).

In addition to region-wide declines in cover of scleractinians, there is evidence from shallow reefs (i.e., ≤ 25 m) that assemblages of scleractinian species have changed compared to that prevailing just a few decades ago (Loya et al. 2001; Coté et al. 2005;

Green et al. 2011; Edmunds 2013). These changes have important implications because they suggest that the goods and services provided by coral reef ecosystems (Moberg and

Folke 1999) may be changing in form as well as magnitude, because of a reduction in scleractinian diversity (Worm et al. 2006). For example, the replacement of massive corals like Orbicella and Diploria in the Caribbean with weedy corals (sensu Knowlton

2001) like Agaricia and Porites (Edmunds and Elahi 2007; Green et al. 2008) probably will create a reef that is less resistant to the mechanical forces created by storms, and will exhibit impaired abilities to calcify and form rugose communities (Alvarez-Filip et al.

2013).

As the global decline in cover and composition of scleractinians has intensified, many Caribbean reefs have undergone a transition from a coral- to macroalgae- dominance (Hughes 1994; Rogers and Miller 2006; Mumby 2009). However, a shift in community structure favoring macroalgae is only one of several possible outcomes following the death of scleractinians, and in a few cases, sponges, ascidians, or non- scleractinian anthozoans have come to dominate the benthos (Norström et al. 2009). In the Caribbean for example, few studies have described changes in abundance of sponges and gorgonians on reefs that have been depleted of scleractinians (Wulff 2006; Norström et al. 2009; Ruzicka et al. 2013).

Gorgonians provide an interesting example of changes in coral reef communities involving taxa other than scleractinians, because there are reasons to expect gorgonians to be less severely affected by disturbances that are detrimental to scleractinians. For instance, gorgonians are less severely affected by bleaching than scleractinians (Lasker et al. 1984, Ruzicka et al. 2013) and hurricanes (Yoshioka et al. 1987; Witman 1992). Their flexible branches can resist the drag associated with waves (Lewis and Wallace 1991;

West 1998), and can exploit storm-mediated fragmentation for asexual reproduction

(Lasker 1984). Compared to scleractinians, gorgonians show strong ability to pre-empt vacant space on benthic surfaces. Presumably in part because of these attributes, gorgonians cover has increased almost three fold in the Florida Keys from 1999-2009 along shallow forereefs (Ruzicka et al. 2013).

Against a backdrop of changes in benthic community structure on Caribbean coral reefs, this study tested the hypothesis that gorgonians have increased in abundance on shallow coral reefs over the last 20 to 45 years. Two approaches were taken. First, changes over time in the abundance (colonies m-2) of gorgonians were assessed on a local-scale (≤ 20 km [Mittelbach et al. 2001]) in St. John, U.S. Virgin Islands, using photoquadrats (0.25 m2) recorded from 1992 to 2012 on shallow reefs along the south shore of the island (Edmunds 2002, 2013). Second, to expand the spatial scale of the analysis, changes over time in the abundance of gorgonians also were evaluated on a regional-scale (i.e., 200-4000 km [Mittelbach et al. 2001]) throughout the Caribbean, using data compiled from the present study as well as peer-reviewed literature and monitoring reports. The data compilation focused on studies reporting gorgonian

densities on reefs at ≤ 20 m depths in units comparable to those obtained for St. John

(i.e., colonies m-2). In this compilation, absolute densities (i.e., individual per unit area) were favored over percentage cover to avoid biased cover estimates arising from arborescent gorgonians (hereafter AGs), which are common in St. John (Fig. 1) and can obscure the benthos beneath their canopy.

Materials and methods

Local-scale assessment

For the local-scale (≤ 10 km) evaluation, gorgonian densities were measured along the south shore of St. John, US Virgin Islands, within the Virgin Islands National

Park (VINP) and Biosphere Reserve (Rogers et al. 2008). Within this protected area, the community structure of scleractinians has been studied since the 1950’s (Randall, 1961;

Collette and Earle 1972) and in a systematic manner since 1987 (Rogers et al. 1991;

Rogers and Beets 2001; Edmunds 2013). Six sites on shallow reefs (7-9 m depth) were selected randomly between Cabritte Horn and White Point in 1992, and have been censused annually thereafter (Fig. 2). At each site, photoquadrats (0.25 m2) were recorded at random positions along a fixed transect, with 20-m transects and 17-20 photoquadrats site-1 between 1992 and 1999, and 40-m transects and 40 photoquadrats from 2000 to present (Edmunds 2002, 2013). Before 2000, photoquadrats were recorded with a Nikonos™ V camera (fitted with a 28-mm lens, two Nikonos SB 105 strobes, and

Kodachrome 64 film) mounted onto a quadrapod that held the camera perpendicular to the seafloor (Edmunds 2002, 2013). In 2000, the method was upgraded to digital images using first a 3.3 megapixel camera (2000-2006, Nikon Coolpix 990) and then a 6.1

megapixel camera (2007-2012, Nikon D70). The camera framer remained unchanged throughout the sampling, and the images allowed objects ≥ 10-mm diameter to be resolved.

In earlier work, the six sites have been analyzed separately (Edmunds 2002), but more recently they have been combined as the “pooled random sites” (hereafter PRS

[Edmunds 2013]) to provide a single description of the near-shore fringing reef habitat.

Photoquadrats from the PRS were used to create a 20-y record in which gorgonian abundances were quantified at 5-y intervals (images archived at http://mcr.lternet.edu/vinp). Although the photoquadrats have been recorded annually, a coarse-grain analysis was used to capture the contrast created between samplings separated by five years: 1992, 1997, 2002, 2007, and 2012.

In 2011, eleven genera of gorgonians were found at the PRS during field surveys:

Antillogorgia (formerly Pseudopterogorgia [Williams and Chen 2012]), Briareum,

Erythropodium, Eunicea, Gorgonia, Muricea, Muriceopsis, Plexaura, Plexaurella,

Pseudoplexaura, and Pterogorgia. Many gorgonians could not be identified to species in situ, which would have required analyses of spicules, and therefore the analyses were constrained to generic resolution. Of the eleven gorgonian genera found on the shallow reefs of St. John, Eunicea, Plexaura, Plexaurella, and Pseudoplexaura could be distinguished from one another as adults, but it proved difficult to identify small colonies

(i.e., < 12 cm tall) with the same resolution; these four genera therefore were pooled and scored as “sea rods” (hereafter SRs). Although Erythropodium has long been present in

St. John (e.g., Witman 1992), it was difficult to detect the encrusting colonies of this species in planar photoquadrats of rugose surfaces, particularly when their polyps were not extended. Further, when colonies of E. caribaeorum were detected in photoquadrats, often it was impossible to resolve the perimeters of colonies for the purpose of counting individuals. Therefore E. caribaeorum was excluded from the analysis.

The aforementioned retrospective analysis was conducted using photoquadrats from 1992 to 2013, and was augmented with field sampling surveys in July and August

2013 that were used to compare field counts with estimates from photographs taken at the same time. To quantify the discrepancy between densities of AGs estimated from photoquadrats versus in situ counts, AGs at the six sites constituting the PRS were censused underwater with 0.25 m-2 quadrats placed at the same positions along the 40 m transect at which photoquadrats were recorded (n = 40). Photoquadrats were used to estimate mean densities of AGs at each of the six sites and mean densities of each genus pooled across sites as described above. The relationship between mean densities of pooled AGs recorded in situ and in photoquadrats using sites as replicate (n = 6) was used to determine the efficacy of estimating AG densities from the photoquadrats. The relationship between mean densities by genus (Antillogorgia, Briareum, Muricea,

Pterogorgia), SRs, and pooled AGs across sites recorded in situ and in photoquadrats was used to determine the efficacy of estimating rare and common AG genera from the photoquadrats.

Despite the utility of photoquadrats as a unique historic record of scleratinian community structure, these records were an imperfect tool for quantifying the abundance of AGs (and Millepora [Brown and Edmunds 2012]). The archived photoquadrats were difficult to use in identifying gorgonians to species (described above), and because the three dimensional structure of AGs could not be captured in planar images. AGs sometimes only filled a portion of the photoquadrat, other times a single colony filled an entire photoquadrat when it was distorted beneath the camera framer, and at other times multiple colonies located outside the photoquadrat all contributed a few branches to the planar image. To avoid these problems, enumeration was constrained to counts of colonies with their basal holdfast within the photoquadrat, and in all cases an individual was defined by the holdfast.

In addition to the surveys conducted in 2013 to establish the efficacy of estimating densities of AGs in photoquadrats, AGs also were surveyed in 2012 and 2013 at Booby Rock, 1.3 km east of Cabritte Horn (Fig. 2). Local knowledge suggested that gorgonians occurred in unusually high abundances in this location (Fig. 1), and therefore

Booby Rock was used to describe the upper range of contemporary gorgonians densities in St. John. Gorgonians were censused using 20 quadrats (0.5 x 0.5 m) placed at random points along a 40 m transect placed along the 7-9 m depth contour. As there were no historic data for this location, gorgonian densities were not used in the contrast of gorgonian abundances over time in St. John, although the results from Booby Rock were included in the regional-scale assessment.

Regional-scale assessment

To test the null hypothesis that densities of AGs have remained unchanged over the last 6 decades throughout the Caribbean, densities of AGs were obtained from 32 studies extracted from the peer-reviewed literature, annual monitoring reports, and the St.

John abundances from the local-scale assessment. These sources were identified from standard bibliographic search techniques, references in key publications, and expert referrals (Table 1). This analysis was restricted to results reporting the abundance of colonies per area, which evaluates population size (i.e., number of individuals) without encountering the bias inherent in using percentage cover for arborescent and flexible taxa forming canopies. This restriction resulted in the exclusion of several studies that have used percentage cover of gorgonians to record abundances (i.e., Ruzicka et al. 2013).

Statistical Analysis

For the local-scale analysis in St. John, the association between densities of AGs obtained from photoquadrats and in situ counts was tested with a Pearson correlation, and the relationship between the two methods described using Model II regression. The equation for this line quantified the accuracy of photoquadrats in assessing population density of AGs, and it was used to adjust density estimates from photographs to more closely resemble in situ counts. To test for changes in the abundance of AGs over time in

St. John, densities were compared among years with a one-way ANOVA, first with AGs pooled among taxa, and then for the most abundant genera (Antillogorgia and Gorgonia) and the pooled sea rods (SRs) category. Where a significant effect of time was detected, post-hoc analyses by Tukey HSD test was used to determine which pairs of years differed

statistically. For the regional-scale analysis, a Model II regression was used to describe the changes of gorgonian densities over time. This analysis was completed first for all gorgonians (i.e., pooled among taxa), then for mean densities of the two most common genera (Antillogorgia and Gorgonia) and SRs found in 14 years between 1973 and 2013.

All analyses were conducted with Systat Version 10 (Systat Software Inc., Chicago,

USA). The statistical assumptions of normality and homoscedasticity were tested through graphical analysis of residuals.

Results

Local-scale assessment

For the analysis comparing methods, the density of all AGs (i.e., pooled among taxa) counted in situ and from photoquadrats at the six sites were positively correlated (r

= 0.99, df = 4, P < 0.001) as were densities separated by Antillogorgia, Gorgonia, and

SRs (r ≥ 0.82, df = 4, P ≤ 0.04). The Model II regression describing the relationship between the two methods was defined by the equations (Fig. 3). These relationships show that mean densities from in situ surveys were predictably higher than were obtained using photoquadrats. The discrepancy between methods was accentuated for rare gorgonians like Muricea or Pterogorgia (densities of ca. 0.03 and 0.02 colonies m-2) compared to the more abundant gorgonians and pooled AGs.

The photoquadrats at the PRS provided 102 to 246 records year-1 for the five years analyzed from 1992 to 2012, and 55% of the photoquadrats (n = 944) contained at least

one gorgonian holdfast. Between 1992 and 2007 (n = 698 photoquadrats), gorgonian holdfasts were found in 49-52% of the photoquadrats in each of the four years analyzed, but in 2012 this proportion increased to 65% (n = 246 photoquadrats). Densities of AGs

(pooled among taxa) differed significantly among the five sampled years (F4,939 = 6.91, P

< 0.01) and post-hoc analyses revealed that densities were higher in 2012 compared to

1997, 2002, and 2007 (P ≤ 0.01); there was a trend for densities in 2012 to be higher than in 1992 (P = 0.06). Densities of AGs did not differ between any other pair of years (P ≥

0.74). Densities of AGs were similar between year 1992 and 2007, varying between 3.51

± 0.46 colonies m-2 (1997) and 4.24 ± 0.57 colonies m-2 (1992), but increased 42% to

6.03 ± 0.45 colonies m-2 in 2012 (mean ± SE, n = 102-246).

The AG fauna was composed largely of Antillogorgia, Gorgonia, and SRs (18%,

17%, and 57% of the colonies, respectively), which together accounted for 92% of all

AGs encountered in 5 census years (n = 4,202 colonies) (Figs. 3). Mean densities of

Antillogorgia, Gorgonia, and SRs displayed dissimilar changes over time across the 20-y study. For Antillogorgia, mean density (± SE) remained relatively constant at 0.78 ± 0.07

-2 colonies m (F4,939 = 0.72, P = 0.58) (Fig. 3B). Mean density of Gorgonia changed significantly from 1992 to 2012 (F4,939 = 3.71, P = 0.01), with different densities in 1992

(0.31 ± 0.12 colonies m-2) versus 2012 (1.01 ± 0.16 colonies m-2) (± SE, n = 102 and 246,

P = 0.02), but not among intervening years (Fig. 3C, P > 0.07). Mean densities of SRs differed among the five years sampled (F4,939 = 7.84, P < 0.01), with mean densities in

2012 (3.79 ± 0.35 colonies y-1) significantly higher than in 1997, 2002, and 2007 (P ≤

0.01), but not 1992 (2.86 ± 0.45 colonies y-1) (P = 0.35). From 1997 to 2007, mean SR

densities remained between 1.64 ± 0.31 colonies m2 (1997) and 2.50 ± 0.22 colonies m2

(2002, n = 107 and n = 248, respectively), but they increased 68% from 2007 (2.25 ±

0.25 colonies m-2) to 2012 (3.79 ± 0.35 colonies m2) (n = 246, P < 0.01) at a rate of ~0.3 colonies m-2 y-1 (Fig. 3D).

Regional-scale assessment

Screening of the literature revealed 32 studies on near-shore reefs throughout the

Caribbean that reported gorgonian abundance (colonies m-2) at sites ≤ 25 m depth for the present study. These studies originated in 28 peer-reviewed articles, 2 online sources from the Coral Reef Evaluation and Monitoring Project (2011 and 2012) of the Florida

Fish and Wildlife Commission/Fish and Wildlife Research Institute, the United Nations

Educational, Scientific and Cultural Organization (Table 15 in the Caribbean Coastal

Marine Productivity Program Data Report 1994-1995), and the present surveys in St.

John. Together, these sources described densities of AGs from 353 surveys conducted at

257 sites throughout the Caribbean over 45-y from 1968 to 2013. The surveys were located in the Florida Keys (50% of total surveys), Cuba (13%), Honduras (7%), Mexico

(7%), Colombia (1%), Belize (2%), Jamaica (7%), Netherland Antilles (2%), Panama

(2%), Bermuda (1%), Dominican Republic (1%), Puerto Rico (1%), Turks and Caicos

Islands (1%), Bahamas (0.5%), Curaçao (0.5%), US Virgin Islands (2%), Trinidad and

Tobago (1%), and Venezuela (1%). Sampling effort increased with time, with 28 surveys from 1965 to 1989, but 310 surveys from 1990 to 2013. The greatest number of surveys was conducted between 2000 and 2004, when 91% of the studies were conducted in the

Florida Keys (n = 100) (Chiappone, 2006; Miller et al., 2006). Only eight articles

contained repeated surveys that monitored changes in benthic communities over 2 to 20-y

(e.g., Keck, 2004). All study sites were in relatively shallow seawater (≤ 25 depth), and the overall mean depth of the studies was 10.9 ± 0.5 m (± SE, n = 176); 22% of the study sites were in seawater ≤ 7 m deep (Fig. 1, Table 1). The study sites were used to provide a mean density of AGs (pooled among taxa, but excluding Erythropodium) by survey year, and when possible they were used to calculate annual mean densities for

Antillogorgia, Gorgonia and SRs, which provided data comparable in taxonomic resolution to that obtained for St. John (Fig. 5B-D). In a few studies, densities of a single genus (e.g., Lasker and Coffroth 1998) or several genera subsampled from the gorgonian fauna were reported (e.g., Dahlgren 1989) and inclusion of these data in our analysis resulted in sample sizes that varied depending on the taxonomic resolution under consideration.

The densities of gorgonians in the data compilation varied nearly three orders of magnitude depending on survey year and location. The lowest densities were recorded at

Isla Lobos Reef at 25 m in Yucatan, Mexico in 1991 (0.03 colonies m-2 [Jordan-Dahlgren

2002]), and the highest densities in Biscayne Bay (Florida Keys, 5 m depth) in 2001

(41.80 colonies m-2; Kuffner et al. 2010) (Fig. 5). The number of surveys conducted throughout the Caribbean that met the selection criteria increased over time, with 18 surveys encountered between 1980 and 1989, but 152 between 1990 and 1999. Survey intensity for each study differed significantly across spatial and temporal scales, with major differences in the inclusion of gorgonian genera and species. The gaps in the data

compilation create limitations for inferences drawn from the trends of gorgonian density over time.

Overall (i.e., pooled among taxa), gorgonian densities increased over time (F1,351

= 14.62, P < 0.01), with densities increasing at 0.17 ± 0.34 colonies m-2 decade-1 between

1968 and 2013 (Fig. 5A). However, the regression explained a small proportion of the variance in gorgonian density (r2 = 0.166) and therefore the estimate of the slope (i.e., change over time) must be interpreted with caution. Of the 32 studies included in the data compilation, densities of gorgonians were available to support separate analyses of

Antillogorgia, Gorgonia, and SRs in 14 studies (Fig. 5B, C and D). There was no association between densities of Antillogorgia, Gorgonia, and SRs and time between

1973 and 2013 (r ≤ 0.486, F1,12 ≤ 3.71, P ≥ 0.08). The lowest mean (densities of

Antillogorgia were 0.08 colonies m-2 in 1987 (Columbia, 9-11 m depth) (Botero, 1987) and the highest was 3.21 colonies m-2 in 2012 (St. John, USVI and Florida Keys, 9-15 m)

(E. Lenz unpublished data; CREMP 2012 Report). For Gorgonia, mean densities were lowest at 0.01 colonies m-2 in 1994 (Colombia, 10.5-20.5 m depth) (Sanchez, 1997) and highest at 5.16 colonies m-2 in 2001 (Key Largo, Florida Keys, 1-15 m depth) (Chiappone et al. 2003). The lowest mean density of SRs was 0.10 colonies m-2 in 1994 (Colombia, 4-

20.5 m depth) (Sanchez et al. 1997) and highest at 8.82 colonies m-2 in 2013 (St. John,

USVI, 9 m depth).

Discussion

The percent cover of scleractinians on reefs throughout the Caribbean has been measured for > 60 years, and the resulting trends illustrate the destructive impacts on coral cover of anthropogenic and natural disturbances (Bellwood et al. 1992; Hughes,

1994). These results underscore the importance of long-term studies in detecting changes in coral community structure (Gardner et al. 2003), but few of these studies have considered benthic organisms other than scleractinians, several functional groups of algae, and a few key invertebrates such as Diadema antillarum (Gardner et al. 2005;

Schutte et al. 2010; Edmunds 2013). This is unfortunate because an important part of understanding the causes of declining coral cover is elucidating the role of other taxa in driving the changes and evaluating their potential to increase in abundance following (or concurrent with) population declines in scleractinians. The role of shifting abundances of echinoids and asteroids in mediating the death of scleractinians (Hughes et al. 1987), and the potential for actiniarians and ascidians to exploit vacant space on denuded reefs

(Chadwick and Morrow 2011), provide examples of the importance of “non- scleractinian” taxa in the changes taking place in coral reef community structure.

My analyses reveal that AGs (pooled among taxa) have become more abundant at both local- and regional-scales (since 1992 and 1968, respectively), with these effects driven largely by Gorgonia and sea rods (but not Antillogorgia). This potential shift in coral reef community structure in favor of AGs has important implications for the functional attributes of contemporary coral reefs. While AGs may have the capability to fulfill some of the habitat provisioning, by creating tall, flexible canopies over the

benthos to sustain the previous trophic functions of scleractinians corals (Rossi et al.

2012), they cannot provide the wave resistant calcareous framework that creates the foundation of a coral reef and provides near-shore protection from damaging waves.

In St. John at the PRS where the abundance of AGs increased, macroalgae cover has increased while scleractinians have remained low without signs of recovery over the last 20-y (Edmunds 2013). The mean percent cover of scleractinians corals on the near shore, shallow reefs (measured at the PRS as in Edmunds 2013) has remained ≤ 4% over the last 20 y, whereas the mean cover of macroalgae has increased from 14.4 ± 1.7 % in

1992 to 25.4 ± 1.2 % in 2011 (± SE, n = 20), equivalent to a rate of 1.1% y-1 (Edmunds

2013). Over a longer period (1987-2011) mean scleractinian cover on one nearby area

(9-m depth) initially dominated by Orbicella annularis (formerly Montastraea annularis) declined 80% from 44.6 ± 3.4% to 6.7 ± 1.7%, while mean macroalgal cover increased from 2.2 ± 0.4% to 37.8 ± 2.4% (all ± SE, n = 30) (Edmunds 2013). Scleractinians generally compete poorly with macroalgae under similar conditions (Tanner 1995;

Lirman 2001), and this has been widely cited as a leading process favoring a change in phase of coral reefs to favor macroalgae (Mumby 2009).

Despite the increase in macroalgae, AG populations do not appear to be negatively effected. Mean abundances of AGs remained similar from 1992 to 2007 at

3.89 ± 0.20 colonies m-2 (± SE, n = 698), but increased 55% in the most recent six years of the study to 6.03 ± 0.57 colonies m-2 (in 2013, ± SE, n = 246). Much of this increase was contributed by the increased abundances of Gorgonia and SRs (described below) in

2012. At the time of the most recent surveys in 2013, ~19% of the gorgonians were juveniles in the small size class (i.e., ≤ 4 cm) (sensu Lasker and Coffroth 1983) which suggests successful recruitment along these shallow reefs. AGs may not be a direct result of changing cover of scleractinians (Gardner et al. 2003). However, it is striking that gorgonians have increased in abundance on reefs that had 26% cover of macroalgae in

2012, and where gorgonians thrive under conditions of potentially strong competition for space on the benthos with macroalgae.

AGs have high linear growth rates and flexible morphologies that allow them to succeed in many reef environments where scleractinians are at a disadvantage (Brazeau and Lasker 1992; Lasker et al. 2003). Unlike scleractinians, AGs can potentially growing as much as 17.8 cm y-1 (Lasker et al. 2003). Established AGs provide examples of tree morphologies (sensu Jackson 1979), that rise above the benthos to exploit resources in multiple layers of seawater up to 1-m in the water column, and create colonies whose distal tissues are heavily dependent on the structural integrity of small areas of basal attachment (Jackson 1979). A few Caribbean scleractinians have similar morphologies – for example, Acropora palmata, A. cervicornis and Dendrogyra cylindrus – but populations of Caribbean acroporids have been greatly depleted by white band disease

(Holden 1996; Patterson et al. 2002), and in general, several decades of disturbances have left reefs substantially reduced in topographic complexity that once was created by scleractinians (Alvarez-Filip et al. 2011). The results imply that AGs are occurring at a population density where colonies may overcome competitive restraints on the benthos with high linear growth of upright branches to exploit the water column and establish

thick canopies or ‘animal forests’ (sensu Rossi 2013). Unlike scleractinians however, the flexible morphology of AGs is resistant to the effects of drag as they deform in water motion (Koehl 1996). Therefore, AGs are more likely than scleractinians to survive the effects of large waves, at least until their diminutive holdfasts eventually rip off the benthos, which appears to be rare and often the result of substratum (rather than gorgonian) failure (Wahle 1985).

This analyses from St. John (i.e., the local-scale) relied heavily on a retrospective analysis conducted using archived planar images (i.e., photoquadrats), yet these images were an imperfect means to quantify erect organisms with tree-like morphologies. I did not expect counts of AGs from photoquadrats and from in situ surveys to be identical, but my cross-calibration analysis revealed a strong association between the two techniques, with a downward bias in densities estimated from photoquadrats that intensified with AG density (i.e., the calibration line had a slope < 1 and passed approximately through the origin). This discrepancy appeared largely to be a result of a limited ability to detect the smallest colonies, with this effect intensifying with colony densities that made it more difficult to see the underlying benthos. Based on a more detailed analysis using the best color images available from St. John (Cabritte Horn, 2012), only 18% of all AGs (n =

150 colonies) were in the smallest size class (estimated to be < ~ 4 cm height) which suggests the potential bias in my method attributed to the smallest colonies is small.

Although it is not possible to test the efficacy of the photoquadrat sampling in the legacy images retrospectively, inspection of the original 35 mm slides (recorded using

Kodachrome 64 color slide film that contains ~20 megapixels of data) suggested that the

smallest colonies were recorded with similar efficiency in the older film and in the most recent digital images.

Based on the empirical calibration relationship generated from comparing AG counts from photoquadrats and in situ surveys with both conducted in 2013 (Fig. 3A), the downward bias in estimates of AG densities from photoquadrats was ~ 15% at 5 colonies m-2, ~19% at 10 colonies m-2, and ~32% at 15 colonies m-2. Thus, the bias dependes upon the mean actual density of AG encountered, but at the mean density encountered in

2013 (6.03 colonies m-2), the downward bias caused by dependence on photoquadrats would be ~16%, and at the density estimated for 1992 (4.24 colonies m-2), the downward bias would be about 20%. Correcting for these effects would result in an increase in density of AGs to a mean of 5.09 colonies m-2 in 1992 to 6.99 colonies m-2 in 2012. The calibration relationships generated from comparing rare and common AG genera from photoquadrats and in situ surveys conducted in 2013 (Fig. 3B) also revealed downward biases in estimates of AG densities from photoquadrats. Uncommon genera, such as

Muricea and Pterogorgia at densities of 0.03 and 0.02 colonies m-2 measured in photoquadrats, respectively, had large downward biases as high as ~88% at mean (±SE) densities of 0.13 ± 0.66 colonies m-2 in situ. For more common genera, such as

Antillogorgia (1.35 colonies m-2) and Gorgonia (1.35 colonies m-2), the downward bias decreased to ~30% and 28%, respectively. Critically, the upward adjustments suggested by the comparison of AG counts measured in the field versus photographs does not alter the principal conclusions of my study.

The regional-scale analysis yields results for pooled AGs that are consistent with the local-scale analysis and demonstrate a significant increase in AG density over time, here over six decades. Although reported on a different scale (i.e., percent cover) similar results have been reported for gorgonians on shallow reefs in the Florida Keys from 1999 to 2009 (Ruzicka et al. 2013). The regional analysis of decadal-scale changes in density of AGs relied on data garnered from peer-reviewed literature, published reports, and online resources. Often data on densities of AGs were marginalized to small tables or anecdotal comments addressing coral reef community ecology, and this created challenges to finding suitable data for the present compilation. These issues were made more acute by restricting the analysis to surveys reporting AGs as densities of individuals rather than percentage cover (e.g., Ruzicka et al. 2013). My analysis therefore should not be construed as a summary of all possible data on these topics that could be secured.

Rather this data should be considered as a compilation that represents a trend of changing densities over time. Perusal of this compilation reveals high within-time variance that I suspect would not be reduced substantially with more data, and which is likely to reflect the diversity of methods employed to conduct the original surveys. Some of the earliest surveys in the late 1960’s and early1970’s, utilized destructively sampling by clearing plots (Opresko 1973, 1974) which yielded exceptionally high densities (e.g., Lasker and

Coffroth 1982) that are difficult to reconcile with subsequent surveys conducted with more standardized ecological techniques.

Analysis of AGs at a lower taxonomic level to species was challenging given the inherent difficulty of identifying members of this taxon in the field, and the near-

impossibility of identification in photoquadrats. With the benefit of expert identification of voucher specimens from the shallow reefs of St. John (by H. Lasker), and the comparison of such voucher specimens with color images, it was possible to achieve genus, and sometimes species-level resolution of adult colonies, but this capacity did not extend to smaller colonies. Faced with these limitations, a consensus lower-taxonomic resolution consisting of two genera (Antillogorgia and Gorgonia) and one functional group (“sea rods” or SRs) was applied.

The coarse-grain resolution of this study revealed that changing population density over time was heterogeneous within the genera and functional group of AGs.

Gorgonia and SRs sustained significant increases in density over time at both the local and regional scale, whereas Antillogorgia maintained fairly constant population densities.

Antillogorgia has higher growth rates than Gorgonia and SRs (Yoshioka and Yoshioka,

1991) and may be considered a weedy genus that can sustain densities despite disturbances. The branches of Antillogorgia have differential growth rates according to their age, size, and proximity to the stipe (Lasker et al. 2003). The flexible, plume-like branches of Antillogorgia may be beneficial by sweeping over the benthos potentially reducing recruitment and competition of other organisms. Differences in abundance at the genus level may be attributed to dissimilarities in growth rates, recruitment patterns, and habitat preference across gorgonians. Further studies are required to support greater taxonomic resolution of SRs through tagging of individuals in the field to understand population dynamics of each genus. At the local and regional scales, there were dominant genera consistently found within the gorgonian community with these

including Antillorgorgia, Gorgonia, and SRs. Often however, the taxonomic detail of historic studies was insufficient to fully compare changes at the genus level.

Since 2011, the long-term monitoring of coral reefs have expanded beyond macroalgae and scleractinian percent cover in St. John and efforts have begun measuring both densities and percent cover of gorgonians. If percent cover of scleractinians remains at ~10 % or continues to decline in abundance because of anthropogenic and natural disturbances (Hughes et al. 2000; Gardner et al. 2003; Edmunds 2007; Schutte et al.

2010), it is essential for monitoring efforts throughout the Caribbean to expand to include benthic categories beyond scleractinians, macroalgae, and bare space. Expanding monitoring efforts will support effective descriptions and predictions of changes in coral reef ecosystems. Further work is needed to test the capacity of gorgonians to withstand the stressors (e.g. thermal bleaching and competition) that have caused declines in scleractinians. Such mechanisms include further testing of competition between gorgonians and macroalgae (De Nys et al. 2006), particularly as it might affect gorgonian success through outgrowth over macroalgae and resistance to wave stress (Jeyasuria and

Lewis 1987). Further research on gorgonian thermal tolerance is needed to understand the mechanistic differences that allow their success in ENSO events. Such studies may include identifying differences in the strong association between Symbiodinium Clade B in the tissue of resistant gorgonian species compared to scleractinians. Future studies should also investigate heterotrophy and feeding rates of gorgonians as a strategy to mitigate potentially harmful effects responsible for mass scleractinian bleaching (Grottoli et al. 2006).

The community ecology of Caribbean coral reefs has changed as a result of natural and anthropogenic disturbances causing the decline in scleractinian cover

(Jackson et al. 2013). Other benthic taxa may be favored as the disturbances negatively affect scleractinians, in part because other taxa can withstand disturbances. Despite the increasing densities of gorgonians on shallow reefs in St. John and throughout the

Caribbean, understanding of ecological services provided by gorgonians to reef ecosystems requires further research. As the morphologies of gorgonians includes encrusting, plumes, fans, and elongated branching, it is important to understand how these morphologies contribute to the construction of “animal forests” (sensu Rossi et al.

2012), which potentially will serve as an alternative benthic community to that formerly created by scleractinians. However, the transition into a reef dominated by the soft- bodied arborescent structure of gorgonians may be subalternate to the calcareous and topographical complex reefs that scleractinians provided for numerous reef organisms.

Although gorgonians have traits to withstand common disturbances, the animal forest of gorgonians is unequal to the invaluable ecosystem services scleractinians have long supplied.

35 Table 1: The 32 studies used to assess the abundance of arborescent gorgonians on a regional-scale throughout the Caribbean showing

author (year of publication, name of study site surveyd as described in the article, region of survey (country), year the survey was

conducted, depth, and abundance measured standardized to colonies m-2.

Year of Depth Density Author(Year) Study Site Name Location/Country survey (m) (colonies m-2)

Kinzie RA III (1973 ) Acropora cervicornis Discovery Bay, Jamaica 1968 15 3.75 Rock Discovery Bay, Jamaica 1968 15 10.63 Lagoon Discovery Bay, Jamaica 1968 1-3 2.04 Lagoon Discovery Bay, Jamaica 1968 1-3 2.37 Lagoon Discovery Bay, Jamaica 1968 1-3 2.42 Lagoon Discovery Bay, Jamaica 1968 1-3 1.16 Lagoon Discovery Bay, Jamaica 1968 1-3 1.65 Lagoon Discovery Bay, Jamaica 1968 1-3 1.17 Lagoon Discovery Bay, Jamaica 1968 1-3 1.1 Acropora cervicornis zone 1 Discovery Bay, Jamaica 1968 20-25 1.25 Acropora cervicornis zone 2 Discovery Bay, Jamaica 1968 20-25 1.02 Acropora cervicornis zone 3 Discovery Bay, Jamaica 1968 20-25 1.34 Acropora cervicornis zone 4 Discovery Bay, Jamaica 1968 20-25 2.35 Acropora cervicornis zone 5 Discovery Bay, Jamaica 1968 10-15 2.66 Acropora cervicornis zone 6 Discovery Bay, Jamaica 1968 10-15 0.93 Acropora cervicornis mound A Discovery Bay, Jamaica 1968 20-25 2.21 Acropora cervicornis mound B Discovery Bay, Jamaica 1968 20-25 1.8 Acropora cervicornis mound C Discovery Bay, Jamaica 1968 20-25 2.03 Acropora cervicornis mound D Discovery Bay, Jamaica 1968 20-25 2.28

Goldberg WM (1973) Outer reef platform Boca Raton, Florida 1973 16-20 15.2 Patch reefs Boca Raton, Florida 1973 9 34.3 Preston EM, Preston Gorgonian patch reef complex Negro Bank, Puerto Rico 1973 5.43 15-18 JL (1975) Wahle CM (1985) Mixed fore reef Discovery Bay, Jamaica 1977 7 14.6 Rear Zone 1 Discovery Bay, Jamaica 1977 1 0.3 Rear Zone 2 Discovery Bay, Jamaica 1977 1 0.6 Terrace fore reef Discovery Bay, Jamaica 1977 15 2.9 Lasker HR, Coffroth Forereef Carrie Bow Cay, Belize 1982 10 17.44 MA (1983) Forereef ridge Carrie Bow Cay, Belize 1982 18 7.4 Patch reef Carrie Bow Cay, Belize 1982 3-5 5.5 Sand-flat Carrie Bow Cay, Belize 1982 2-3 39.93 Yoshioka PM, Media luna Puerto Rico 1983 6.7 3.53 Yoshioka BB (1991) San Cristobal Puerto Rico 1983 10.6 11.9 Botero L (1987) Chengue b Colombia 1987 9-10 1.24 Cinto Colombia 1987 9-11 2.78 Chiappone M, Craig Key Florida Keys, Florida 1989 2 3.6 Sullivan KM (1994) Fiesta Key Florida Keys, Florida 1989 2 1.5 Dahlgren EJ (1989) Breaker (Acropora/Millepora) Yucatan Peninsula, Mexico 1989 0.4-1.2 1.19 Fore reef 10m Yucatan Peninsula, Mexico 1989 7.5-12.5 2.14 Fore reef 15m Yucatan Peninsula, Mexico 1989 12.6-17.5 1.82 Fore reef 20m Yucatan Peninsula, Mexico 1989 17.6-22.5 2.02 Fore reef 25m Yucatan Peninsula, Mexico 1989 22.6-26 1.95 Fore reef 5m Yucatan Peninsula, Mexico 1989 3.6-7.5 0.88 Lagoon Yucatan Peninsula, Mexico 1989 0.1-6 1.01 Rear reef Yucatan Peninsula, Mexico 1989 1-3 0.54 Chiappone M, Craig Key Florida Keys, Florida 1990 2 5.3 Sullivan KM (1994) Fiesta Key Florida Keys, Florida 1990 2 2.6 Coffroth MA, Lasker Korbiski San Blas Islands, Panama 1990 2-4 0.67 HR (1998) Macaroon San Blas Islands, Panama 1990 3-4 0.08 Marsarkantupo San Blas Islands, Panama 1990 5-10 0 Niatupo San Blas Islands, Panama 1990 3-4 0.57

Pinnacles San Blas Islands, Panama 1990 7-9 0.03 Sail Rock San Blas Islands, Panama 1990 5-10 0.06 Salar San Blas Islands, Panama 1990 5-10 0 Three Sisters Florida Keys, Florida 1990 NA 0.02 Tiantupo San Blas Islands, Panama 1990 2-4 0.13 Jordan-Dahlgren E Alacranes reef Yucatan Peninsula, Mexico 1991 25 2.04 (2002) Cayo Arenas reef Yucatan Peninsula, Mexico 1991 25 1.29 Cayos Arcas reef Yucatan Peninsula, Mexico 1991 25 2.39 Chiappone M, Craig Key Florida Keys, Florida 1991 2 5.9 Sullivan KM (1994) Fiesta Key Florida Keys, Florida 1991 2 0.7 Fish Cay Channel Reef Turks and Caicos Island 1991 3-4.9 1.83 Dahlgren EJ (2002) Isla Lobos reef group Yucatan Peninsula, Mexico 1991 25 0.03 Triangulos reef group Yucatan Peninsula, Mexico 1991 25 1.73 Tuxpan reef group Yucatan Peninsula, Mexico 1991 25 0.41 Ver-Al reef group Yucatan Peninsula, Mexico 1991 25 0.4 Sullivan KM et al. Ambergris Patch Reef Turks and Caicos Island 1991 5-6 1.22 (1996) Leeward Fish Cay Reef Turks and Caicos Island 1991 0.1-3.9 0.67 Admirals Aquarium Turks and Caicos Island 1991 1-1.5 0.03 Ambergris Channel Reef Turks and Caicos Island 1991 5 0.46 Blair SM et al. Station A Poststorm Dade County, Florida 1992 10-12 4.67 (1994) Station A Prestorm Dade County, Florida 1992 10-12 3.62 Station G poststorm Dade County, Florida 1992 18-20 14.14 Station G prestorm Dade County, Florida 1992 18-20 21.86 Station H poststorm Dade County, Florida 1992 18-20 8.38 Station H prestorm Dade County, Florida 1992 18-20 9.9 Station I poststorm Dade County, Florida 1992 16-18 5.33 Station I prestorm Dade County, Florida 1992 16-18 5.81 Station K poststorm Dade County, Florida 1992 18-20 25.62 Station K prestorm Dade County, Florida 1992 18-20 27.76 Station L poststorm Dade County, Florida 1992 16-18 8.19 Station L prestorm Dade County, Florida 1992 16-18 8.48

Station M poststorm Dade County, Florida 1992 10-12 8.81 Station M prestorm Dade County, Florida 1992 10-12 9.38 Station N poststorm Dade County, Florida 1992 16-18 5.81 Station N prestorm Dade County, Florida 1992 16-18 7.95 Chiappone M, Craig Key Florida Keys, Florida 1992 2 4.9 Sullivan KM (1994) Fiesta Key Florida Keys, Florida 1992 2 1.4 Lenz EA, Edmunds PRS St. John, USVI 1992 4.24 9 PJ Unpublished Alcolado PM et al. Cayos Esquivel A10 Sabana-Camaguey Archipelago, Cuba 1994 10-11 8.33 (2008) Cayos Esquivel A20 Sabana-Camaguey Archipelago, Cuba 1994 19-20 9.29 Cayos Esquivel A5 Sabana-Camaguey Archipelago, Cuba 1994 4-6 4.65 Cayos Fragoso B1 Sabana-Camaguey Archipelago, Cuba 1994 1-2 6.07 Cayos Fragoso B10 Sabana-Camaguey Archipelago, Cuba 1994 10-11 10.38 Cayos Fragoso B20 Sabana-Camaguey Archipelago, Cuba 1994 19-21 13.15 Cayos Fragoso B5 Sabana-Camaguey Archipelago, Cuba 1994 4-5 5.56 Cayo Francés C10 Sabana-Camaguey Archipelago, Cuba 1994 10-11 9.91 Cayo Francés C20 Sabana-Camaguey Archipelago, Cuba 1994 14-16 7.61 Cayo Francés C5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 7.19 Cayo Caimán Grande D1 Sabana-Camaguey Archipelago, Cuba 1994 1-2 3.23 Cayo Caimán Grande D10 Sabana-Camaguey Archipelago, Cuba 1994 10 3.8 Cayo Caimán Grande D20 Sabana-Camaguey Archipelago, Cuba 1994 20 2.86 Cayo Caimán Grande D5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 4.83 Oeste de Cayo Guillermo E1 Sabana-Camaguey Archipelago, Cuba 1994 1-2 1.35 Oeste de Cayo Guillermo E10 Sabana-Camaguey Archipelago, Cuba 1994 10-11 4.5 Oeste de Cayo Guillermo E15 Sabana-Camaguey Archipelago, Cuba 1994 14-15 3.03 Oeste de Cayo Guillermo E5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 4.25 Cayo Coco F1 Sabana-Camaguey Archipelago, Cuba 1994 1-2 0.93 Cayo Coco F10 Sabana-Camaguey Archipelago, Cuba 1994 9-10 3.78 Cayo Coco F15 Sabana-Camaguey Archipelago, Cuba 1994 12-15 4.86 Cayo Coco F5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 5.31 Cayo Paredón G1 Sabana-Camaguey Archipelago, Cuba 1994 1-2 1.71 Cayo Paredón G10 Sabana-Camaguey Archipelago, Cuba 1994 10-11 3.06

Cayo Paredón G15 Sabana-Camaguey Archipelago, Cuba 1994 14-16 6.38 Cayo Paredón G5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 3.82 Cayo Confites H1 Sabana-Camaguey Archipelago, Cuba 1994 1-2 1.74 Cayo Confites H10 Sabana-Camaguey Archipelago, Cuba 1994 9-10 5.61 Cayo Confites H20 Sabana-Camaguey Archipelago, Cuba 1994 20 5.85 Cayo Confites H5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 5.16 Cayo Guajaba I10 Sabana-Camaguey Archipelago, Cuba 1994 10-11 8.5 Cayo Guajaba I5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 5.94 Cayo Sabinal J10 Sabana-Camaguey Archipelago, Cuba 1994 10 14.45 Cayo Sabinal J20 Sabana-Camaguey Archipelago, Cuba 1994 18-20 11.87 Cayo Sabinal J5 Sabana-Camaguey Archipelago, Cuba 1994 5-6 10.28 Sanchez JA et al. Lagoonal patch reefs Colombia 1994 4-11 0.98 (1997) Leeward fore-reef terraces Colombia 1994 10.5-20.5 1.06 Windward fore-reef terraces Colombia 1994 10.5-20.5 1.02 UNESCO Bahamas site 2 San Salvador, Bahamas 1994 1.5 0.54 Unpublished Bahamas site 1 San Salvador, Bahamas 1994 1.5 1.26 Belize site 1 Carrie Bow Cay, Belize 1994 NA 1.04 Belize site 2 Carrie Bow Cay, Belize 1994 9-12 1 Bermuda site 1 Twin & Hog Breaker Reefs, Bermuda 1994 7-9 1.7 Bermuda site 2 Twin & Hog Breaker Reefs, Bermuda 1994 7-9 3.64 Colombia site 1 Chengue Bay, Colombia 1994 9-12 2.4 Colombia site 2 Chengue Bay, Colombia 1994 9-12 0.3 Cuba site 1 Cayo Coco, Cuba 1994 NA 0.63 Cuba site 2 Cayo Coco, Cuba 1994 NA 0.61 Bahamas site 1 San Salvador, Bahamas 1995 1.5 0.48 Bahamas site 2 San Salvador, Bahamas 1995 1.5 0.24 Belize site 1 Carrie Bow Cay, Belize 1995 NA 1.19 Belize site 2 Carrie Bow Cay, Belize 1995 NA 0.68 Cuba site 1 Cayo Coco, Cuba 1995 NA 0.5 Cuba site 2 Cayo Coco, Cuba 1995 NA 0.56 Curacao site 1 Santa Barbara, Curacao 1995 NA 1.28

Curacao site 2 Santa Barbara, Curacao 1995 NA 0.72 Jamaica site 1 West Forereef, Jamaica 1995 NA 0.52 Jamaica site 2 West Forereef, Jamaica 1995 NA 0 Trinidad & Tobago site 1 Outer and Eastern Buccoo Reefs, Trinidad and Tobago 1995 NA 0.3 Trinidad & Tobago site 2 Outer and Eastern Buccoo Reefs, Trinidad and Tobago 1995 NA 0.26 Venezuela site 1 Isla Margarita, Venezuela 1995 NA 2.02 Venezuela site 2 Isla Margarita, Venezuela 1995 NA 0.92 Keck JR (2004) Fish Den Roatan, Honduras 1996 12 2 Pillar coral Roatan, Honduras 1996 12 2.53 Tuk's treasure Roatan, Honduras 1996 12 2.53 Lybolt M Middle Deep Florida Keys, Florida 1996 11-15 11.05 Unpublished Middle Hardbottom Florida Keys, Florida 1996 4 8.93 Middle Patch Florida Keys, Florida 1996 4-6 13.64 Middle Shallow Florida Keys, Florida 1996 4-6 6.93 Upper Deep Florida Keys, Florida 1996 12-17 11.91 Upper-Patch Florida Keys, Florida 1996 1-6 11 Upper-Shallow Florida Keys, Florida 1996 1-6 7.97 Torres M et al. La Raya Parque Nacional del Este, Dominican Republic 1996 15-20 2.6 (2001) Ruben Parque Nacional del Este, Dominican Republic 1996 15-20 3.3 Dominicus Parque Nacional del Este, Dominican Republic 1996 15-20 3.1 El Toro Parque Nacional del Este, Dominican Republic 1996 16-20 8.5 Yoshioka PM (2005) Media Luna Puerto Rico 1996 6.7 26.16 San Cristobal Puerto Rico 1996 10.6 27.66 Keck JR (2004) Fish Den Roatan, Honduras 1997 12 2.73 Pillar coral Roatan, Honduras 1997 12 2.67 Tuk's treasure Roatan, Honduras 1997 12 2.67 Lenz EA, Edmunds PRS St. John, USVI 1997 3.51 9 PJ Unpublished Keck JR (2004) Fish Den Roatan, Honduras 1998 12 3.07 Pillar coral Roatan, Honduras 1998 12 2.67 Tuk's treasure Roatan, Honduras 1998 12 2.87 Lybolt M Middle Deep Florida Keys, Florida 1998 11-15 7.9

Unpublished Middle Hardbottom Florida Keys, Florida 1998 4 10.1 Middle Patch Florida Keys, Florida 1998 4-7 2.6 Middle Shallow Florida Keys, Florida 1998 4-6 6.98 Upper Deep Florida Keys, Florida 1998 12-17 11.48 Upper-Patch Florida Keys, Florida 1998 1-6 12.91 Upper-Shallow Florida Keys, Florida 1998 3-8 8.35 Sanchez JA et al. Mid-depth reefs N. lagoon and deep Providence Island, Colombia 1998 2.88 (1998) leeward terrace 6-22 Shallow and mid-depth mixture Providence Island, Colombia 1998 1-8 4.1 Shallow barrier reef pinacles and spurs Providence Island, Colombia 1998 0.27 1-4

Keck JR (2004) Fish Den Roatan, Honduras 1999 12 2.67 Pillar coral Roatan, Honduras 1999 12 2.53 Tuk's treasure Roatan, Honduras 1999 12 1.87 Lybolt M Middle Deep Florida Keys, Florida 1999 11-15 6.56 Unpublished Middle Hardbottom Florida Keys, Florida 1999 4 7.96 Middle Patch Florida Keys, Florida 1999 4-7 9.71 Middle Shallow Florida Keys, Florida 1999 4-6 7.4 Upper Deep Florida Keys, Florida 1999 12-17 11.61 Upper-Patch Florida Keys, Florida 1999 1-6 9.79 Upper-Shallow Florida Keys, Florida 1999 3-7 7.73 Sanchez JA (1999) Plateau Baru Island, Colombia 1999 17-18 0.49 Slope Baru Island, Colombia 1999 21-24 0.99 Keck JR (2004) Fish Den Roatan, Honduras 2000 12 2.8 Pillar coral Roatan, Honduras 2000 12 2.73 Tuk's treasure Roatan, Honduras 2000 12 2.4 Chiappone M et al. Carysfort Reef Florida Keys, Florida 2001 1-15 8.75 (2003) Conch Reef Florida Keys, Florida 2001 1-15 9.28 Davis Reef Florida Keys, Florida 2001 1-15 8.51 Eastern Dry Rocks Florida Keys, Florida 2001 1-15 0.63 Eastern Sambo Florida Keys, Florida 2001 1-15 1.01 Elbow Reef Florida Keys, Florida 2001 1-15 10.15

Fished areas Florida Keys, Florida 2001 1-15 2.83 Fished areas (2) Florida Keys, Florida 2001 7.5-9 8.3 Fished areas (3) Florida Keys, Florida 2001 1-15 10.01 Fished areas (4) Florida Keys, Florida 2001 1-15 8.99 Fished areas (5) Florida Keys, Florida 2001 1-15 11.92 Fished areas (6) Florida Keys, Florida 2001 1-15 12.62 High-relief spur & groove Florida Keys, Florida 2001 1-15 5.19 Low-relief hard-bottom Florida Keys, Florida 2001 1-15 11.02 Lower Keys Region Florida Keys, Florida 2001 1-15 1.99 Lower Keys Region (2) Florida Keys, Florida 2001 1-15 8.99 Middle Keys Region Florida Keys, Florida 2001 1-15 4.3 Middle Keys Region (2) Florida Keys, Florida 2001 1-15 10.98 Molasses Reef Florida Keys, Florida 2001 1-15 7.06 No-fishing zones Florida Keys, Florida 2001 1-15 1.06 No-fishing zones (2) Florida Keys, Florida 2001 1-15 2.31 No-fishing zones (3) Florida Keys, Florida 2001 1-15 8.67 No-fishing zones (4) Florida Keys, Florida 2001 1-15 8.89 Sand Key Florida Keys, Florida 2001 1-15 1.43 Sombrero Key Florida Keys, Florida 2001 1-15 2.31 Upper Keys Region Florida Keys, Florida 2001 1-15 9.25 Upper Keys Region (2) Florida Keys, Florida 2001 1-15 12.62 Western Sambo Florida Keys, Florida 2001 1-15 1.18 Chiappone M et al. Pickles Reef Control Reef 3 Florida Keys, Florida 2001 7.5-9 7.1 (2006) Pickles Reef Control Reef 4 Florida Keys, Florida 2001 7.5-9 4.3 Pickles Reef Experimental Reef 1 Florida Keys, Florida 2001 7.5-9 3.88 Pickles Reef Experimental Reef 2 Florida Keys, Florida 2001 7.5-9 8.5 Fernandez LH et al. Reef crest Florida Keys, Florida 2001 2-4 4.11 (2011) Shallow forereef (9-15m) Florida Keys, Florida 2001 1-15 7.87 Keck JR (2004) Fish Den Roatan, Honduras 2001 12 2.8 Pillar coral Roatan, Honduras 2001 12 2.93 Tuk's treasure Roatan, Honduras 2001 12 2.53

Chiappone M (2006) Carysfort Reef SPA* Florida Keys, Florida 2002 7.62-7.92 5.38 Keck JR (2004) Fish Den Roatan, Honduras 2002 12 2.93 Pillar coral Roatan, Honduras 2002 12 3 Tuk's treasure Roatan, Honduras 2002 12 3.8 Lenz EA, Edmunds PRS St. John, USVI 2002 3.6 9 PJ Unpublished Lybolt M Upper Deep Florida Keys, Florida 2002 12-17 16.04 Unpublished Middle Deep Florida Keys, Florida 2002 11-15 7.27 Middle Hardbottom Florida Keys, Florida 2002 4 11.19 Middle Patch Florida Keys, Florida 2002 4-7 11.24 Middle Shallow Florida Keys, Florida 2002 4-6 9.9 Upper-Patch Florida Keys, Florida 2002 1-6 10.96 Upper-Shallow Florida Keys, Florida 2002 3-8 9.1 Miller SL et al. Carysfort Reef SPA* Florida Keys, Florida 2002 16.76-18.90 14.06 (2006) Ajax Reef Florida Keys, Florida 2002 6.10-7.32 14.81 Between Molasses & French Reef Florida Keys, Florida 2002 10.97-11.89 10.13 Between Star and Triumph Reef Florida Keys, Florida 2002 10.97-12.50 10.69 Between Star and Triumph Reef Florida Keys, Florida 2002 15 4.69 Bird Key Reef Florida Keys, Florida 2002 11.89-14.63 31.92 Coal Bin (East of Cogsrove Shoal) Florida Keys, Florida 2002 16.76-17.37 23 Conch Reef Research Only* Florida Keys, Florida 2002 14.33-15.54 19.81 Cosgrove Shoal Florida Keys, Florida 2002 19.81-21.94 12.88 Dixie Showl Florida Keys, Florida 2002 14.63-15.85 10.56 Dry Tortugas National Park Florida Keys, Florida 2002 12.80-13.72 21.94 Dry Tortugas National Park Florida Keys, Florida 2002 14.33-14.94 8.19 Dry Tortugas National Park Florida Keys, Florida 2002 14.33-14.94 25.33 Dry Tortugas National Park Florida Keys, Florida 2002 15.85-17.37 26.83 East of Pelican Shoal Florida Keys, Florida 2002 11.89-13.41 10.25 Inshore and SW of Pacific Reef Florida Keys, Florida 2002 8.23-10.36 19.88 Inshore of Pacific Reef Florida Keys, Florida 2002 10.06-10.97 11 Ledberry Reef Florida Keys, Florida 2002 14.63-15.24 15.06 Molasses Reef** Florida Keys, Florida 2002 15.85-18.59 10.56

North Dixie Shoal Florida Keys, Florida 2002 14.63-15.85 9.75 North of Ajax Reef Florida Keys, Florida 2002 7.01-8.84 10.94 North of Fowey Rocks Florida Keys, Florida 2002 10.67-11.58 10.88 Offshore Pacific Reef Florida Keys, Florida 2002 15.85-17.68 27.85 Seaward of Watson's Reef Florida Keys, Florida 2002 10.36-10.97 16.13 South of Fowey Rocks Florida Keys, Florida 2002 15.54-19.20 27.19 Southern DTNP (just outside park) Florida Keys, Florida 2002 17.37 19.17 Southwest of Brewster Reef Florida Keys, Florida 2002 5.18-6.10 9.56 Southwest of Pacific Reef Florida Keys, Florida 2002 11.89 17.1 Star Reef Florida Keys, Florida 2002 11.28-12.19 11 West of West Washerwoman Florida Keys, Florida 2002 6.10-7.62 11.44 West Washerwoman Shoal Florida Keys, Florida 2002 5.18-7.01 27.88 French Reef** Florida Keys, Florida 2002 16.46-19.51 6.13 East of Biscayne Bay Florida Keys, Florida 2002 8.84-10.36 8.94 Keck JR (2004) Fish Den Roatan, Honduras 2003 12 2.93 Pillar coral Roatan, Honduras 2003 12 3.27 Tuk's treasure Roatan, Honduras 2003 12 5.33 Kuffner IB et al. November Biscayne Bay, Florida 2003 3.71 28.3 (2010) Bravo Biscayne Bay, Florida 2003 3.69 30.5 Delta Biscayne Bay, Florida 2003 4.13 23 Echo Biscayne Bay, Florida 2003 4.48 29 Golf Biscayne Bay, Florida 2003 4.71 41.8 Hotel Biscayne Bay, Florida 2003 4.37 23.5 India Biscayne Bay, Florida 2003 3.86 34.3 Juliet Biscayne Bay, Florida 2003 4.59 40 Kilo Biscayne Bay, Florida 2003 4.07 29.3 Lima Biscayne Bay, Florida 2003 5.33 37.9 Oscar Biscayne Bay, Florida 2003 4.28 29.8 Papa Biscayne Bay, Florida 2003 5.46 36.5 Etnoyer PJ et al. Fore-reef A12 Saba Bank, Netherland Antilles 2007 11-20 4.19 (2010) Fore-reef A7 Saba Bank, Netherland Antilles 2007 11-20 3.55

Fore-reef CV1 Saba Bank, Netherland Antilles 2007 11-20 4.77 Fore-reef CV2 Saba Bank, Netherland Antilles 2007 11-20 4.64 Plateau D9 Saba Bank, Netherland Antilles 2007 >20 1.5 Plateau E3 Saba Bank, Netherland Antilles 2007 >20 0.18 Plateau E4 Saba Bank, Netherland Antilles 2007 >20 0.46 Plateau Void Saba Bank, Netherland Antilles 2007 >20 0.18 Lenz EA, Edmunds PRS St. John, USVI 2007 4.23 9 PJ Unpublished Espinosa YO et al. Aguadores este Reserva Ecológica Siboney-Juticí, Santiago de Cuba, Cuba 2009 12-17 7.17 (2010) Aguadores oeste Reserva Ecológica Siboney-Juticí, Santiago de Cuba, Cuba 2009 12-17 7.58 El Magnel Reserva Ecológica Siboney-Juticí, Santiago de Cuba, Cuba 2009 12-17 3.58 Juticí Reserva Ecológica Siboney-Juticí, Santiago de Cuba, Cuba 2009 12-17 4.19 Sardinero este Reserva Ecológica Siboney-Juticí, Santiago de Cuba, Cuba 2009 12-17 4.59 Sarinero oeste Reserva Ecológica Siboney-Juticí, Santiago de Cuba, Cuba 2009 12-17 4.54 CREMP Unpublished Admiral Florida Keys, Florida 2011 < 7 2.83 Carysfort Deep Florida Keys, Florida 2011 10-20 9.23 Carysfort Shallow Florida Keys, Florida 2011 < 7 6.45 Cliff Green Florida Keys, Florida 2011 < 7 7 Dustan Rocks Florida Keys, Florida 2011 < 7 3.73 Loggerhead Patch Florida Keys, Florida 2011 < 7 8.74 Porter Patch Florida Keys, Florida 2011 < 7 7.35 Sand Key Deep Florida Keys, Florida 2011 10-20 0.15 Sand Key Shallow Florida Keys, Florida 2011 < 7 6.56 Temptation Rock Florida Keys, Florida 2011 < 7 3.08 Texas Rock Florida Keys, Florida 2011 < 7 0.3 The Maze Florida Keys, Florida 2011 < 7 2.53 West Turtle Shoal Florida Keys, Florida 2011 < 7 6.2 West Washer Women Florida Keys, Florida 2011 < 7 3.58 Western Sambo Deep Florida Keys, Florida 2011 10-20 12.83 Western Sambo Shallow Florida Keys, Florida 2011 < 7 7.18 White Shoal Florida Keys, Florida 2011 < 7 2.3 Admiral Florida Keys, Florida 2012 < 7 12.43

Bird Key Reef Florida Keys, Florida 2012 < 7 11.38 Black Coral Rock Florida Keys, Florida 2012 10-20 10.03 Carysfort Deep Florida Keys, Florida 2012 10-20 16.08 Carysfort Shallow Florida Keys, Florida 2012 < 7 11.28 Cliff Green Florida Keys, Florida 2012 < 7 7.45 Dustan Rocks Florida Keys, Florida 2012 < 7 6.25 Loggerhead Patch Florida Keys, Florida 2012 < 7 0.6 Mayer's Peak Florida Keys, Florida 2012 < 7 14.28 Palmata Patch Florida Keys, Florida 2012 < 7 1.52 Porter Patch Florida Keys, Florida 2012 < 7 10.43 Sand Key Deep Florida Keys, Florida 2012 10-20 13.08 Sand Key Shallow Florida Keys, Florida 2012 < 7 7.33 Sombrero Deep Florida Keys, Florida 2012 10-20 9.85 Temptation Rock Florida Keys, Florida 2012 < 7 3.2 Tennessee Deep Florida Keys, Florida 2012 10-20 14.07 Texas Rock Florida Keys, Florida 2012 < 7 0.25 The Maze Florida Keys, Florida 2012 < 7 4.08 West Turtle Shoal Florida Keys, Florida 2012 < 7 8.25 West Washer Women Florida Keys, Florida 2012 < 7 4.88 Western Sambo Deep Florida Keys, Florida 2012 10-20 11.28 Western Sambo Shallow Florida Keys, Florida 2012 < 7 6.65 White Shoal Florida Keys, Florida 2012 < 7 2.28 Lenz EA, Edmunds Booby Rock St. John, USVI 2012 9 17.2 PJ Unpublished PRS St. John, USVI 2012 9 6.03 Booby Rock St. John, USVI 2013 9 21.65 PRS St. John, USVI 2013 9 8.6

Legends

Figure 1. Representative photograph of a shallow (8-m depth) fringing reef at Booby

Rock, St. John (N18º 18.133’, W64º 42.600’), where some of the highest population densities of gorgonians were found in August 2012. In this location, gorgonians occurred at densities as high as 17 colonies m-2 and were represented mostly by Sea Rods (33%),

Antillogorgia (19%), Eunicea (16%), and Gorgonia (15%). Together, these gorgonians formed a canopy ~ 1-2 m high. For scale, the central Dendrogyra cylindrus colony is about 1 m tall (photo credit: P.J. Edmunds).

Figure 2. Map showing the location of the study sites for the local-scale assessement in

St. John (a), and the sites for the regional-scale assessment across the Caribbean (b;

Table 1). The local-scale analysis from St. John is based on surveys conducted at six sites at 7-9 m depth,which are combined as the Pooled Random Sites (PRS) (Edmunds,

2013): Cabritte Horn = N18° 18.404’, W064° 43.308’, West Tektite = N18° 18.760’,

W064° 43.383’, Wast Tektite = N18° 18.682’, W064° 43.386’, West Little Lameshur =

N18°19.028’, W064° 43.667’, Europa = N18° 18.976’, W064° 43.784, and White Point =

N18°18.910’.

Figure 3. Mean (± SE) densities in St. John, USVI of mean (a) pooled arborescent gorgonians (AGs), and (b) Antillogorgia, (c) Gorgonia, and (d) Sea Rods (SRs) from in situ surveys and photoquadrats (n = 240, 0.25 m-2) that were recorded in 2013 at the six survey sites in St. John (PRS). The best-fit linear relationship from Model II linear regression describes the relationship between the two methods where y is the mean density estimated from the photoquadrats and x is the mean density estimated in situ at each of the six sites.

Figure 4. Mean gorgonian densities from 1992 to 2012 at the PRS (± SE, n = 102-248) used for the local-scale assessment. Gorgonian densities are based on analyses of photoquadrats in 5-y increments, with results shown for: (a) Antillogorgia, Briareum,

Eunicea, Gorgonia, Muricea, Pterogorgia and Sea Rods combined (AGs), (b)

Antillogorgia, (c) Gorgonia, and (d) Sea Rods (SRs). Together, Antillogorgia, Gorgonia, and SRs accounted from 92% of all gorgonians encountered. Overall, gorgonian densities differed among years (F4,939 = 6.912, P < 0.001), with significantly higher densities in 2012 compared to 1997.

Figure 5. Densities of gorgonians (n = 353 surveys) from in situ surveys at a study sites across the Caribbean reported from 32 studies. Densities were taken from peer-reviewed literature, reports, and online sources (Table 1), and included both retrospective and contemporary analyses reported for St. John. Densities of pooled gorgonians (minus

Erythropodium) (a) and mean densities (n = 14) of (a) Antillogorgia, (b) Gorgonia, (c) and (d) Sea Rods (SRs) for each year. The relationship is described by the best-fit line from a Model II linear regression for density against time, from 1968 to 2013.

Figure 1.

Figure 2.

a. St. John, USVI

St. John

1 km N

’ ’ 18o 19.002 N o 64 43.453 W Booby Rock

Great Lameshur Bay

WLL Europa West Tektite White 300 m Point East Tektite

Cabritte Horn

b. Caribbean

Gulf of Atlantic Ocean 25° N Mexico

20° N

Caribbean Sea 15° N

500 km

10° N

85° W 80° W 75° W 70° W 65° W 60° W

Figure 3.

a. AGs b. Antillogorgia 7 y = 0.73x + 0.3 2 y = 0.64x - 0.05 r = 0.99 r = 0.82 6 P < 0.001 P = 0.044

1.5 5

4 1 3

2 0.5 )

-2 1

0 0 0 1 2 3 4 5 6 7 0 0.5 1 1.5 2 c. Gorgonia d. SRs Photoquadrats 2 y = 0.94x + 0.09 5 y = 0.75x + 0.14 (colonies 0.25 m r = 0.92 r = 0.99 P = 0.011 P = 0.001 4 1.5

1 2

0.5 1

0 0 0 0.5 1 1.5 2 0 1 2 3 4 5 In situ (colonies 0.25 m-2)

Figure 4.

a. AGs b. Antillogorgia 7 F4,939 = 6.912 F4,939 = 0.715 P < 0.001 P = 0.582 6

-2 1

c. Gorgonia d. SRs

Colonies m 7 F4,939 = 3.714 F4,939 = 7.844 P = 0.005 P < 0.001 6

0 1992 1997 2002 2007 2012 1992 1997 2002 2007 2012 Year

Figure 5.

a. AGs b. Antillogorgia 50 7 r = 0.20 r = 0.49 F1,338 = 8.611 F1,12 = 3.706 P = 0.004 6 P = 0.078 40 5

30 4

3 20

2 10

-2 1

0 0

c. Gorgonia d. SRs

Colonies m 7 7 r = 0.36 r = 0.33 F1,12 = 1.769 F1,12 = 1.426 6 P = 0.208 6 P = 0.255

5 5

4 4

3 3

2 2

1 1

0 0 1970 1980 1990 2000 2010 2020 1970 1980 1990 2000 2010 2020 Year

Chapter 3

Contrasting morphologies of the Pacific coral

Porites rus are not affected by ocean acidification

Introduction

The morphology of sessile colonial organisms has been studied quanitatively since the 1970’s (Prosser 1973; Wainwright et al. 1976; Chappell 1980). Morphology has been exploited as a part of a physiological response by organisms to overcome stress associated with environmental and biological factors, such as fluid mechanics (Wainwright 1976;

Patterson 1992) and predation (Jackson 1979), respectively. Jackson (1979) codified this focus by identifying six common and predictable morphologies utilized by sessile marine organisms. Tropical hermatypic scleractinians provide prime examples of morphological diversity that is strongly influenced by environmental gradients such as light, hydrodynamic conditions, and sedimentation (Chappell 1980). Examples of this morphological diversity are demonstrated by the contrast between branched and plated morphologies (Jaubert 1977; Jackson 1979), which allow scleractinians to play a key role as ecosystem engineers and create topographically complex habitats (Jones et al. 1997;

Bruno and Bertness, 2000) facilitating high biodiversity (Idjadi and Edmunds 2006).

However, with increased anthropogenic disturbances changing the climate and oceans on a global scale (e.g.. increased sea surface temperature and ocean acidification), structure of scleractinian communities on tropical reefs is being drastically altered.

Formerly dominant species such as weedy and branching corals, like Acropora spp., are

declining in abundance as other corals such as mounding and encrusting corals, like massive Porites spp., persist (Loya et al. 2001). Despite the long history of studies on morphological variation in scleractinians, the current threats of global climate change and

OA suggest it might be timely to reevaluate the role of scleractinian morphology and the parameters shaping their success under projected environmental conditions. Morphology has yet to be investigated as a potential variable for categorizing the vulnerability of scleractinians to OA across a range of functional groups.

Studies of tropical scleractinians have become more common in the last decade, in large part because of the coral reef crisis produced by anthropogenic activities. Dramatic declines in the calcification of hermatypic corals are predicted as OA intensifies (Hough-

Guldberg et al. 2007). The current rates of rising atmospheric CO2 are unparalleled in the past 300 million years (Hönisch et al. 2012; Zeebe 2012) and the ocean has absorbed ~26-

30% of atmospheric CO2. As CO2 dissolves in seawater, the ocean carbonate chemistry is altered as the dissolved inorganic carbon equilibrium is reestablished. Ocean pH has already declined by 0.1 units since the Industrial Revolution (Sabine et al., 2004), with the expectation of additional decreases of 0.10-0.35 by the end of the 21st century under

Representative Concentration Pathways (RCPs) (Moss 2010). These changes will be accompanied by declines in saturation states of calcium carbonate (Ω) from 3 to 1.5-2 in tropical regions (Orr et al. 2005; Ridgwell and Schmidt 2010). Recent studies have shown that calcification of scleractinians generally is negatively impacted by the implications of

OA, but there is considerable variation in responses both within and among taxonomic groups (Melzner et al. 2009; Kroeker et al. 2010; Comeau et al. 2013; Kroeker et al.

2013). Scleractinians have been studied extensively in laboratory settings and these studies reveal high variation in physiological responses, particularly in calcification rates

(Comeau et al. 2013). These differences in responses reflect the wide spectrum of sensitivities and tolerances due to characteristics and mechanisms yet to be defined fully

(Cohen and Holcomb 2009; Comeau et al. 2013). Studies from the early 21st century on ocean acidification have investigated the direct effects of altered carbonate chemistry on the calcification scleractinians, but they lack the application of ecologically relevant environmental parameters in coral reef ecosystems, such as natural levels of light and flow, which are necessary for testing interactive effects.

Light strongly affects scleractinians and their morphologies at the organismal and community level, such as through light enhanced calcification and zonation with depth

(Dustan 1975; Jaubert 1977; Chappell 1980; Muscatine 1990). Tropical hermatypic scleractinians are geographically restricted to clear, oligotrophic waters, and must rely on light and a limited variety of heterotrophic resources for energy. As light diminishes with depth, coral morphology tends to change from highly-branched to foliose forms to reduce photoinhibition and maximize surface area at light-limited depths, respectively (Jaubert,

1977; Chappell, 1980). This morphological zonation is common across and within coral species, particularly those that exhibit a high capacity for morphological plasticity in response to light intensities (Jaubert 1977; Muko et al., 2000; Kaniewska et al. 2013;

Padillo-Gamiño et al. 2013). As coral morphology is coupled strongly with light, both morphology and light levels need to be considered in studies of climate change effects to properly assess responses at the community level. Recent OA studies on coral biology are

expanding beyond the direct effects of elevated PCO2 at the organismal level by incorporating interactive effects, such as temperature and light (Edmunds et al. 2012;

Dufault et al. 2013; Comeau et al. 2014). Calcification rates of Pocillopora damicornis spat (< 4 mm diameter) varied in a hyperbolic manner in response to elevated PCO2 at differential light intensities, which suggested that the response to PCO2 was light-dependent

(Dufault et al. 2013). A comparison between coral genera found that high light intensity mitigated the negative effects of OA on the calcification rates of Acropora horrida but not

Porites cylindrica (Suggett 2013). As coral morphology is strongly coupled with light, both parameters need to be considered in studies of climate change effects to properly assess responses at the community level.

The size and shape of scleractinian colonies are also influenced by other key environmental parameters such as flow, with consequences to physiological performance

(Patterson and Sebens 1989; Patterson 1992; Lesser et al. 1994; Nakamura et al. 2005;

Goldenheim and Edmunds 2011). More specifically, flow across colonies of differing shapes and sizes strongly influences and determines diffusion boundary layers and mass transfer rates (Patterson 1992). This is particularly important for sessile, soft-bodied marine organisms that rely on the delivery of dissolved solutes, such as metabolites and nutrients, such as bicarbonate and nitrogen, to their tissues by transport across external boundary layers (Lesser et al. 1994). In response to varying flow rates, skeletal morphology and physiology of coral colonies can change through phenotypic plasticity, thereby maintaining Reynold’s number that reduce diffusion boundary layers (Patterson

1992; Lesser et al. 1994). For OA experiments, it is important to investigate how flow

2- and scleractinian morphology interact under conditions of reduced Ω and CO3 to determine mitigating or exacerbating negative effects (Langdon and Atkinson 2005).

As the environment can influence skeletal morphology of corals, I compared the calcification response of branched and plated morphologies to ambient and elevated PCO2.

I selected the morphologically plastic coral, Porites rus (Forskâl 1775), which is dominant along shallow fringing reefs (< 4-m depth) in Moorea, French Polynsesia (Jaubert 1977;

Comeau et al. 2013). This study tested the hypothesis that corallum morphology would

differ in sensitivity to elevated PCO2. To test this hypothesis, calcification rates of contrasting morphologies within a single coral species were compared during four OA experiments. The first two experiments were focused on the influence of light under OA conditions on the branched versus plated morphologies. The second two experiments tested the interactive effects of flow and OA on contrasting morphologies.

Materials and methods

A series of experiments was conducted to test the effects of combinations of light, flow, PCO2 on branches and plates of Porites rus. Four combinations of the factors were

tested: Experiment 1 tested for the interactive effects of light and PCO2; Experiment 2, the

interactive effects of light, PCO2, and temperature; and Experiment 3, the interactive effects

of flow and PCO2. These three experiments were conducted in indoor mesocosms.

Experiment 4, tested the interactive effects of flow and PCO2 outdoors under natural irradiance levels. P. rus is an abundant coral in the South Pacific and was used as a model system because of its high capacity for morphological plasticity. The plasticity found in

P. rus consists of branched and plated morphologies, that are formed within a single colony in response to high and low irradiances, respectively. Experiments were conducted at the Richard B. Gump South Pacific Station in Moorea, French Polynesia from April to May 2012 (Experiment 1), January to February 2013 (Experiment 2), and

April to May 2013 (Experiment 3 and 4) at ambient and elevated PCO2 (700 and 1000

µmol) over 2 to 3 week periods.

Collection and acclimation to experimental conditions

For each experiment, branches and plates of Porites rus (Forskål 1775) 3-5 cm in diameter and length were collected haphazardly along a fringing reef in Cook’s Bay

(17°48.96S, 149°81.88W). Branches and plates of P. rus were selected from low and high light regions of the fringing reef at 0.5-2.0 m depth. Mean irradiance where branches and plates were collected was 251 and 1013 µmol photons m-2 s-1, respectively, at noon on a sunny day. Irradiance was measured with 4π MkV-L logging light sensors (JFE

Advantech Co., Kobe, Japan), which measured PAR with a 4π spherical quantum PAR sensor for 60 hours (Fig.1). Samples of P. rus were placed in plastic bags and transported to the laboratory in a cooler filled with seawater. Each coral piece was attached to a plastic base with Z-spar (A-788 Splash Zone Compound) to position the corals upright and allow the coral to be handled without touching the tissue.

After allowing the Z-spar to cure for 48 h, corals were transferred to a 1,000 L tank where experimental irradiance and temperature conditions were maintained, and corals could be placed on a circular table that rotated at two revolutions min-1. Coral

placed on the rotating table were exposed equally to light from four 75-W Light-Emitting

Diode units position directly above (AquaIllumination® LED System Model: Sol Blue,

Ames, IA). The LED lamps operated on a 12:12h light:dark cycle with irradiance gradually increasing from 0 to 100% of maximum output declining gradually increasing over the first 4 h of the day, remaining at 100% for 4 h, and declining in intensity over the final 4 h of the day (Fig. 1).

Light levels were based on the intensities measured with a 4π quantum sensor at

0.5 m and 2 m depth where branches and plates of P. rus were collected as previously mentioned (Fig. 1). For Experiment 1 and 2, half of the branching and plating nubbins were placed directly underneath the LED lamps to acclimate to high light conditions

(~700 µmol photons m-2 s-1) and the other half was placed at the perimeter of the tank in low light conditions (300 µmol photons m-2 s-1) (Fig. 1). Prior to Experiment 3 and 4, corals were placed throughout the acclimation tank to acclimate to light and ambient temperature conditions and to alleviate stress induced by collection. Acclimation lasted one week.

Maintenance of Experimental Conditions

Custom-made mesocosms consisting of 150-L flow-through tanks were used for

Experiment 1 (4 tanks per PCO2), Experiment 2 (3 tanks per PCO2) and Experiment 3 (3

-1 tank per PCO2) (Fig. 2A). Seawater flowed into the tanks at ~200 mL min and was pumped from Cook’s Bay through a filter (100-µm mesh). Each tank was controlled independently for temperature and irradiance. Temperature was measured with a certified

digital thermometer (Fisher Scientific, 15-077-8), and irradiance with the 4π quantum sensor. In all experiments, except Experiment 2 (1 tank per treatment), water temperature was maintained at 28 ºC, Experiment 2 contrasted 30 ºC and 27 ºC.

Ambient PCO2 of 400 µatm was used as the control represening present day PCO2 for

all experiments. The elevated PCO2 for Experiment 1 was targeted at 700 µatm, which is

the PCO2 expected at the end of this century (Representative Concentration Pathways

[RCP] Scenario 6.0). In Experiments 2, 3, and 4, PCO2 was targeted at 1000 µatm to represent a “business as usual” worst-case scenario (RCP Scenario 8.5) (Moss et al. 2010).

Ambient and elevated PCO2 were maintained through bubbling of ambient air and CO2 enriched air into the tanks. Enriched CO2 was mixed by a solenoid-gas regulation system

(Model A352, Qubit, Ontario, Canada), which blended pure CO2 with ambient air.

Experiment 4 involved recirculation of seawater from a 150-L reservoir through a submersible pump (Aqueon Quietflow Submersible Aquarium Utility Pump 1700) with direct bubbling of ambient and CO2 mixed gas. Every other day, 80% of the seawater was replaced with sand-filtered seawater pumped from Cook’s Bay. The temperature (28 ºC) for this experiment was maintained with chillers (Aqua Logic Inc. Delta Star Chillers 1/3

HP).

Seawater carbonate chemistry

Temperature, salinity, total alkalinity (AT) and pH were monitored daily (08:00 h and 18:00 h) by sampling 250 mL of seawater from each tank. When experimental

carbonate chemistry became stable, AT and salinity were assessed every other day (Table

1, 4, 7, and 11). Temperature was measured with a digital thermometer (Fischer Scientific model 15-077-9) and salinity with a YSI 3100 Conductivity Meter. Seawater pHT and AT was calculated from titrations using the gran function in an open-cell potentiometric automatic titrator (T50, Mettler-Toledo International Inc., Ohio, USA) fitted with a

DG115-SC pH probe that was calibrated with TRIS buffers (Dr. Andrew Dickson, Scripps

Institution of Oceanography). Standard operating procedures 3b (Dickson et al. 2007) with certified acid titrant (0.1N HCl and 0.6M NaCl from Dickson Laboratory, Scripps) were followed. AT of certified reference materials (Dickson Laboratory, Scripps) was determined prior to seawater titrations and provided values ± 3.0 µmol kg-1 (Experiment 1 and 2, Batch 108, n = 10) and ± 1.0 µmol kg-1 (Experiment 3 and 4, Batch 122, n = 25) of

- -2 certified values. Carbonate seawater chemistry (PCO2, HCO3 , CO3 , aragonite saturation

-1 state [ΩArag], and DIC) was calculated using AT (µmol kg ), pH, salinity and temperature with the R software package “seacarb” (Lavigne and Gattuso 2011).

Experiment 1 & 2: Effects of light and PCO2

To test for differential calcification responses between branches and plates of

Porites rus, two experiments were conducted at high and low light intensities under OA conditions. Experiment 1 tested the effects of light and PCO2 to evaluate the response of P. rus under ecologically relevant light intensities and elevated PCO2 values expected in 100 y

(700 µatm). Experiment 2 tested the interactive effects of increased PCO2 (1000 µatm) and high temperature (30 ºC).

For Experiment 1, irradiances were ~215 µmol photons m-2 s-1 (along the margin of the tank) and ~1,006 µmol photons m-2 s-1 (directly under the LED lamp). Light was measured weekly, with the sensor placed in the center and along the front of each tank where corals were located for high and low light conditions. Additionally, light was measured continuously for two days with MkV-L logging light sensors (JFE Advantech

Co., Kobe, Japan) that recorded PAR with a 4π spherical quantum sensor (Fig. 1).

Experiments measuring the dissolution rates of clod cards (Thomson and Glenn 1994) determined that corals under the low light condition, along the margin of the tanks, experienced similar flow as the corals in the center of the tank (middle: 1.17 ± 0.11 g h-1 and front: 1.30 ± 0.06 g h-1, mean ± SE; P = 0.341, df = 3). Three plates and branches of

P. rus each were placed randomly in both high and low light conditions within each of the

8 tanks. Corals were rearranged by hand daily to avoid position effects.

For Experiment 2, three branches and three plates of P. rus were placed in each light condition within each of the 6 tanks. A neutral density screen was placed over half of the top of each tank to shade half of the tank to create a low light condition (Fig. 2B).

Irradiance was measured every third day with a 4π light sensor, and the low and high light were 158 ± 3 µmol m-2 s-1 and 758 ± 24 µmol m-2 s-1, for the low and high light, respectively (mean ± SE; P < 0.001, n = 12).

Experiment 3 & 4: Effects of flow and PCO2

To test for the effects of flow on the calcification of branches and plates of Porites rus, two experiments were conducted using high and low flow rates at ambient and high

PCO2. These experiments were designed to test for mass transfer limitation at high PCO2

conditions. For Experiment 3, three branches and three plates were randomly attached in a row to a plastic strip (3 cm wide x 30 cm long) that was inserted inside clear UV- transparent extruded acrylic tubes (o.d. 7.62 cm, i.d. 6.35 cm, 30 cm length); McMaster-

Carr, Santa Fe Springs, CA). Branches and plates of P. rus were detached and rearranged on the plastic strip every other day to avoid position effects created by the location of nubbins relative to the pump at the end of each tube. A small pump (Aquatop Aquarium

Submersible Pump NP80 and NP302) produced unidirectional flow at approximately 5.5 or 24.6 cm s-1 (Fig. 4A: P < 0.001, df = 8). Flow rates were measured prior to the experiment by photographing hydrated Artemia spp. cysts and analyzing images with

ImageJ software (Sebens and Johnson 1991). The pumps were fitted with sponge filters that were cleaned every other day. Irradiance within the tubes was measured every three days with a cosine quantum PAR sensor on a Diving-PAM sensor (Waltz, GmbH,

Effeltrich, Germany) that was calibrated with a LI-192 Underwater Quantum Sensor fitted to a LI-1400 Datalogger (LI-COR Inc., Nebraska, USA).

Experiment 4 tested for the effects of flow and PCO2 under natural light conditions.

Four pieces of each morphology were placed in each of four flumes for 14 d. Each flume created a different treatment (one flume per treatment): low flow-ambient PCO2 (LF-

ACO2), high flow-ambient PCO2 (HF-ACO2), low flow-high PCO2 (LF-HCO2), and high flow-high PCO2 (HF-HCO2). The flumes held 5.3 L of seawater (9.5 cm depth, 9.0 cm width, and 60 cm length) that poured over a lip into a 150-L reservoir tank that was filled with seawater equilibrated to the desired PCO2. Water was pumped from the reservoir to the flumes with pumps (Aqueon Quietflow Submersible Aquarium Utility Pump 1700) to

produce low and high flow rates of ~5.3 and ~17.5 cm s-1, respectively (Fig. 5B: P < 0.01, df = 59), and were measured as previously described (Sebens and Johnson 1991). During preliminary experiments the natural irradiances caused the corals to bleach, and therefore the flumes were shaded with a black mesh to reduce light to 1229 ± 54 µmol m-2 s-1 for

-2 -1 branches and 209 ± 10 µmol m s for plates (P < 0.001, df = 11). The corals were arranged in the flume in a row parallel to the direction of flow, and each day their positions were randomly changed to avoid position effects. Irradiance in the flumes were measured every three days with a cosine quantum PAR sensor fitted to a Diving-PAM, within the sensor placed in both the unshaded and shaded locations where branches and plates were placed, respectively.

Response variables

Buoyant weights of corals were measured at the beginning and end of each experiment. To determine net calcification, the difference in the initial and final buoyant weight was converted to dry weight using the density of aragonite (2.93 g cm-3, Davies,

1989) and normalized to area estimated using aluminum foil (Marsh, 1970).

Tissue biomass was assessed to test for differences between the morphologies.

During Experiment 1, after surface area was measured, the branches and plates of P. rus were fixed in formalin (5% in seawater) for 48 hours to preserve the tissue. The coral fragments were then rinsed with distilled water and placed in 10% HCl (in freshwater) to decalcify. Decalcification typically required 72 h. The preserved tissue was rinsed with distilled water and endoliths were removed with forceps. For Experiment 1, the tissue

was then placed in a 15-mL centrifuge tube with 5 mL of distilled water for homogenization using a Fisher Scientific Model 500 Ultrasonic Dismembrator fitted with a 3.2-mm diameter probe. An aliquot (1 mL) of the homogenate was dried to a constant weight at 60 °C to determine biomass, which then was normalized to surface area

(mg cm-2). For Experiment 2, 3, and 4, coral tissue was fixed in formalin (5% in filtered seawater) for 48 h and then dissolved in 10% HCl for up to 72 h. The coral tissue was cleaned of endolithic algae using forceps, rinsed in deionized water, and dried to a constant weight at 60 ºC for 48 hours. Tissue samples were weighed after 48 and 72 hours to ensure weight tissue was thoroughly dried. The difference in the initial and final weight was used to estimate biomass normalized to surface area (mg cm-2).

During Experiment 3, dark respiration of the corals also was measured. The respiration chamber was of a design modified after Patterson et al. (1981), and allowed flow to be regulated between 7.3 and 21.7 cm s-1 using a variable voltage supplied to a submersible pump (up to 1,360 L h-1). The chambers had a volume of 2.3 L and were filled with seawater from the treatment tanks in which corals were incubated. The seawater used in the system was acquired from the PCO2 equilibrated tanks where the corals were incubated. Corals were kept in darkness for 30 min to avoid stimulatory effects of light on respiration (Edmunds and Davies 1988), and dark respiration was determined at ambient temperature (28 °C) maintained by an aquarium heater and chiller.

Flow speeds were assessed as described previously. The depletion rate of oxygen was determined using oxygen optrodes (Foxy-R, 1.58 mm diameter, Ocean Optics) connected to spectrophotometers (USB2000 and NeoFox, Ocean Optics) that logged percentage

saturation onto a personal computer running Ocean Optics software (OOISensors and

NeoFox Viewer, Ocean Optics). Oxygen probes were calibrated with water-saturated air

1 (100%) and a zero solution of sodium sulfite (Na2SO3) and 0.01 mol L- sodium tetraborate (Na2B4O7). O2 saturation was maintained between 80 – 100% to avoid effects

-1 of hypoxia on respiration. O2 saturation was converted to concentration (µmol O2 mL ) using values for O2 solubility [N. Ramsing and J. Gundersen at Unisense, http://www.unisense.com/Default.aspx?ID=1109 (Garcia and Gordon, 1992)]. The rate of

O2 consumption was estimated by linear regression of O2 concentration on time, and rates were corrected for control values in respirometers filled with filtered seawater alone.

Final values were standardized to the surface area of the coral tissue and expressed in

-1 -2 -1 units of µmol O2 mL cm h .

Statistical analysis

For the physical parameters of the experiments, PCO2 treatments, and temperature, compared with tanks as replicates with a one-way ANOVA. Light in Experiment 1 and 2, and flow rates in Experiment 3 and 4, were compared with a t-test. A split-plot ANOVA was used to test for the effects of morphology, temperature, and light or flow on calcification rates and biomass in Experiment 1, 2, and 3. In these analyses, tank was the

random effect nested in PCO2 or PCO2 x temperature, morphology was the fixed, between- plot effects, and light or flow were the split plot effects. For Experiment 3, a three-way

ANOVA was used to test the effects of PCO2, morphology and flow on dark respiration.

Although Experiment 4 had one replicate per treatment and was pseudoreplicated, a one- way ANOVA was used to determine effects of each treatment (flow x PCO2) on

calcification rates and biomass of branches and plates of P. rus. Results from Experiment

4 should be considered with caution. Post-hoc Tukey HSD tests were used when interactive effects were significant in each trial to test for differences between specific pairs. Statistical analyses were performed using Systat 11. Assumptions of normality and homoscedasticity were tested by visually inspecting plotted residuals.

Results

Seawater carbonate chemistry

For Experiment 1, mean (± SE) PCO2 values in treatments were regulated for ambient conditions of 367 ± 5 to 384 ± 7 µatm (F1,156 = 1.701, P = 0.169) and elevated

PCO2 from 703 ± 16 to 720.2 ± 14 µatm (n = 4, P = 0.756) both at 28 ± 0.1°C (Table 1).

Light levels were 1007 ± 53µmol photons m-2 s-1 (mean ± SE, n = 24) in the center of each tank, and 215 ± 7 µmol photons m-2 s-1 (mean ± SE, n = 72) along the front perimeter of the tank where corals in low light conditions were placed.

The four treatments for Experiment 2 consisted of ambient temperature-ambient

PCO2 (AT-ACO2), ambient temperature-high PCO2 (AT-HCO2), high temperature-ambient

PCO2 (HT-ACO2), high temperature-high PCO2 (HT-HCO2) with 2 tanks per treatment.

PCO2 treatments were maintained at ambient and levels, with ambient PCO2 at 380 ± 3 µatm

(AT-ACO2) and 406 ± 3 µatm (HT-ACO2) and elevated PCO2 at 1108 ± 15 and 1107 ± 16

µatm (Table 4) (F1,286 = 4001.685, P < 0.001). Light intensities were 728 ± 24 µmol photons m-2 s-1 (mean ± SE, n = 8) in the center of each tank and 154 ± 4 µmol photons m-

2 s-1 (mean ± SE, n = 8) along the front perimeter of the tank where corals in low light

conditions. Mean ambient temperature was maintained at 27 °C and elevated temperature at 30 °C (F1,250 = 2570.096, P < 0.001).

For Experiment 3, the indoor flow study, mean (± SE) PCO2 values in treatments

were regulated for ambient conditions of 385 ± 3 µatm and elevated PCO2 of 965 ± 19

µatm (F1,120 = 928.00, P < 0.001). The temperature for all tanks were consistent at 28.61

± 0.03 °C (Table 7; F15,396 = 1.608, P = 0.157). Light levels were 181 ± 13 µmol photons m-2 s-1 (mean ± SE, n = 42) in the center of each acrylic tube holding the samples of P. rus for the two rates of flow, 24.6 ± 0.2 and 5.3 ± 0.3 cm s-1 (P < 0.001).

Ambient and elevated PCO2 values in treatments for Experiment 4 were maintained at targeted levels of 400 and 1000 µatm (F3,54 = 103.685, P < 0.001). Mean (± SE) PCO2 ambient conditions were 425 ± 15 µatm and 433 ± 15 µatm. Mean (± SE) PCO2 for elevated conditions were 1028 ± 33 µatm and 1088 ± 62 µatm. All experimental tanks had a temperature of 28.01 ± 0.13 °C (F3,54 = 0.892, P = 0.451). Light levels were 1230 ±

55 µmol photons m-2 s-1 (mean ± SE, n = 9) for the branches, and 209 ± 10 µmol photons m-2 s-1 (mean ± SE, n = 9) for the plates. Mean (± SE) flow rates were 17.5 ± 0.2 and 5.3 ± 0.3 (P < 0.001).

Experiment 1: Effects of light and PCO2 on coral growth

For Experiment 1, there were 4 replicates treatment-1, with 3 subsamples within each tank for the 21-day period coral samples were in PCO2 conditions of 400 and 700

µatm under high and low light conditions. Area-normalized net calcification was not

significantly affected by PCO2 for either plates or branches (Fig 2A, F1,6 = 0.241, P =

0.641) and there was no interactions among PCO2, morphology, and light (F1,60 = 0.155, P

= 0.708). However, there was an interactive effect of light and morphology for area- normalized net calcification (F1,76 = 6.081, P = 0.049). Mean (± SE) area-normalized net

2- -1 calcification of branches was 45.5% higher in high light (0.64 ± 0.05 mg CaCO3 day

-2 2- -1 -2 cm ) than low light (0.44 ± 0.04 mg CaCO3 day cm ) (Fig. 3A, Post-hoc HSD Test: P

= 0.005). Area-normalized net calcification averaged across the treatments ranged from

2- -1 -2 2- -1 -2 0.49 ± 0.07 mg CaCO3 day cm to 0.60 ± 0.05 mg CaCO3 day cm for plates, and

2- -1 -2 2- -1 -2 ranged from 0.38 ± 0.03 mg CaCO3 day cm to 0.67 ± 0.05 mg CaCO3 d cm (mean

± SE) for branches. Area-normalized biomass in branches and plates was not affected by

light or PCO2 (Fig. 2B, F1,62 = 0.074, P = 0.795 and F1,6 = 0.36, P = 0.570, respectively).

Mean biomass of branches was similar among treatments, ranging from 4.13 ± 0.4 mg cm-

2 to 4.7 ± 0.56 mg cm-2 while for plates it ranged 4.82 ± 0.56 mg cm-2 to 6.29 ± 1.43 mg cm-2 (Fig. 2B, ± SE, n=44 and 47, respectively). Biomass was 19.4% higher in plates than branches, but only marginally significant (F1,62 = 4.805, P = 0.071). Tissue biomass of plates was 5.13 ± 0.28 mg cm-2, =(± SE, n = 47) while branches were 4.12 ± 0.28 mg cm-2

(Fig 3B; ± SE, n = 47 and 44, respectively).

Experiment 2: Effects of temperature, light, and PCO2 on coral growth

In Experiment 2, calcification rates in branches and plates were not affected by the interactive effects of light, temperature, and PCO2 (Fig. 3C; F1,66 = 0.029, P = 0.874).

However, there was a marginally significant difference in calcification between morphologies. Branches had 41% slower calcification than plates (Fig. 3C; F1,66 = 37.612

P = 0.004). Mean (± SE) area-normalized calcification rates for branches were 0.70 ±

2- -1 -2 2- -1 -2 0.05 mg CaCO3 d cm and for plates were 1.19 ± 0.06 mg CaCO3 d cm . Area- normalized calcification rates of both branches and plates incubated in low light conditions were 20% less than rates from the high light conditions, but this was not significant (F1,66 = 0.260, P = 0.976). After 14 d, area-normalized biomass of branches and plates was not affected by the interactive and direct effects of light, temperature, and

PCO2 (Table 3, Fig. 3D; F4,68 = 0.249, P = 0.909). Mean (± SE) area-normalized biomass of branches did not differ among treatments, ranging from from 4.4 ± 0.5 to 5.5 ± 0.8 mg

-2 cm . Plates had 46% more biomass than branches (F1,68 = 179.361, P < 0.001). Mean (±

SE) area-normalized biomass of plates ranged from 7.3-10.8 ± 1.5-1.3 (Fig. 3D).

Experiment 3: Effects of flow and PCO2 on coral growth

For Experiment 3, mean (± SE) area-normalized calcification of branches ranged

2- -1 -2 from 0.72 ± 0.06 to 1.17 ± 0.06 mg CaCO3 d cm and plates ranged from 1.34 ± 0.09 to

2- -1 -2 1.41 ± 0.05 mg CaCO3 d cm (Fig. 5A). Branches and plates were not affected by PCO2

(F1,47 = 0.249, P =0.644) or by the interactive effects of PCO2 and flow (Table 8; F1,47 =

1.649, P = 0.268). Under ambient conditions, calcification of branches differed between

2- -1 -2 flow conditions with rates at 1.17 ± 0.06 mg CaCO3 d cm under low flow (n = 9) and

2- -1 -2 0.76 ± 0.09 mg CaCO3 d cm under high flow (F1,47 = 18.261; P = 0.001). Under

2- -1 -2 elevated PCO2, calcification of branches was 0.72 ± 0.06 mg CaCO3 d cm in high flow,

2- -1 -2 and 0.91 ± 0.04 mg CaCO3 d cm in low flow, but did not differ significantly between flows (P = 0.97). Calcification rates of plates were similar in high and low flow

2- -1 -2 treatments, with the mean (± SE) rate at 1.36 ± 0.01 mg CaCO3 d cm (P = 0.97).

There was no effect of flow and PCO2 on the biomass of branches and plates (F1,2 = 8.705,

P = 0.098). As seen in Experiment 1 and 2, biomass was marginally significantly higher in plates than in branches (Fig. 5B; F1,29 = 14.814, P = 0.061).

Mean dark respiration rates (± SE) of branches were highest in low flow, ambient

-1 -2 -1 PCO2 at 0.95 ± 0.10 µmol mL cm h (n = 5), and lowest in high flow, high PCO2 at 0.39

± 0.08 µmol mL-1 cm-2 h-1 (n = 5) (Fig. 5C; Table 9). Mean dark respiration (± SE)

-1 -2 -1 declined from low flow, ambient PCO2 at 1.26 ± 0.09 µmol mL cm h to high flow, high

-1 -2 -1 PCO2 at 0.79 ± 0.26 µmol mL cm h (Fig. 5C; Table 9). Respiration rates did not significantly differ between morphologies despite the 40% higher rates in plates compared to branches (Table. 10; F1,30 = 17.21, P < 0.001) with plates respiring at 1.01 ± 0.05 µmol

-1 -2 -1 -1 -2 -1 mL cm h and branches at 0.61 ± 0.06 µmol mL cm h . Elevated PCO2 negatively impacted respiration rates. In branches, dark respiration rates were lower in elevated PCO2 conditions in both low and high flow at 0.39 ±0.08 and 0.42 ± 0.11 µmol mL-1 cm-2 h-1

(mean ± SE), respectively than compared to low and high flow in the ambient PCO2 at 0.95

-1 -2 -1 ± 0.10 and 0.68 ± 0.06 µmol mL cm h , respectively (F1,30 = 10.900, P < 0.001).

Experiment 4: Effects of flow and PCO2 on coral growth under natural light

For Experiment 4, the effects of flow and PCO2 on branches and plates were tested separately because each morphology was under their natural light conditions. For branches, area-normalized calcification rates were similar at low-flow, ambient PCO2,

-2 high-flow, ambient PCO2, and high-low, high PCO2 and averaged at 0.38 ± 0.01 mg CaCO3 d-1 cm-2 (mean ± SE n = 4 treatment-1, P > 0.12). Mean (± SE) area calcification rates

-2 -1 were lowest for branches in low-flow, high PCO2 conditions at 0.16 ± 0.07 mg CaCO3 d

-2 cm (Fig. 6A; F3,12 = 5.10, P = 0.02). Area-normalized biomass of branches was not effected by the flow and PCO2 (Fig. 6B; F3,12 = 0.99, P = 0.43). For plates, mean (± SE) area-normalized calcification rates did not differ (Fig. 6C; F3,12 = 0.596, P = 0.630),

-2 -1 -2 -2 -1 -2 ranging from 0.27 ± 0.07 mg CaCO3 d cm to 0.42 ± 0.04 mg CaCO3 d cm .

Biomass of plates were not affected by the PCO2 and/or flow (Fig. 6D; F3,12 = 0.43, P =

0.73).

Discussion

Morphology is a fundamental feature of tropical reefs affecting structural complexity of coral colonies and reef communities. Reef morphology is an emergent property of the organisms building the reef framework and is a result largely of plasticity and physiological responses of sessile coral colonies to the environment to maximize efficiency, such as for mass transfer rates and light acquisition. The objective of this study was to test for differences in calcification rates of two contrasting skeletal morphologies exposed to OA coupled with ecologically relevant conditions of light and water flow. Porites rus is an ideal model system for this study as it is an abundant reef building coral with high morphological plasticity (Jaubert 1977) and is representative of the morphological complexity found on the reef among and within coral species (Todd

2008). Branches and plates of P. rus were used for this experiment to test the null hypothesis that calcification rates would not differ between the two morphologies in response to elevated PCO2. There was a series of 4 experiments applied to test for differences in calcification rates between contrasting morphologies exposed to OA under

different combinations of flow and light. The results of these experiments show that branches and plates of P. rus were insensitive to elevated PCO2 at 800 µatm and 1000 µatm for 2 to 3 weeks. Therefore the null hypothesis that skeletal morphology would not have an effect on calcification response to OA was not rejected.

Experiment 1 and 2 tested for differences in calcification rates in branches and plates of P. rus to elevated PCO2 in high and low light. In Experiment 1, light conditions were manipulated as a treatment within ambient and elevated PCO2 for 3-weeks from April to May 2012. In Experiment 2, ambient and elevated temperature was added to the design and 2-week incubations were run from January to February in 2013. In both experiments,

PCO2 had no interactive or direct effect on calcification and biomass of the two morphologies. Branches exhibited light-enhanced calcification (LEC) with higher calcification rates in high light than compared to low light. Plates, however, had similar calcification rates regardless of treatment, and higher biomass than branches. Branches exhibited stronger LEC than plates possibly becauseas they are more dependent on photosynthetic carbon to meet their metabolic needs than plates, which acquire their metabolic carbon from heterotrophy (Padillo-Gamiño et al. 2012).

Regardless of the temperature effects added to the experiments, calcification rates of both morphologies did not reveal differential responses to the interactive or individual effects of OA and temperature. Hermatypic corals respond to climate change and ocean acidification in a variety of ways. Edmunds et al. (2012) reported that for branches of P. rus, calcification was high under elevated temperatures (29.3 ºC) and ambient PCO2 (411

µatm), but significantly decreased when exposed to elevated PCO2 (804 µatm) at the same temperature. When P. rus was exposed to low temperatures (25.6 ºC), there were no differences in calcification despite the ambient and elevated PCO2 treatments (Edmunds et al. 2012). Some disparity in the responses of corals to OA and climate change is attributed to the susceptibility of the symbiont and the environmental history of the holobiont (Putnam et al. 2011), the symbiont composition, and types of Symbiodinium clades associated within the tissue of the coral host (Putnam et al. 2012). Corals with a specialized relationship with a dominant symbiont clade (i.e., specifist [Franklin et al.,

2012]) appear to be less sensitive to thermal stress than generalists (Putnam et al. 2012).

P. rus is considered a specifist in term of having a strong association with a dominant clade and sustains high specificity with Symbiodinium C15 (Lajuenesse et al. 2003;

Franklin et al. 2012; Padillo-Gamiño et al. 2012; Putnam et al. 2012). High fidelity of P. rus with Symbiodinium C15 may have coevolved to be successful in variable environments (van Woesik et al. 2011; Putnam et al. 2012).

The second set of experiments addressed the effects of OA and flow speeds on the calcification of branches and plates of P. rus. Skeletal morphology might be of importance as it affects mass transfer characteristics, modulating the flux of metabolites and ions adjacent to the coral tissue directly affecting coral growth and symbiont photosynthesis (Patterson 1992; Lesser et al. 1994). Flow and morphology can greatly influence the delivery and flux of metabolites necessary for holobiont respiration, coral biomineralization, and photosynthesis by the symbionts (Lesser et al. 1994). Velocity of water passing over an object and the ratio of advective to diffusive forces is described as

the Reynold’s number (Re) and is the size of the object (height in cm) multiplied by the flow speed (cm s-1) divided by the kinematic viscosity (cm2 s-1). Maintaining a Re that effectively reduces diffusion boundary layers and an optimal Sherwood’s number (Sh), which is the ratio of mass flux per unit area by water motion to diffusivity (White 1988:

Patterson 1992; Lesser et al. 1994). For instance, Pocillopora damicornis displayed morphological plasticity under different water flow environments in nature, which allowed it to achieve a constant Re that mitigated the risk of mass transfer limitations

- when [HCO3 ] was high in demand by Symbiodinium spp. for photosynthesis (Lesser et al.

1994).

2- + - In acidified conditions, [CO3 ] and Ωarag declines while [H ] and [HCO3 ] increases. Recent studies and reviews on hermatypic corals have proposed that

2- calcification is not controlled by the availability of CO3 in the water column, rather

- 2- calcification is influenced by the acquisition of HCO3 , its conversion to CO3 , and the efflux of excess [H+] from the coral tissue (Atkinson et al. 2008; Comeau et al. 2013;

Jokiel 2013; Venn et al. 2013). The processes by which hermatypic corals acquire, transport, and convert DIC species from the seawater through the tissue layers to the site of calcification in the calicoblastoderm remains unresolved. Potentially, in acidified conditions the efflux of H+ may become more energetically costly as the gradient of [H+] between inside and outside of the holobiant tissue is reduced with ocean acidification

- (Jokiel 2013; Venn et al. 2013). Increased water flow may assist in the delivery of HCO3 and the efflux of H+ to sustain calcification by the coral and photosynthesis by

Symbiodinium spp. Dissimilar coral morphologies may exhibit differential calcification

responses to OA as they are dissimilarly influenced by flow. The present study tested if branches and plates of P. rus are differentially susceptible to elevated PCO2 because of potential mass transfer limitations by reduced H+ efflux (Jokiel 2013).

Flow speeds for Experiment 3 and 4 generated different turbulent flows across branches and plates (i.e., ~5 cm s-1 and ~25 cm s-1). Branches experienced Re of ~11,327 in the high flow and ~2,265 in the low flow. Re of plates were ~2,831 in high flow and

~566 in low flow. The limitations of the experimental design caused branches and plates to experience different Re values. Regardless of the high and low flow speeds, DIC appeared not to be limiting or H+ accumulating as calcification rates were not depressed by elevated PCO2 in either branches or plates. Plates had similar calcification and dark respiration rates across PCO2 and flow treatments. Although calcification rates of branches were unaffected by PCO2, dark respiration rates were significantly lower in elevated PCO2 regardless of flow rates in Experiment 3. Unlike plates, branches reduced aerobic metabolic activity and demand for oxygen at high PCO2. Experiment 4, the outdoor experiment under natural light cycles revealed that calcification of branches were reduced in low flow, elevated PCO2 suggesting that increased flow may mitigate the negative effects of OA on branches. Sensitivity of branches to OA in low flow environments may have been more apparent under natural high light conditions because of the photosynthetic

- demand from the symbiont for CO2 from the conversion of stored HCO3 by the enzyme carbonic anhydrase (Weis et al. 1989). However, it is important to note that Experiment 4 was pseudoreplicated and further studies with replicates are needed to accurately descrbe the response of branches and plates to flow and OA under natural light conditions.

- In low flow under natural light conditions, the mass transfer of HCO3 for use in photosynthesis by Symbiodinium was potentially limited, and this could have reduced

- LEC. In high flow, there is delivery of HCO3 and O2 to the coral tissue at night that can increase photosynthesis and improve metabolic performance (Gardella and Edmunds,

1999). High flow is also necessary for removing photosynthetic inhibiting radicals, such as reactive oxygen species and hydroxyl, during the day (Yamasaki 2000; van Woesik et al. 2001). High water flow may alleviate negative impacts of OA and climate change

-2 caused by reduced pH and available CO3 (Langdon and Atkinson 2005; Marubini et al.

2008; Jokiel 2013). The understanding of how inorganic carbon is acquired by corals and delivered to the site of calcification in the calicoblastic epithelium for coral biomineralization remains underdeveloped (Allemand et al. 2011). Hermatypic corals and their symbiotic relationship with Symbiodinium adds complexity in understanding the effects of OA on calcifying organisms because of the delivery and use of DIC for calcification by the coral holobiont (Allemand et al. 2011; Jokiel 2013). The capacity to

- use increased HCO3 by Symbiodinium in hospite within corals under high light conditions may be a key trait in explaining variation in susceptibility to OA and climate change

(Comeau et al. 2013).

Previous studies have shown a variety of responses in calcification rates of P. rus upon exposure to elevated PCO2. In previous studies, P. rus was affected negatively by elevated PCO2 at 850 µatm (Edmunds et al. 2012) and ~2000 µatm (Comeau et al. 2013).

In other cases, branches of P. rus were not affected by OA conditions at ~1000 µatm PCO2

(Comeau et al. 2013). Calcification rates of P. rus branches were unaffected by OA regardless of paired autotrophic and heterotrophic treatments (Comeau et al. 2013). The variability in response of calcification of P. rus to OA conditions in previous studies and this present analysis begs for further investigation in order to identify the key traits, which leads to resistance in the face of OA. P. rus is a perforate coral with high biomass and has a strong association with the symbiont Symbiodinium C15 known to be tolerant of warming stress (Putnam et al. 2012). These characteristics may contribute to P. rus being a “winner” (Fabricius et al. 2011) as environmental conditions, such as increased SST and

OA, are expected to worsen.

Scleractinian diversity supports topographical structure at the scale of whole reefs, which is critical in sustaining ecosystem function, services, and high biodiversity (Connell

1978). As ocean acidification and climate change are expected to continue (Hoegh-

Gulberg et al. 2007), these press disturbances will contribute to reduce percent cover of scleractinians. Individual species and functional groups of scleractinian morphologies, are anticipated to differ in their capacity to acclimatize to rapid changes in seawater carbonate chemistry and SST (Hofmann et al. 2010). There could be substantial reduction in species richness and structure of reefs (Knowlton and Jackson 2008; Anthony et al. 2011).

Enormous progress has been made in understanding and describing scleractinian tolerances to OA (Fabricius et al. 2011; Comeau et al. 2013; Shamberger et al. 2014).

However, little is known about the response of coral traits such as morphological plasticity and acclimation capacity to identifying important contribute to coral tolerance to changing environments. Further studies are necessary to explore relative susceptibilities

of corals with contrasting corallum morphologies to OA. A comparative study of multiple species with high phenotypic plasticity would be valuable in determining if selection of corals with different morphologies can be a useful tool for improved conservation and the management of corals faced with climate change and ocean acidification.

Tables

Table 1. Summary of carbonate chemistry (n=160) and light (n=20) conditions in Experiment 1 where 8 tanks assigned randomly to create four treatments of ACO2 = ambient CO2 and HCO2 = high CO2 for 21 d. Within each of the 6 tanks, there were high and low irradiance levels. Mean ± SE.

Treatment T Sal pH TA PCO2 ΩArag Light Irradiance -1 -2 -1 (°C) (µmol kg ) (µatm) Treatment (µmol m s )

ACO2 28 36.10 8.07 2335 ± 1 374 ± 3 3.97 ± 0.02 High 1017 ± 20

HCO2 28 36.10 7.84 2332 ± 1 712 ± 8 2.63 ± 0.02 Low 216 ± 5

Table 2. Results of the 3-Way Split Plot ANOVA for Experiment 1 comparing area-normalized calcification (mg cm-2 d-1) to determine the effects of light and PCO2 on the branched and plated morphologies of Porites rus. Tanks were nested within each PCO2 treatment. Bold indicates significance at α = 0.05.

Source SS df MS F-Ratio P-Value MS Denominator

Between CO2 0.020 1 0.02 0.241 0.641 Tank(CO2)

Plots Tank(CO2) 0.502 6 0.084 1.465 0.205 Residual

Morphology 0.069 1 0.069 1.259 0.305 Morph x Tank(CO2)

Light 0.171 1 0.171 3.223 0.123 Light x Tank(CO2)

Morphology x Light 0.326 1 0.326 6.081 0.049 Morphology x Light x Tank(CO2)

Morphology x CO2 0.000 1 0 0 0.995 Morphology x Tank(CO2)

Within Light x CO2 0.112 1 0.112 2.096 0.198 Light x Tank(CO2)

Plots Morphology x Light x CO2 0.008 1 0.008 0.155 0.708 Morphology x Light x Tank(CO2)

Morphology x Tank(CO2) 0.327 6 0.054 1.596 0.292 Morphology x Light x Tank(CO2)

Light x Tank(CO2) 0.322 6 0.054 1.983 0.213 Morphology x Light x Tank(CO2)

Morphology x Light x Tank(CO2) 0.318 6 0.053 0.928 0.482 Residual Residual 3.428 60 0.057

Table 3. Results of the 3-Way Split Plot ANOVA for Experiment 1 comparing area-normalized tissue biomass (mg cm-2) to determine the effects of light and PCO2 on the branched and plated morphologies of Porites rus. Tanks were nested within each PCO2 treatment. Bold indicates significance at α = 0.05.

Source SS df MS F-Ratio P-Value MS Denominator

Between CO2 2.976 1 2.976 0.36 0.570 Tank(CO2) Plots Tank(CO2) 49.613 6 8.269 1.332 0.257 Residual

Morphology 13.862 1 13.862 4.805 0.071 Morph x Tank(CO2)

Light 0.139 1 0.139 0.074 0.795 Light x Tank(CO2)

Morphology x Light 1.096 1 1.096 0.172 0.693 Morphology x Light x Tank(CO2)

Morphology x CO2 1.323 1 1.323 0.459 0.524 Morphology x Tank(CO2)

Within Light x CO2 0.092 1 0.092 0.049 0.832 Light x Tank(CO2)

Plots Morphology x Light x CO2 1.344 1 1.344 0.211 0.662 Morphology x Light x Tank(CO2)

Morphology x Tank(CO2) 17.31 6 2.885 0.453 0.921 Morphology x Light x Tank(CO2)

Light x Tank(CO2) 11.26 6 1.877 0.295 0.919 Morphology x Light x Tank(CO2)

Morphology x Light x Tank(CO2) 38.178 6 6.363 1.025 0.418 Residual Residual 384.898 62 6.208

Table 4. Summary of the carbonate chemistry (n=72) and light (n=8) conditions applied in Experiment 2 where 8 tanks were assigned randomly to create four treatments AT-CO2 = ambient temperature ambient CO2, HT-ACO2 = high temperature ambient CO2, AT-

HCO2 = ambient temperature high CO2, HT-HCO2 = high temperature high CO2 for 21d. Mean ± SE.

Treatment T Sal pH TA PCO2 ΩArag Light Irradiance (°C) (µmol kg-1) (µatm) Treatment (µmol m-2 s-1)

AT-ACO2 27.32 ± 0.04 35.29 ± 0.81 8.06 2313 ± 2 380 ± 3 3.78 ± 0.02

High 728 ± 24

HT-ACO2 29.82 ± 0.05 35.28 ± 0.78 8.03 2317± 2 406 ± 3 3.92 ± 0.02

AT-HCO2 27.05 ± 0.05 35.29 ± 0.81 7.67 2316 ± 1 1108 ± 15 1.79 ± 0.02

Low 153 ± 4

HT-HCO2 29.80 ± 0.04 35.34 ± 0.09 7.67 2318 ± 2 1107 ± 16 1.99 ± 0.02

Table 5. Reseults of the 4-Way Split Plot ANOVA for Experiment 2 comparing area-normalized calcification (mg cm-2 d-1) to determine the effects of light, temperature, and PCO2 on the branched and plated morphologies of Porites rus. Tanks were nested within each PCO2 x temperature treatment. Bold indicates significance at α = 0.05.

Source SS df MS F-ratio P-value MS Denominator

Between CO2 0.090 1 0.09 1.102 0.353 Tank(Temp x CO2) Plots Temp 0.331 1 0.331 4.055 0.114 Tank(Temp x CO2)

Tank(Temp x CO2) 0.326 4 0.082 0.664 0.619 Residual

Morphology 5.334 1 5.334 37.612 0.004 Morphology x Tank(Temp x CO2)

Light 1.201 1 1.201 5.017 0.089 Light x Tank(Temp x CO2)

Light x Morphology 0.009 1 0.009 0.280 0.976 Morphology x Light x Tank(Temp x CO2)

Morphology x CO2 0.010 1 0.010 0.072 0.802 Morphology x Tank(Temp x CO2) Light x CO 0.194 1 0.194 0.811 0.419 Light x Tank(Temp x CO ) Within 2 2 Plots Light x Morphology x CO2 0.096 1 0.096 0.285 0.622 Morphology x Light x Tank(Temp x CO2) Morphology x Temp 0.052 1 0.052 0.364 0.579 Morphology x Tank(Temp x CO2)

Light x Temp 0.101 1 0.101 0.715 0.446 Light x Tank(Temp x CO2)

Morphology x Light x Temp 0.01 1 0.010 0.029 0.874 Morphoogy x Light x Tank(Temp x CO2)

Morphology x Tank(Temp x CO ) 0.567 4 0.142 0.420 0.789 Morphology x Light x Tank(Temp x CO ) 2 2 Light x Tank(Temperature x CO2) 0.958 4 0.239 0.709 0.627 Morphology x Light x Tank(Temp x CO2) Morphology x Light x Tank(Temp x CO2) 1.351 4 0.338 2.753 0.035 Residual Error 8.101 66 0.123

Table 6. Results of the 4-Way Split Plot ANOVA for Experiment 2 comparing area-normalized tissue biomass (mg cm-2) to determine the effects of light, temperature, and PCO2 on the branched and plated morphologies of Porites rus. Tanks were nested within each

PCO2 x temperature treatment. Bold indicates significance at α = 0.05.

Source SS df MS F-ratio P-value MS Denominator CO 0.055 1 0.055 0.006 0.942 Tank(Temp x CO ) Between 2 2 Plots Temp 9.158 1 9.158 1.002 0.374 Tank(Temp x CO2) Residual Tank(Temp x CO2) 36.567 4 9.142 1.048 0.389 427.19 Morphology 427.191 1 1 179.361 < 0.001 Morphology x Tank(Temp x CO2)

Light 1.023 1 1.023 0.39 0.566 Light x Tank(Temp x CO2)

Light x Morphology 1.035 1 1.035 0.476 0.528 Morphology x Light x Tank(Temp x CO2) Within Plots Morphology x CO2 0.497 1 0.497 0.209 0.671 Morphology x Tank(Temp x CO2) Light x CO2 5.846 1 5.846 2.229 0.21 Light x Tank(Temp x CO2)

Light x Morphology x CO2 1,388 1 1.399 0.643 0.467 Morphology x Light x Tank(Temp x CO2) Morphology x Temp 0.312 1 0.312 0.131 0.736 Morphology x Tank(Temp x CO ) 2 Light x Temp 9.494 1 9.494 3.62 0.13 Light x Tank(Temp x CO2) Morphology x Light x Temp 16.310 1 16.31 7.498 0.052 Morphoogy x Light x Tank(Temp x CO2)

Morphology x Tank(Temp x CO2) 9.527 4 2.382 1.206 0.43 Morphology x Light x Tank(Temp x CO2)

Light x Tank(Temperature x CO ) 10.492 4 2.623 0.39 0.566 Morphology x Light x Tank(Temp x CO ) 2 2 Residual Morphology x Light x Tank(Temp x CO2) 8.701 4 2.175 0.249 0.909 Error 593.12 68 8.722

Table 7. Summary of carbonate chemistry (n=222) of Experiment 3, flow (n=30) and light (n = 6) conditions used inside the

mesocosm set up with 6 tanks, 3 tanks for each PCO2 treatment – ACO2 = ambient CO2 and HCO2 = high CO2 for 21d. Mean ± SE.

Flow Treatment T pH A P Ω Light Irradiance Sal T CO2 Arag Flow Rate (°C) (µmol kg-1) (µatm) Treatment (µmol m-2 s-1) (cm s-1)

28.63 ± 35.37 ± 3.84 ± ACO 8.07 2305 ± 2 385 ± 3 High 25 ± 0.4 2 0.04 0.03 0.02

Light 181 ± 13

28.66 ± 35.37 ± 2.11 ± HCO 7.73 2304 ± 2 965 ± 19 Low 6 ± 0.3 2 0.04 0.03 0.03

Table 8. Results of the 4-Way Split Plot ANOVA for Experiment 3 comparing area-normalized calcification (mg cm-2 d-1) to determine the effects of flow and PCO2 on the branched and plated morphologies of Porites rus. Tanks were nested within each PCO2 treatment. Bold indicates significance at α = 0.05.

Source SS df MS F-ratio P-value MS Denominator

Between CO2 0.028 1 0.028 0.249 0.644 Tank(CO2) Plots Tank(CO2) 0.448 4 0.112 1.419 0.242 Residual

Flow 0.608 1 0.608 4.27 0.108 Flow x Tank(CO2)

Morphology 4.219 1 4.219 73.202 0.001 Morphology x Tank(CO2)

Morphology x Flow 0.374 1 0.374 18.261 0.013 Morphology x Flow x Tank(CO2)

Morphology x CO2 0.095 0.095 1.649 0.268 Morphology x Tank(CO2)

Within Flow x CO2 0.056 1 0.056 0.396 0.563 Flow x Tank(CO2)

Plots Flow x Morphology x CO2 0.142 1 0.142 6.94 0.058 Morphology x Flow x Tank(CO2)

Flow x Tank(CO2) 0.57 4 0.143 6.953 0.043 Morphology x Flow x Tank(CO2)

Morphology x Tank(CO2) 0.231 4 0.058 2.813 0.17 Morphology x Flow x Tank(CO2)

Morphology x Flow x Tank(CO2) 0.082 4 0.02 0.26 0.902 Residual Error 3.706 47 0.079

Table 9. Results of the 4-Way Split Plot ANOVA for Experiment 3 comparing area-normalized biomass (mg cm-2) to determine the effects of flow and PCO2 on the branched and plated morphologies of Porites rus. Tanks were nested within each PCO2 treatment.

Source SS df MS F-ratio P-value MS Denominator

Between CO2 150.14 1 150.14 8.705 0.098 Tank(CO2) Plots Tank(CO2) 34.393 2 17.247 0.386 0.683 Residual

Flow 25.566 1 25.566 0.535 0.536 Flow x Tank(CO2)

Morphology 889.792 1 889.792 14.814 0.061 Morphology x Tank(CO2)

Morphology x Flow 53.526 53.526 0.593 0.522 Morphology x Flow x Tank(CO2)

Morphology x CO2 28.871 1 28.871 0.481 0.56 Morphology x Tank(CO2)

Within Flow x CO2 0.027 1 0.027 0.001 0.983 Flow x Tank(CO2)

Plots Flow x Morphology x CO2 0.013 1 0.013 < 0.001 0.992 Morphology x Flow x Tank(CO2)

Flow x Tank(CO2) 87.804 2 43.902 0.486 0.673 Morphology x Flow x Tank(CO2)

Morphology x Tank(CO2) 120.129 2 60.064 0.665 0.601 Morphology x Flow x Tank(CO2)

Morphology x Flow x Tank(CO2) 180.654 2 90.327 2.021 0.151 Residual Error 1295.843 29 44.683

Table 10. Results of the 3-Way ANOVA for Experiment 3 comparing dark respiration rates (µmol mL-1 cm-2 h-1) to determine the effects of flow and PCO2 on the branched and plated morphologies of Porites rus. Bold indicates significance at α = 0.05.

Source SS df MS F-ratio P-value

CO2 1.03 1 1.03 10.90 < 0.001 Morphology 1.53 1 1.53 16.21 < 0.001 Flow 0.34 1 0.34 3.56 0.07 Morphology x CO2 0.07 1 0.07 0.71 0.41 Flow x CO2 0.05 1 0.05 0.55 0.47 Flow x Morphology 0.01 1 0.01 0.14 0.71 Flow x CO2 x Morphology 0.02 1 0.02 0.21 0.65 Error 2.84 30 0.09

Table 11. Summary of carbonate chemistry (n=28), flow (n=30), and light (n=12) conditions used for the 4 flumes (low flow-ambient

PCO2 [LF-ACO2], high flow-ambient PCO2 [HF-ACO2], low flow-high PCO2 [LF-HCO2], high flow-high PCO2 [HF-HCO2]) in each flume exposed to the natural light cycles for 14 d for Experiment 4. Mean ± SE.

Flow Treatment T Sal pH A P Ω Flow Light Irradiance T CO2 Arag Rate (°C) (µmol kg-1) (µatm) Treatment (µmol m-2 s-1) (cm s-1)

28.09 35.81 ± HF-ACO 8.06 2317 ± 6 425 ± 15 3.65 ± 0.06 2 ± 0.15 0.12 17.50 ± High High 1230 ± 54.5 0.2 28.34 35.71 ± LF-ACO 8.06 2309 ± 4 433 ± 15 3.62 ± 0.07 2 ± 0.12 0.11

27.69 35.73 ± 1028 ± HF-HCO 7.75 2325 ± 8 1.97 ± 0.05 2 ± 0.50 0.12 33 5.34 ± Low Low 209 ± 10.0 0.3 27.88 35.86 ± 1088 ± LF-HCO 7.71 2308 ± 6 1.92 ± 0.11 2 ± 0.11 0.14 62

Table 12. Results of area-normalized calcification from the One Way ANOVA of each treatment (low flow-ambient PCO2 [LF-ACO2], high flow-ambient PCO2 [HF-ACO2], low flow-high PCO2 [LF-HCO2], high flow-high PCO2 [HF-HCO2]) in each flume (n = 1).

Branched (A) and plated (B) morphologies of Porites rus were analyzed separately because they were exposed to different irradiance levels. Bold indicates significance at α = 0.05.

A. Branched Source SS df MS F-ratio P-value Treatment 0.152 3 0.051 5.104 0.017 Error 0.119 12 0.010 B. Plated Source SS df MS F-ratio P-value Treatment 0.063 3 0.021 0.596 0.630 Error 0.422 12 0.035

Table 13. Results of area-normalized biomass from the One Way ANOVA of each treatment (low flow-ambient PCO2 [LF-ACO2], high flow-ambient PCO2 [HF-ACO2], low flow-high PCO2 [LF-HCO2], high flow-high PCO2 [HF-HCO2]) in each flume (n = 1).

Branched (A) and plated (B) of Porites rus were analyzed separately because they were exposed to different irradiance levels.

A. Branched Source SS df MS F-ratio P-value Treatment 7.104 3 2.368 0.992 0.430 Error 28.659 12 2.388 B. Plated Source SS df MS F-ratio P-value Treatment 17.282 3 5.760 0.434 0.733 Error 159.443 12 13.287

Figures

-2 -1 Figure 1. Irradiance (µmol photons m s ) measured over 3 d in 2012 in tanks

(experimental conditions of high light [Exp-High] and low light [Exp-Low]) and in the natural environment where branches and plates of P. rus were collected at 0.5 and 2 m depth, respectively (high light [Field-High] and low light [Field-Low]) with MkV-L logging light sensors (JFE Advantech Co., Kobe, Japan) that measured PAR with a 4-π spherical quantum PAR sensor).

Figure 2. The flow-through system with maintained seawater carbonate chemistry in twelve 150 L tanks with independently controlled temperature, light, and CO2 for

Experiments 1, 2, and 3 (A). Seawater equilibrated from this system was also used for

Experiment 4. Black mesh was attached to the aquarium covers of each tank for shading

-2 -1 purposes to produce contrasting light conditions of ~158 and ~758 µmol photons m s for low and high light conditions for Experiment 2 (B).

Figure 3. Mean (± SE) area-normalized calcification (mg cm-2 d-1) and tissue biomass

(mg cm-2) of branches (light grey) and plates (dark grey) of Porites rus from Experiment

1 (A and B) and Experiment 2 (C and D). Branches and plates were incubated under high and low irradiance in ambient (400 µatm, ACO2) and elevated (700 µatm, HCO2) PCO2 for

Experiment 1. In experiment two, corals were incubated under ambient temperature- ambient PCO2 (AT-ACO2), high temperature-ambient PCO2 (HT-ACO2), ambient temperature-high PCO2 (AT-HCO2), and high temperature-high PCO2 (HT-HCO2). The number of replicates is shown within each bar (n=8-12 for Experiment 1 and 5-6 for

Experiment 2).

Figure 4. Experimental set up to test for the effects of flow and PCO2 on branches and plates of Porites rus. For Experiment 3 three samples of branches and three plates were placed inside clear acrylic tubes with aquarium pumps producing low and high flow rates

(A). Experiment 4 tested for the effects of flow and PCO2 under natural light conditions, four samples of branches and plates were placed in each of the four flumes. The flumes were placed over a 150L reservoir tank with seawater being equilibrated at experimental

PCO2. Water was pumped from the reservoir to the flumes with submersible pumps to produce low and high flow rates (B).

Figure 5. Mean (± SE) area-normalized calcification (mg cm-2 d-1) (A), area-normalized biomass (mg cm-2) (B), and dark respiration (µmol mL-1 cm-2 h-1) (C) of branches (light grey bars) and plates (dark grey bars) of Porites rus in Experiment 3. Branches and plates were exposed to low and high flow conditions in ambient (400 µatm) and elevated

(1000 µatm) PCO2 conditions (low flow-ambient PCO2 [LF-ACO2], high flow-ambient

PCO2 [HF-ACO2], low flow-high PCO2 [LF-HCO2], and high flow-high PCO12 [HF-HCO2].

The number of replicates is indicated within each bar.

Figure 6. Mean (± SE) area-normalized calcification (mg cm-2 d-1) and area-normalized biomass (mg cm-2) of branches (light grey bars) (A and B) and plates (dark grey bars) (C and D) of Porites rus. This study tested the effects of flow and PCO2 on branched and plated morphologies under natural light conditions. Branches and plates were placed in low flow-ambient PCO2 (LF-ACO2), high flow-ambient PCO2, low flow-high PCO2 (LF-

HCO2), and high flow-high PCO2 (HF-HCO2). Number of replicates is indicated in each bar.

Figure 1.

2800 Exp. High Exp. Low Field High 2400 Field Low

) 2000 -1 s -2 m 1600

1200 Irradiance

800 (µmol photons

400

0 27 April 28 April 29 April

Date

Figure 2.

100

Figure 3.

101

Figure 4.

102

Figure 5.

103

Figure 6.

104

Chapter 4

Concluding Remarks

Hermatypic corals are central in maintaining the structural integrity, high biodiversity, and ecosystem services in tropical coral reef ecosystems (Connell 1978;

Moberg and Folke 1999). However the corals that create the structural foundation for tropical reef communities are weakened in strength and declining in cover from increasing threats posed by multiple disturbances (Hughes 1994; Alvarez-Filip et al.

2009). Natural disturbances such as hurricanes and crown-of-thorn seastar (Acanthaster planci) outbreaks have been long driven the inherent cycles in coral populations.

However, anthropogenic activity, such as fossil fuel use, has greatly exacerbated the negative effects of natural disturbances in eroding reef resilience (De’ath et al. 2012).

The increase in greenhouse gas emissions has augmented the occurrence of coral bleaching resulting from increased sea surface temperature (Hoegh-Guldberg 1999; Van

Woesik et al. 2011) and reduced calcification rates from acidified oceans (Hoegh-

Guldberg et al. 2007; Comeau et al. 2013). In recent decades, hermatypic coral communities have changed in species composition (Bellwood et al. 2004) and declined in cover at high rates worldwide as a result of sequential disturbance eventsm, and there are few signs of successful recovery (Gardner et al. 2003; Bruno and Selig 2007; De’ath et al. 2012; Ruzicka et al. 2013; Jackson et al. 2014). As these disturbances persist, it is important to determine how reef communities and structure will continue to change.

While hermatypic corals in shallow reefs have declined in cover, it is important to assess how reef communities and abundances of other benthic taxa, such as ascidians,

105

sponges, gorgonians are also being affected by such disturbances. Alterations in reef communities and relative abundance of benthic sessile organisms can change the function and ecological trajectory of coral reefs (Norström et al. 2009; Rossi et al. 2012; Rossi

2013). While shallow reef corals of the Caribbean have declined in coral cover from numerous disturbances such as yellow and white band disease outbreaks (Holden 1996;

Patterson et al. 2002; Aronson and Precht 2001), hurricanes (Woodley et al. 1981; Blair et al. 1994), large-scale bleaching (Glynn 1993; Lesser 2011), and competition with macroalgae (Jompa and McCook 2003), taxa such as gorgonians (Ruzicka et al. 2013;

Inoue et al. 2013), sponges (Lesser 2006; Pawlik 2011), and ascidians (Norström et al.

2009) may have traits allowing their populations to be resistant to the same stressors.

Relative to studies on scleractinians in the Caribbean, little is known about other reef taxa beyond macroalgae and stony corals (Norström et al. 2009).

For this study, I assessed changes in gorgonian abundance in St. John, US Virgin

Islands, and throughout the Caribbean to describe abundances of benthic taxa, aside from macroalgae and scleractinians. My study demonstrates that gorgonians have been able to persist and increase in abundance despite the numerous factors that have caused scleractinians to decline in abundance (Edmunds 2013; Ruzicka et al. 2013). The persistence of gorgonians in a changing environment may provide a three-dimensional habitat for shallow reef organisms. However, these alternative reefs will unlikely equal the skeletal structure and ecological function that scleractinians have long provided in supporting complex tropical coral reef ecosystems. It should also be noted that while gorgonians appear abundant and less affected than scleractinians by thermal stress and

106

hurricanes, this might be a short-term deferral to the long-term devastation of reefs if disturbances continue to escalate from poor mitigation of carbon emissions and protection of reefs.

Enormous efforts have been made to understand and rank the varied responses of scleractinian calcification to future climate change and acidified conditions. In this study, calcification rates of branches and plates of Porites rus were measured to determine how morphology of scleractinians will be affected in future oceans impacted by acidification and increased temperature. P. rus is abundant along fringing reef environments exposed to high sedimentation and run off in Moorea, French Polynesia (Padillo-Gamiño et al.

2012). Although P. rus is structurally delicate and colonies can have high turnover rates, this coral has ecological value in providing microhabitats for numerous reef invertebrates and vertebrates (Holbrook et al. 2002; Brooks et al. 2007). Branching and “weedy” corals are considered to be more vulnerable to climate change and ocean acidification than massive, slow growing corals (Comeau et al. 2013). However, in this study P. rus was generally unaffected by acidification or interactive effects with light, flow, and temperature. Differential responses in calcification between the contrasting morphologies of P. rus were not detected. It is important for future OA studies to consider morphology of hermatypic corals to assess how reef structure will change with increasing abiotic pressures. Such studies can identify tolerances within morphological groups and further explore the capacity of acclimatization for sessile organisms with high phenotypic plasticity subjected to stochastic environments.

107

Hermatypic corals that once thrived in tropical coral reef ecosystems have declined in abundance and reefs are projected to undergo continual reductions in biodiversity and topographical complexity (Jones et al. 2007). This study highlights current and potential changes in tropical reef communities. In particular, the increase in the abundance of gorgonians in the Caribbean and resistance of P. rus morphologies to

OA conditions in Moorea, French Polynesia, both draw attention to the hardy fauna that may persist in threatened habitats. These cnidarians are examples of “winning” taxa that have high recruitment, fast growth, and physiological tolerance against climate change conditions. Although it is good news that there are benthic taxa robust to disturbances, the next “generation” of reef communities may be greatly impaired by the loss of species richness and durable calcium carbonate structure. Communities comprised of gorgonians or scleractinians like P. rus will be young communities, with higher turn over rates with little long-term investment into durable structural reefs. Future studies are required to understand physiological thresholds of these “tough” organisms. Moreover, further research needs to encourage effective local and global management to mitigate disturbances that cause organisms to perform at the edge of their biological thresholds.

108

Literature Cited

Alcolado, P.M., García-Parrado, P., Hernández-Muñoz, D. 2008. Estructura y composición de las comunidades de gorgonias de los arrecifes del Archipiélago Sabana- Camagüey, Cuba: conectividad y factores determinantes. Nol. Invest. Mar. Cost. 37: 11- 29.

Allemand, D., È, Tambutté, D. Zoccola, and S. Tambutté. 2011. Coral calcification: cells to reefs. Coral Reefs: An Ecosystem in Transition. 119-150.

Alvarez-Filip, L., N. Dulkvy, I. Cote, A. Waktinson, and J. Gill. 2011. Coral identity underpins architectural complexity on Caribbean reefs. Ecol. App. 21: 2223-2231.

Alvarez-Filip, L., Dulvy, N.K., Gill, J.A., Coˆte, I.M., and Watkinson, A.R. 2009. Flattening of Caribbean coral reefs: region-wide declines in architectural complexity. P. Roy. Soc. B. Biol. 276: 3019–3025.

Anthony, K., D. Kline, G. Diaz-Pulido, S. Dove, and O. Hoegh-Gulberg. Ocean acidification cause bleaching and productivity loss in coral reef builders. P. Natl. A. Sci. 105: 17442-17446.

Aronson, R.B. and Precht, W.F. 2001. White-band disease and the changing face of Caribbean coral reefs. The Ecology and Etiology of Newly Emerging Marine Diseases. pp. 25-38. Springer Netherlands.

Atkinson, M.J. and Cuet, P. 2008. Possible effects of ocean acidification on coral reef biogeochemistry: topics for research. Mar. Ecol. Prog. Ser. 373: 249-256.

Baskin, Y. 1998.Winners and losers in a changing world. BioSci. 48: 788-792.

Bellwood, D.R., Hughes, T.P., Folke, C., and Nyström, M. 2004. Confronting the coral reef crisis. Nature. 429: 827-833.

Bertness, M. and Callaway, R. 1994. Positive interactions in communities. TREE. 9: 191-193.

Blair, S.M., McIntosh, T.L., and Mostkoff, B.J. 1994. Impacts of Hurricane Andrew on the Offshore Reef Systems of Central and Northern Dade County, Florida. B. Mar. Sci. 54: 961-973.

Bond, Z.A., Cohen, A.L., Smith, S.R., and Jenkins, W.J. 2005. Growth and composition of high-Mg calcite in the skeleton of a Bermudian gorgonian (Plexaurella dichotoma): Potential paleothermometry. Geochem. Geophys. Geosy. 6: 1-10.

109

Botero, L. 1987. Zonación de octocorales gorgonáceos en el area de Santa Marta y Parque Nacional Tayrona, costa Caribe colombiana. An. Inst. Inv. Mar. Punta de Betín. 17: 60-61.

Brazea, D.A. and Lasker, H.R. Growth rates and growth strategy in a clonal marine invertebrate, the Caribbean octocoral Briareum asbestinum. Biol. Bull. 183: 269-277.

Brooks, A.J., Holbrook, S.J., and Schmitt, R.J. 2007. Patterns of microhabitat use by fishes in the patch-foring coral Porites rus. Raffles Bull. Zool. 14: 245-254.

Brown, D.J. and Edmunds, P.J. 2013. Long-term changes in the population dynamics of the Caribbean hydrocoral Millepora spp. J. Exp. Mar. Biol. Ecol. 441: 62-70.

Bruno, J. and Edmunds. P.J. 1997. Clonal variation for phenotypic plasticity in the coral Madracis mirabilis. Ecology. 28: 2177-2190.

Bruno, J., Petes, L.E., Harvell, C.D., and Hettinger, A. 2003. Nutrient enrichment can increase the severity of coral diseases. Ecol. Lett. 12: 1056-1061.

Cesar, H.J.S., Burke, L., and Pet-Soede, L. 2003. The Economics of Worldwide Coral Reef Degradation. Cesar Environmental Economics Consulting, Arnhem, and WWF Netherlands, Zeist, The Netherlands. 23 pp.

Chappell, J. 1980. Coral morphology, diversity, and reef growth. Nature. 286:249-252.

Chiappone, M. and Sullivan, K.M. 1994. Ecological structure and dynamics of nearshore hard-bottom communities in the Florida Keys. B. Mar. Sci. 54: 747-756.

Chiappone, M., Geraldes, F., Rodriguez, Y., and Vega, M. 2001. Sedimentation as an important environmental influence on Domincan Republic reefs. B. Mar. Sci. 69: 805- 818.

Chiappone, M., Dienes, H., Swandson, D., and Miller, S. 2003. Density and gorgonian host-occupation patterns by flamingo tongue snails. Caribb. J. Sci.. 39: 116-127.

Cohen, A.L. and Holcomb, M. 2009. Why corals care about ocean acidification, uncovering the mechanism. Oceanography. 22: 118-127.

Coffroth, M.A. and Lasker, H.R. 1998. Population structure of a clonal gorgonian coral: the interplay between clonal reproduction and disturbance. Evolution. 379-393.

Colvard, N.B. and Edmunds, P.J. 2007. Decadal-scale changes in abundance of non scleractinian invertebrates on a Caribbean coral reef. J. Exp. Mar. Biol. Ecol. 397: 153 160.

110

Comeau, S., Edmunds, P.J., Spindel, N.B., Carpenter, R.C. 2013. The responses of eight coral reef calcifiers to increasing partial pressure of CO2 do not exhibit a tipping point. Limnol. Oceanogr. 58: 388-398.

Comeau, S., Carpenter, R.C., and Edmunds, P.J. 2013. Coral reef calcifiers buffer their response to ocean acidification using both bicarbonate and carbonate. P. Roy. Soc. B. Biol. 280: 20122374

Comeau, S., Carpenter, R.C., and Edmunds, P.J. 2013. Effects of feeding and light intensity on the responses of the coral Porites rus to ocean acidification. Mar. Biol. 160: 1127-1134.

Connell, J.H. 1978. Diversity in tropical rain forests and coral reefs. Science 199:1302- 1310.

Connell, J.H. 1997. Disturbance and recovery of coral assemblages. J. Exp. Mar. Biol. Ecol. 209: 261-278.

Crawley, A., D. Kline, S. Dunn, K. Anthony, and S. Dove. 2010. The effect of ocean acidification on symbiont photorespiration and productivity in Acropora formosa. Glob. Change Biol. 16: 851-863

Dahlgren, EJ. 1989. Gorgonian community structure and reef zonation patterns on Yucatan coral reefs. Bull. Mar. Sci. 45: 678-696.

Dahlgren, EJ. 2002. Gorgonian distribution patterns in coral reef environmentsof the Gulf of Mexico: evidence of sporadic ecological connectivity? Coral Reefs. 21: 205- 215.

Davies P. S. 1989 Short-term growth measurements of corals using an accurate buoyant weighing technique. Mar. Biol. 101: 389–395.

De Nys, R., Coll, J.C., and Price, I.R. 1991. Chemically mediated interactions between the red alga Plocanium namatum (Rhodophyta) and the octocoral Sinularia cuciatu (Alcyonacea). Mar. Biol. 108: 315-320.

Diaz, M.C. and Rützler, K. 2001. Sponges: an essential component of Caribbean coral reefs. Bull. Mar. Sci. 69: 535-546.

Dickson, A. C. Sabine, and J. Christian [eds.] 2007. Guide to best practices for ocean CO2 measurements. PICES Special Publication 3.

Doney, S.C., Fabry, V.J., Feely, R.A., and Kleypas, J.A. 2009. Ocean acidification: the other CO2 problem. Mar. Sci. 1: 169-192.

111

Dufault, A.M., Ninokawak, A., Bramanti, L., Cumbo, V.R., Fan, T. Y., and Edmunds, P.J. 2013. The role of light in mediating the effects of ocean acidification on coral calcification. J. Exp. Biol. 216: 1570-1577.

Dustan, P., and Halas, J.C. 1987. Changes in the reef-coral community of Carysfort, Reef, Key Largo, Florida: 1974 to 1982. Coral Reefs. 6: 91-106.

Edmunds, P.J. 2002. Long-term dynamics of coral reefs in St. John, US Virgin Islands. Coral Reefs. 21: 357-367.

Edmunds, P.J. 2007. Evidence for decadal-scale decline in the growth rtes of juvenile scleractinian corals. Mar. Ecol. Prog. Ser. 341: 1-13.

Edmunds, P. 2011. Zooplanktivory ameliorates the effects of ocean acidification on the reef coral Porites spp. Limnol. Oceanogr. 56: 2402-2410.

Edmunds, P.J. 2013. Decadal-scale changes in the community structure o coral reefs of St. John, US Virgin Islands. Mar. Ecol. Prog. Ser. 489: 107-123.

Edmunds, P.J., Aronson, RB., Swanson, DW., Levitan, DR., and Precht, WF. 1998. Photographic versus visual census techniques for the quantification of juvnile corals. Bull. Mar. Sci. 62: 937-946.

Edmunds, P.J., Brown, D.B., and Moriarty, V.M. 2012. Interactive effects of ocean acidification and temerpature on two scleractinian corals from Moorea, French Polynesia. Glob. Change Biol. 18: 2173-2183.

Edmunds, P.J. and Elahi, R. 2007. The demographics of a 15-year decline in cover of the Caribbean reef coral Montastraea annularis. Ecol. Monogr. 77: 3-18.

Enriquez, S., Méndez, E.R., Iglesias-Prieto, R. 2005. Multiple scattering on coral skeletons enhances light absorption by symbiotic algae. Limnol. Oceanogr. 50: 1025- 1032.

Etnoyer, P.J., Wirshing, H.H., and Sánchez, J.A. 2010. Rapid assessment of octocoral diversity and habitat on Saba Bank, Netherlands Antilles. PloS One. 5: e10668.

Erez, J., S. Reynaud, J. Silverman, K. Scheider, and D. Allemand. 2011. Coral calcification under ocean acidification and global change in Z. Dubinsky. N. Stambler (Eds) Coral Reefs: An Ecosystem in Transition. Springer Science, New York. 151-176.

112

Fabricus, K., C. Langdon, S. Uthicke, C. Humphrey, A. Noonan, G. De’ath, R. Okazaki, N. Muehllehner, M. Glas, and J. Lough. 2011. Losers and winners in coral reefs acclimatized to elevated carbon dioxide concentrations. Nature Clim. Change.1: 165-169.

Fabry, V., B. Seibel, R. Feely, and J. Orr. 2011. On the potential for ocean acidification to be a general cause of ancient reef crises. Glob. Change Biol. 17: 56-67.

Falkowski, P. and Z. Dubinsky. 1981. Light-shade adaptation of Stylophora pistillata, a hermatypic coral from the Gulf of Eilat. Nature. 289: 172-174

Falkowski, P., Z. Dubinsky, L. Muscatine, and J. Porter. 1984. Light and the bioenergetics of a symbiotic coral. BioScience. 34: 705-709.

Feely, R.A., Doney, S.C., Cooley, S.R. 2009. Ocean acidification: present conditions and future changes in a high-CO2 world. Oceanography. 22: 36-47.

Fernández, L.H., Bermejo, M.G., Barreto, R.A., and Alonso, L.C. 2011. Composición de las comunidades de octocorales y corales pétreos y la incidencia del blanqueamiento del 2005 en Jardines de la Reina, Cuba. Revista Ciencias Marinas Costeras. 3: 77-90.

Ferrier-Pagès, C. M. Googenboom, and F. Houlbrèque. 2010. The role of plankton in coral trophodynamics, p. 215-229. In Z. Dubinsky and N. Stambler [eds.], Coral reefs: An ecosystem in transition. Springer.

Foster, A. 1979. Phenotypic plasticity in the reef corals Montastaea annularis (Ellis and Solander) and Siderastrea siderea (Ellis and Solander). J. Exp. Mar. Biol. Ecol. 39: 25-54.

Franklin, E.C., Stat, M., Pochon, X., Putnam, H.M., and Gates, R.D. 2012. GeoSymbio: a hybrid, cloud-based web application of global geospatial bioinformatics and ecoinformatics for Symbiodinium-host symbioses. Mol. Ecol. Res. 12:369-373

Fujioka, Y. (1999) Mass destruction of the hermatypic corals during a bleaching event in Ishigaki Island, Southwestern Japan. Galaxea. JCRS. 1: 41-50.

Gardner, T., Côté, I.M., Gill, J.A., Grant A., and Watkinson, A.R. 2003. Long-term region-wide declines in Caribbean corals. Science. 301: 958-960.

Gardner, T. Côté, I.M., Gill, J.A., Grant, A., and Watkinson, A.R. 2005. Hurricanes and Caribbean coral reefs: impacts, recovery patterns, and role in long-term decline. Ecology. 86: 174-184.

Gates, R.D., Baghadasarian, G., and Muscatine, L. 1992. Temperature stress causes host cell detachment in symbiotic cnidarians: implications for coral bleaching. Biol. Bull. 182: 324-332.

113

Gates, R.D. and P. Edmunds. 1999. The physiological mechanisms of acclimatization in tropical reef corals. Amer. Zool. 39: 30-43.

Gattuso, J., M. Frankignoulle, I. Bourge, S. Romaine, and R. Buddemeier. 1998. Effect of calcium carbonate saturation of seawater on coral calcification. Global Planet.Change. 18: 37-46.

Gienapp, P., C. Teplitsky, J. Alho, J. Mills, and J. Merilä. 2008. Climate change and evolution: disentangling environmental and genetic responses. Mol. Ecol. 17: 167-178.

Godinot, C., C. Ferrier-Pagès, and R. Grover. 2009. Control of phosphate uptake by zooxanthellae and host cells in the scleractinian coral Stylophora pistillata. Limnol. Oceanogr. 54: 1627-1633.

Goulet, T.L., LaJeunesse, T.C., and Fabricius, K.E. 2008. Symbiont specificity and bleaching susceptibility among soft corals in the 1998 Great Barrier Reef mass coral bleaching event. Mar. Biol. 154: 795-804.

Gledhill, D.K., Wanninkhof, R., Millero, F.J., and Eakin, M. 2008. Ocean acidification of the greater Caribbean region 1996-2006. J. Geo Research. 113: C10031.

Goldenheim, W.M., and Edmunds, P.J. 2011. Effects of flow and temperature on growth and photophysiology of scleractinian corals in Moorea, French Polynesia. Biol. Bull. 221: 270-279.

Goldberg, W.M. 1973. The ecology of the coral-octocoral communities off the southeast Florida coast: geomorphology, species composition, and zonation. Bull. Mar. Sci. 23: 465-488.

Goreau, T.F. 1964. Mass expulsion of zooxanthellae from Jamaican reef communities after Hurricane Florida. Science. 145: 383-386.

Green, D.H., Edmunds, P.J., and Carpenter, R.C. 2008. Increasing relative abundance of Porites astreoides on Caribbean reefs mediated by an overall decline in coral cover. Mar. Ecol. Prog. Ser. 359: 1-10.

Grottoli, A.G., Rodrigues, L.J., and Palardy, J.E. 2006. Heterotrophic plasticity and resilience in bleached corals. Nature. 440: 1186-1189.

Helmuth, B.S.T., Sebens, K.P., and Daniel, T.L. 1997. Morphological variation in coral aggregation: branch spacing and mass flux to coral tissues. J. Exp. Mar. Biol. Ecol. 209: 233-259.

114

Hoegh-Guldberg, O., P. Mumby, A. Hooten, R. Steneck, P. Greenfield, E. Gomez, C. Harvell, P. Sale, A. Edwards, K.Caldeira, N. Knowlton, C. Eakin, R. Iglesias-Prieto, N. Muthiga, R. Bradbury, A. Dubo, and M. Hatziolos. 2007. Coral reefs under rapid climate change and ocean acidfication. Science. 318: 1737-1742.

Hofmann, G.E., Barry, J.P., Edmunds, P.J., Gates, R.D., Hutchins, D.A., Klinger, T., and Sewell, M.A. 2010. The effect of ocean acidification on calcifying organisms in marine ecosystems: an organism-to-ecosystem perspective. Ann. Rev. Ecol. Evol. Syst. 41: 127-147.

Hofmann, G.E., Smith, J.E., Johnson, K.S., Send, U., Levin, L.A., Micheli, F., Paytan, A., Price, N.N., Peterson, B., Takeshita, Y., Matson, P.G., Crook, E.D., Kroeker, K.J. et al. 2011. High-frequency dynamics of ocean pH: a multi-ecosystem comparison. PlosONE. 6: e28983

Holbrook, S.J., Brooks, A.J., and Schmitt, R.J. 2003. Variation in structural attributes of patch-forming corals and in patterns of abundance of associated fishes. Mar. Fresh. Research. 53: 1045-1053.

Holden, C. 1996. Coral disease hot spots in the Florida Keys. Science. 274: 778-795.

Hönisch, B. A. Ridgwell, D. Schmidt, E. Thomas, S. Gibbs, A. Sluijs, L. Kump, R. Martindale, S. Greene, W. Kiessling, J. Ries, J. Zachos, D. Royer, S. Barker, T. Marchitto, Jr., R. Moyer, C. Pelejero, P. Ziveri, G. Foster, and B.Williams. 2012. The geological record of ocean acidification. Science. 335: 1058-1063.

Hoogenboom, M., S. Connolly, and K. Anthony. 2008. Interactions between morphological and physiological plasticity optimize energy acquisition in corals. Ecology. 89: 1144- 1154.

Holden, C. 1996. Coral disease hot spot in the Florida Keys. Science. 274: 776-795.

Hughes, T.P. 1994. Catastrophes, phase shifts, and large-scale degredation of a Caribbean coral reef. Science. 265: 1547-1551.

Hughes, T.P. and Tanner, J.E. 2000. Recruitment failure, life histories, and long-term decline of Caribbean corals. Ecology. 81: 2250-2263.

Idjadi, J.A. and Edmunds, P.J. 2006. Scleractinian corals as facilitators for other invertebrates on a Caribbean reef. MEPS. 319: 117-127.

Iglesias-Prieto, R., V. Beltrán, T. LaJeunesse, H. Reyes-Bonilla, and P. Thomé. 2004. Different algal symbionts explain the vertical distribution of dominant reef corals in the eastern Pacific. Proc. Royal Soc. 271: 1757-1763.

115

Inoue, S., Kayanne, H., Yamamoto, S., and Kurihara, H. 2013. Spatial community shift from hard to soft corals in acidified water. Nature Climate Change. 3: 683-687.

Jackson, J.B.C. 1979. Morphological strategies of sessile animals. Biology and systematics of colonial organisms. Academic Press, London. 499-555.

Jackson et al. 2014. Status and trends of Caribbean Coral Reefs: 1970-2012. Ed. Jackson, J.B.C, Donovan, M., Cramer, K., and Lam, V. GCRMN.

Jackson, J.B.C., Kirby, M.X., Berger, W.H., Bjorndal, K.A., Botsford, L.W., Bourque, B.J., Bradbury, R.H. et al. 2001. Science. 293: 629-637.

Jaubert, J. 1977. Light, metabolism, and growth forms of the hermatypic scleractinian Synaraea convexa (Verrill) in the lagoon of Moorea (French Polynesia). Proc. 4rd Int. Coral Reef Symp. 1: 483-488.

Jeng, M.S., Huang, H.D., Hsiao, Y.C., and Benayahu, Y. 2011. Sclerite calcification and reef-building in the fleshy octocoral genus Sinularia (Octocorallia: Alcyonacea). Zool. Stud. Env. Biol. Fish. 50: 457-462.

Jaubert, J. 1977. Light, metabolism, and growth forms of the hermatypic scleractinian coral Synarea convexa Verill in the lagoon of Moorea (French Polynesia). P. Third Int. Coral Reef Symp. 483-488.

Jokiel, P.L. 2013. Coral reef calcification: carbonate, bicarbonate and proton flux under conditions of increasing ocean acidification. P. Roy. Soc. B. Biol. 280: 1471-2954.

Jompa, J. and McCook, L.J. 2003. Coral-algal competition: macroalgae with different properties have different effects on corals. Mar. Ecol. Prog. Ser. 258: 87-95.

Jones, C., J. Lawton, and M. Shachak. 1997. Positive and negative effects of organisms as physical ecosystem engineers. Ecology. 78: 1946-1957.

Jones, P., T. Osborn, and K. Briffa. 2001. The evolution of climate over the last millennium. Science. 292: 662-667. 86: 1704-1714.

Jones, R.J., Hoegh-Guldberg, O., Larkum, A.W.D., and Schreiber, U. 1998. Temperature-induced bleaching of corals begins with the impairment of the CO2 fixation mechanism in zooxanthellae. Plant Cell Environ. 21: 1219-1230.

Kaandorp, J.A. 1999. Morphological analysis of growth forms of branching marine sessile organisms along environmental gradients. Mar. Biol. 134: 295-306.

Keck, J. Unpublished, MS Thesis 2004. Changes in coral populations on the northwest coast of Roatán, Bay Islands, Honduras, subsequent to 1998 bleaching event and Hurricane Mitch.

116

Kelly, M. and Hofmann, G.E. 2011. Adaptation and the physiology of ocean acidification. Funct. Ecol. 27: 980-990.

Kinzie, R.A. 1973. Coral reef project – paper in memory of Dr. Thomas F. Goreau. 5. The zonation of West Indian gorgonians. Bull. Mar. Sci. 23: 93-155.

Kleypas, J.A, McManus, J., and Meñez, L. 1999. Environmental limits to coral reef development; Where do we draw the line? Amer. Zool. 39: 146-159.

Kleypas, J.A., Buddemeier, R.W., Gattuso, A.D., Langdon, C., and Opdyle, B.N. 1999. Geochemical consequences of increased atmospheric carbon dioxide on coral reefs. Science. 284: 118-120.

Kleypas, J. Buddemeier, R.W., Gattuso, J.P. 2001. The future of coral reefs in an age of global change. Int. J. Earth Sci. 90: 426-437.

Kline, D.I., Teneva, L., Schneider, K., Niard, T., Chai, A., Marker, M. Headley, K. et al. 2012. A short-term in situ CO2 enrichment experiment on Heron Island (GBR). Sci. Reports. 2: 1-9.

Knowlton, N. 2001. The future of coral reefs. P. Natl. Acad. Sci. 98: 5419-5425

Knowlton, N. and Jackson, J.B.C. 2008. Shifting baselines, local impacts, and global change on coral reefs. PloS Biol. 6: e54.

Koehl, M.A.R. 1984. How do benthic organisms withstand moving water? Amer. Zool. 24: 57-70.

Koehl, M.A.R. 1996. When does morphology matter? Ann. Rev. Ecol. Syst. 27: 501- 542.

Krief, S. E. Hendy, M. Fine, R. Yam, A. Meibom, G. Foster, and A. Shemesh. 2010. Physiological and isotopic responses of scleractinian corals to ocean acidification. Geo et Cosmo Acta. 74: 5988-5001.

Kim, K., Harvell, C.D., Kim, P.D., Smith, G.W., and Merkel, S.M. 2000. Fungal disease resistance of Caribbean sea fan corals (Gorgonia spp.) Mar. Biol. 136: 259-267.

Kroeker, K.J., Kordas, R.L., Crim, R.N., and Singh, G.G. 2010. Meta-analysis reveals negative yet variable effects of ocean acidification on marine organisms. Ecol. Lett. 13: 1419-1434.

117

Kroeker, K.J., Kordas, R.L., Crim, R., Hendriks, I.E., Ramajo, L., Singh, G.S., Duarte, C.M., and Gattuso, J.P. 2013. Impacts of ocean acidification on marine organisms: quantifying sensitivies and interactions with warming. Glob. Change Biol. 19: 1884- 1894.

Kuffner, I.B. Grober-Dunsmore, R., Brock, JC, and Hickey, T.D. 2010. Biological community structure on patch reefs in Biscayne National Park, FL, USA. Environmental monitoring and assessment. Coral Reefs. 164: 513-531.

Lajuenesse, T.C., Loh, T.C., van Woesik, R., Hoegh-Guldberg, O., Schmidt, G.W., and Fitt, W.K. 2003. Low symbiont diversity in souther Great Barrier Reef corals, relative to those in the Caribbean. Limnol. Oceangr. 48: 2046-2054.

Langdon, C. and Atkinson, M.J. 2005. Effect of elevated pCO2 on photosynthesis and calcification of corals and interactions with seasonal change in temperatureirradiance and nutrient enrichment. J. Geophys. Res. 110: 1978-2012.

Lasker, H.R. 2005. Gorgonian mortality during a thermal event in the Bahamas. Bull. Mar. Sci. 76: 155-162.

Lasker, H.R and Coffroth, M.A. 1983. Octocoral distributions at Carrie Bow Cay, Belize. Mar. Ecol. Prog. Ser. 13: 21-28.

Lasker, H.R., Peters, E.C., and Coffroth, M.A. 1984. Bleaching of reef coelenterates in the San Blas Islands, Panama. Coral Reefs. 3: 183-190.

Lasker, H.R. 2003. Zooxanthellae densities within a Caribbean octocoral during bleaching and non-bleaching years. Coral Reefs. 22: 23-26.

Lasker, H.R., Boller, M.L., Castanaro, J., and Sánchez, J.A. 2003. Determinate growth and modularity in a gorgonian octocoral. Biol. Bull. 205: 319-330.

Lesser, M.P. 2006. Benthic-pelagic coupling on coral reefs: feeding and growth of Caribbean sponges. J. Exp. Mar. Biol. Ecol. 328: 277-288.

Lesser, M.P., Weis, V., Patterson, M., and Jokiel, J. 1994. Effects of morphology and water motion on carbon delivery and productivity in the reef coral, Pocillopora damicornis: diffusion barriers, inorganic carbon limitation, and biochemical plasticity. J. Exp. Mar. Biol. Ecol. 178: 153-179.

Lessios, H.A., Cubit J.D., Robertson, D.R., Shulman, H.J., Parker, M.R., Garrity, S.D., and Levings, S.C. 1984. Mass mortality of Diadema antillarum on the Caribbean Coast of Panama. Coral Reefs. 3: 173-182.

Lewis, C.L. and Coffroth, M.A. 2004. The acquisition of exogenous algal symbionts by an octocoral after bleaching. Science. 304: 1490-1492.

118

Lirman, D. 2001. Competition between macroalgae and corals: effects of herbivore exclusion and increased algal biomass on coral survivorship and growth. Coral Reefs. 19: 392-399.

Loh, T.L. and Pawlik, J.R. 2014. From the cover: chemical defenses and resources trade- offs structure sponge communities on Caribbean coral reefs. Proc. Nat. Acad. Sci. 10.1073/pnas.

Loya, Y., K. Sakai, K. Yamazato, Y. Nakano, H. Sambali, and R. van Woesik. 2001. Coral bleaching: the winners and losers. Ecol. Lett. 4: 122-131.

Lybolt, M.J. Unpublished. MS Thesis 2003. Count or pointcount: Is percent octocoral cover an adequate proxy for octocoral abundance?

Marsh, J.A. 1970. Primary productivity of reef-building calcareous red algae. Ecology. 51: 255-263.

Marubini, F., Ferrier-Pagès, C., Furla, P., and Allemand, D. 2008. Coral calcification responds to seawater acidification: a working hypothesis towars a physiological mechanism. Coral Reefs. 27: 491-499.

Moberg, F. and C. Folke. 1999. Ecological goods and services of coral reef ecosystems. Ecol. Econ. 29: 215-233.

Muko, S., K. Kawasaki, F. Takasu, and N. Shigesada. 2000. Morphological plasticity in the coral Porites sillimaniani and its adaptive significance. Bull. Mar. Sci. 66: 224-239.

Mullen, K.M., Peters, E.C., Harvell, C.D. 2004. Coral resistance to disease. Coral health and disease. Springer Berlin Heidelberg. 377-399.

Mumby, P.J. 2009. Phase shifts and community stability of macroalgal communities on Caribbean coral reefs. Coral Reefs. 28: 761-773.

Munday, P.L., Warner, R.R., Monro, K., Pandolfi, J.M., and Marshal, D.J. 2013. Predicting evolutionary responses to climate change in the sea. Ecol. Letters. 16: 1488- 1500.

Muscatine, L. 1980. Productivity of zooxanthellae. Primary Prod. In the Sea. 281-402. Orr, J., V. Fabry, O. Aumont, L. Bopp, S. Doney, R. Feely, A. Gnandesikan, N. Gruber, A. Ishida, F. Joos, R. Key, L. Lindsay, E. Maier-Reimer. R. Matear, R. Monfray, A. Mouchet, R. Najjar, G. Plattner, K. Rodgers, C. Sabine, J. Sarmiento, R. Schlitzer, R. Slater, I. Totterdell, M. Weirig, Y. Yamanaka, A. Yool. 2005. Anthropogenic ocean acidification over the twenty-first century and its impacts on calcifying organisms. Nature. 437: 681-686.

119

Muscatine, L. 1990. The role of symbiotic algae in carbon and energy flux in reef corals. Ecosystems of the world. 25: 75-87.

Muscatine, L., Falkowski, P.G., Porter, J.W., and Dubinsky, Z. 1984. Fate of photosynthetic fixed carbon in light-and shade-adapted colonies of the symbiotic coral Stylophora pistillata. P. Roy. Soc. Lond. B. Bio. 222:181-202.

Muscatine, L., McCloskey, L.R., and Marian, R.E. 1981. Estimating the daily contribution of carbon from zooxanthellae to coral animal respiration. Limnol Oceanogr. 26:601-611.

Nakamura, T. and van Woesik, R. 2001. Water-flow rates and passive diffusion partially explain differential survival of corals during the 1998 bleaching event. Mar. Ecol. Prog. Ser. 212: 301-304.

Nakamura, T. and Yamasaki, H. 2005. Requirement of water-flow for sustainable growth of Pocilloporid corals during temperature periods. Mar. Pol. Bull. 50: 1115-1120.

NOAA. Coral Reef Evaluation and Monitoring Project (2012) Florida Fish and Wildlife Commission/Fish and Wildlife Research Institute. http://myfwc.com/research/habitat/coral

Nyström, M., Folke, C., and Moberg, F. 2000. Coral reef disturbance and resilience in a human-dominated environment. Trends Ecol. Evol. 15: 413-417.

Norström, A.V., Nyström, M., Lokrantz, J., and Folke, C. 2009. Alternative states on coral reefs: beyond coral-macroalgal phase shifts. Mar. Ecol. Prog. Ser. 376: 295-306.

Opresko D.M. 1973. Abundance and distribution of shallow-water gorgonians in the area of Miami, Florida. Bull. Mar. Sci. 23: 535-558.

Opresko, D.M. 1974. Recolonization and regrowth of a population of the gorgonian Plexaura homomalla. Stud. Trop. Oceanogr. Miami. 12: 101-110.

Orr, J.C., Fabry, V.J., Aumont, O., Bopp, L., Doney, S.C., Feely, R.A., Gnanadeskian, A., et al. 2005. Anthropogenic ocean acidification over the twenty-first century and its impact on calcifying organisms. Nature. 437: 681-686.

Padillo-Gamiño, J.L., Hanson, K.M., Stat, M., and Gates, R. 2012. Phenotypic plasticity of the coral Porites rus: Acclimatization. J. Exp. Mar. Bio. Ecol. 434: 71-80.

Patterson, K.L., Porter, J.W., Ritchie, K.M., Polson, S.W., Mueller, E., Peters, E.C., Santavy, D.L., and Smith, G.W. 2002. The etiology of white pox, a lethal disease of the Caribbean elkhorn coral, Acropora palmata. Proc. Natl. Acad. Sci. 99: 8725-8730.

120

Patterson, M. 1992. A chemical engineering view of cnidarian symbioses. Amer. Zool. 32: 566-582.

Pawlik, J.R., Burch, M.T., and Fenical, W. 1987. Patterns of chemical defense among Caribbean gorgonian corals: a preliminary survey. J. Exp. Mar. Biol. Ecol. 108: 55-66.

Pearse, V.B. and Muscatine, L. 1971. Role of symbiotic algae (zooxanthellae) in coral calcification. Biol. Bull. 141: 350-363.

Perry, C.T., Murphy, G.N., Kench, P.S., Smithers, S.G., Edinger, E.N., Steneck, R.S., and Mumby, P.J. 2013. Caribbean-wide decline in carbonate production threatens coral reef growth. Nat. Commun. 4: 1402.

Pespeni, M.H., Sanford, E., Gaylord, B., Hill, T.M., Hosfelt, J.D., Jaris, H.K., LaVigne, M., Lenz, E.A., Russell, A.D., Young, M.K., and Palumbi, S.R. 2013. Evolutionary change during experimental ocean acidification. P. Natl. Acad. Soc. 11: 6937-6942.

Putnam H. and P. Edmunds. 2011. The physiological response of reef corals to diel fluctuations in seawater temperature. J. Exp. Mar. Biol. Ecol. 396: 216-223.

Putnam, H., Stat, M., Pochon, X., and Gates, R.D. 2012. Endosymbiotic flexibility associates with environmental sensitivity in scleractinian corals. Proc. R. Soc. B. 279: 4352-4361

Preston, E.M. and Preston, P.J. 1975. Ecological structure in a West Indian Gorgonian Fauna. Bull. Mar. Sci. 25: 248-258.

Quinn, G. and M. Keough. 2002. Experimental design and data analysis for biologists. Cambridge University Press.

Reusch, T.B.H. 2013. Climate change in the oceans: evolutionary versus phenotypically plastic responses of marine animals and plants. Evol. App. 7: 104-122

Ridgwell, A. and D. Schmidt. 2010. Past constraints on the vulnerability of marine calcifiers to massive carbon dioxide release. Lett. Nat. GeoSci. 3: 196-200.

Ries, J.B., Cohen, A.L., and McCorkle, D.C. 2009. Marine calcifiers exhibit mixed responses to CO2 induced ocean acidification. Geology. 37: 1131-1134.

Rogers C.S., McLain, L.N., and Tobias, C.R. 1991. Effects of Hurricane Hugo (1989) on a coral reef in St. John, USVI. Mar. Ecol. Prog. Ser. 78: 189-199.

Rogers, C.S. and Beet, J. 2001. Degradation of marine ecosystems and decline of fishery resources in marine protected areas in the US Virgin Islands. Environ. Conserv. 28: 312-322.

121

Rogers C.S. and Miller J. 2006. Permanent ‘phase shifts’ or reversible declines in coral cover? Lack of recovery of two coral reefs in St. John, US Virgin Islands. Mar. Ecol. Prog. Ser. 306: 103-114.

Rogers, C.S., Miller, J., NS Muller, E.M. 2008. Coral diseases following massive bleaching in 2005 cause 60 percent decline in coral cover and mortality of the threatened species, Acropora palmate, on reefs in the US Virgin Islands. USGS Fact Sheet. 3058p.

Rossi, R. Bramanti, L., Broglio, E., and Gili, J.M. 2012. Trophic impact of long-lived species indicated by population dynamics in the short-lived hydrozoan Eudendrium racemosum. Mar. Ecol. Prog. Ser. 84: 97-111.

Rossi, S. 2013. The destruction of the ‘animal forests’ in the oceans: towards an over- simplification of the benthic ecosystems. Ocean & Coastal Manag. 84: 77-85.

Ruzicka, R.R., Colella, M.A., and Porter, J.W. 2013. Temporal changes in benthic assemblages of Florida Keys reefs 11 years after the 1997/1998 El Niño. Mar. Ecol. Prog. Ser. 489: 125-141.

Sabine, C.L., Feely, R.A., Gruber, N., Key, R.M., Lee, K., Bullister, J.L., Wanninkhof, R., Wong, C.S., Wallace, D.W.R., Tilbrook, B., Millero, F.J., Peng, T.H., Kozyr, A., Ono, T., and Rios, A.F. 2004. The oceanic sink for anthropogenic CO2. Science. 305: 367-371.

Sánchez, J.A., Díaz, J.M., and Zea, S. 1997. Gorgonian communities in two contrasting environments on oceanic atolls of the Southwestern Caribbean. Bull. Mar. Sci. 61: 453- 465.

Sánchez, J.A., Zea, S., and Díaz, J.M. 1998. Patterns of octocoral and black coral distribution in the oceanic barrier reef-complex of Providencia Island, Southwestern Caribbean. Caribb. J. Sci. 34: 240-264.

Sánchez, J.A. 1999. Black coral-octocoral distribution patterns on Imedla Bank, a deep- water reef, Colombia, Caribbean Sea. Bull. Mar. Sci. 65: 215-225.

Sebens, K.P. and Johnson, A.S. 1991. Effects of water movement on prey capture and distribution of reef corals. Hydrobiologia. 226: 91-101.

Schutte, V.G.W., Selig, E.R., and Bruno, J.F. 2010. Regional spatio-temporal trends in Caribbean coral reef benthic communities. Mar. Ecol. Prog. Ser. 402: 115-122.

Shamberger, K.E.F, Cohen, A.L., Golbuu, Y., McCorkle, D.C., Lentz, S., and Barkley, H.C. 2014. Diverse coral communities in naturally acidified waters of a Western Pacific reef. Geophys. Res. Lett. 41: 499-504.

122

Suggett, D.J., Dong, L.F., Lawson, T., Lawrenze, E., Torres, L., and Smith, D.J. 2013. Light availability determines susceptibility of reef building to ocean acidification. Coral Reefs. 32: 327-337.

Sullivan, K.M., Chiappone, M., and Ninnes, C. 1996. Characteristics of hard-bottom assemblages for resource mapping of the Caicos Bank. Proceedings of the 44th Gulf and Caribbean Fisheries Institute, St. Croix. 44: 143-158.

Sunday, J.M., Calosi, P., Dupont, S., Munday, P.L., Stillman, J., and Reusch, T.B.H. 2014. Evolution in an acidifying ocean. Trends Ecol. Evol. 29: 117-125.

Tambutté, S., M. Holcomb, C. Ferrier-Pagès, S. Reynaud, É. Tambutté , D. Zoccola, and D. Allemand. 2011. Coral biomineralization: from the gene to the environment. J. Exp. Mar. Biol. Ecol. 408: 58-78.

Tanner, J.E. 1995. Competition between scleractinian corals and macroalgae: An experimental investigation of coral growth, survival and reproduction. J. Exp. Mar. Biol. Ecol. 190: 151-168.

Todd, P. 2008. Morphological plasticity in scleractinian corals. Biol. Rev. 83: 315-337.

Torres, R., Chiappone, M., Geraldes, F., Rodriguez, Y., and Vega, M. 2001. Sedimentation as an important environmental influence on Domincan Republic reefs. Bull. Mar. Sci. 69: 805-818.

United Nations Educational, Scientific and Cultural Organization: Coastal region and small island paper 3. Caribbean Coastal Marine Productivity Program Data Report 1992- 1995. van Woesik, R., Sakai, K., Ganase, A., and Loya, Y. 2011. Revisiting the winners and the losers a decade after coral bleaching. Mar. Ecol. Prog. Ser. 434: 67-76.

Vermeij, M.J.A. and Bak, R.P.M. 2002. How are coral populations structured by light? Marine light regimes an the distribution of Madracis. Mar. Ecol. Prog. Ser. 233: 104-116.

Veron, J. 1986. Corals of the World. Vol. 3. Australia: Australian Institute of Marine Sciences and CRR Qld Pty. Ltd.

Wahle, C.M. 1985. Habitat-related patterns of injury and mortality among Jamaican gorgonians. Bull. Mar. Sci. 37: 905-927.

Wall, C.B. and Edmunds, P.J. 2013. In situ effects if low pH and elevated HCO3- on juvenile Massive Porites spp. in Moorea, French Polynesia. Biol. Bull. 225: 92-101.

123

West, J.M. 1997. Plasticity in the sclerites of a gorgonian coral: tests of water motion, light level, and damage cues. Biol. Bull. 192: 279-289.

West, J.M. 1998. The dual roles of sclerites in the gorognian coral: conflicting functions of support and defense. Evol. Ecol. 12: 803-821.

Wild C., Hoegh-Guldberg O., Naumann M.S., Colombo-Pallotta M.F., Ateweberhan M., Fitt W.K., Iglesias-Prieto R., Palmer C., Bythell J.C., Ortiz J.C., Loya Y., van Woesik R. 2011. Climate change impedes scleractinian corals as primary reef ecosystem engineers. Mar. Fresh. Res. 62: 205-215.

Witman, J.D. 1992. Physical disturbance and community structure of exposed and protected reefs: a case study from St. John, US Virgin Islands. Amer. Zool. 32: 641-654.

Woodley, J.D., Chornesky, E.A., Cliffo, P.A., Jackson, J.B.C., Kaufman, L.S., Knowlton, N., et al. Hurricane Allen’s Impact on Jamaican Coral Reef. Science. 214: 749-755.

Wulff, J.L. 2006. Rapid diversity and abundance decline in a Caribbean coral reef sponge community. Biol Conserv. 127: 167-176.

Yoshioka, P.M., and Yoshioka, B.B. 1989. A multispecies, multiscale analysis of spatial pattern and its application to a shallow-water gorgonian community. Mar. Ecol. Prog. Ser. 54: 257-264.

Yoshioka, P.M. and Yoshioka, B.B. 1991. A comparison of the survivorship and growth of shallow-water gorgonian species of Puerto Rico. Mar Ecol. Prog. Ser. 69: 253-260.

Yoshioka, P.M. 2005. Biotic neighborhoods of shallow-water gorgonians of Puerto Rico. Bull. Mar. Sci. 76: 625-636.

Zeebe, R. 2012. History of seawater carbonate chemistry, atmospheric CO2, and ocean acidification. Ann. Rev. Earth Planet. Sci. 40: 141-165.

124