Walsh and Bowers 1971)

The University of Maine DigitalCommons@UMaine

Honors College

Spring 5-2018

A Histological Assessment of the Effects of Elevated Temperature and Nitrogen on the Symbiodinium of Palythoa Toxica (Walsh and Bowers 1971)

Molly Westbrook University of Maine

Follow this and additional works at: https://digitalcommons.library.umaine.edu/honors

Part of the Marine Biology Commons

Recommended Citation Westbrook, Molly, "A Histological Assessment of the Effects of Elevated Temperature and Nitrogen on the Symbiodinium of Palythoa Toxica (Walsh and Bowers 1971)" (2018). Honors College. 360. https://digitalcommons.library.umaine.edu/honors/360

This Honors Thesis is brought to you for free and open access by DigitalCommons@UMaine. It has been accepted for inclusion in Honors College by an authorized administrator of DigitalCommons@UMaine. For more information, please contact [email protected].

A HISTOLOGICAL ASSESSMENT OF THE EFFECTS OF ELEVATED

TEMPERATURE AND NITROGEN ON THE SYMBIODINIUM OF PALYTHOA

TOXICA (WALSH AND BOWERS 1971)

Molly Westbrook

A Thesis Submitted in Partial Fulfillment of the Requirements for a Degree with Honors (Marine Science)

The Honors College

University of Maine

May 2018

Advisory Committee: Dr. Ian Bricknell, Professor of Aquaculture Dr. Rhian Waller, Associate Professor of Marine Sciences Dr. Seth Tyler, Professor of Zoology, Cooperating Professor of Marine Sciences Dr. Heather Hamlin, Associate Professor of Aquaculture Dr. François Amar, Dean of the University of Maine Honors College, Professor of Chemistry

ABSTRACT

Coral reefs around the world are suffering mass bleaching events caused by a combination of stressors, including rising ocean temperatures, acidity, pollution, increased suspended sediments, and increasing nitrogen levels. Corals harbor a complex microbial ecosystem consisting of bacteria, and algal symbionts known as Symbiodinium.

This study examines the effects of elevated temperature, a known cause of bleaching, and elevated nitrogen, an increasingly important potential stressor for reefs, on the

Symbiodinium of a zoanthid coral, Palythoa toxica. A total of 65 polyps were subjected to

5 different water treatments of 2 levels of elevated nitrogen, and 2 levels of elevated temperature in populations of 12. Samples were removed every four days and at the experiment’s termination. The samples were fixed for histological examination.

Symbiodinium size was analyzed using Feret diameter. The results of this experiment confirm the effect of elevated temperature as a bleaching mechanism for P. toxica and provide evidence that elevated nitrate is a potential bleaching trigger in prolonged exposure. Furthermore, there is evidence that combined elevated nitrate and temperature compound bleaching mechanisms, with the highest amount of bleaching occurring in the most stressed treatments. When exposed to low levels of nitrate and low levels of temperature elevation, there is a trend of Symbiodinium Feret diameter increase. With time, however, there is an eventual collapse of the system, resulting in Symbiodinium

Feret diameter decrease and an increase of algal mass potentially from cell lysis within bleached Symbiodinium colonies remaining between tentacles and within the coelenteron.

ACKNOWLEDGEMENTS

Thank you to Dawna Beane for histological fixation of slides, Robert Harrington for system design and construction help, Scarlett Tudor for statistical analysis and results guidance, Jeremy Rich for microscope use, Corinne Mooring and the Bricknell laboratory team of summer 2017 for system care, and Ashley Rossin for methodology of size- frequency analysis. The author would also like to graciously thank Rhian Waller, Seth

Tyler, Heather Hamlin and François Amar for their knowledge, resources, and invaluable input to this project. Finally, the author would like to express their endless gratitude to

Ian Bricknell for helping to make this project a reality, and providing guidance and enthusiasm at every step. Thank you.

iii

TABLE OF CONTENTS

Table of Figures v Introduction 1 Anthropogenic Effects on Coral Reefs 1 Symbiodinium and the Adaptive Bleaching Hypothesis 2 The Interaction Between Nitrogen, Coral, and Symbiodinium 5 Palythoa toxica Overview and Study Objectives 6 Materials and Methods 8 System Design and Sampling 8 Symbiodinium Cell Size Analysis 10 Statistical Analysis 11 Results 12 Experiment One 12 Effects of Elevated Nitrogen 14 Effects of Elevated Temperature 17 Variance Among Treatments 20 Experiment Two 21 Effects of Combined Elevated Nitrogen and Temperature 22 Variance Among Treatments 27 Discussion 28 Nitrogen Effects on P. toxica 28 Temperature Effects on P. toxica 30 Combined Stress Effects 31 Conclusions 33 References 35 Appendices 40 Appendix 1: ImageJ Feret Diameter Calculations 41 Appendix 2: R Script for Statistical Analysis 45 Author’s Biography 49

TABLE OF FIGURES

Figure 1. Image of P. toxica 6 Figure 2. Image of care system 8 Figure 3. Image of experimental system 9 Table 1. Treatment water quality parameters of experiment 1 and 2 9 Figure 4. Images of histological slide s (1) 12 Figure 5. Images of histological slides (2) 12 Figure 6. Images of histological slides (3) 14 Figure 7. Images of histological slides (4) 14 Figure 8. Images of histological slides (5) 15 Figure 9. Box plot of Symbidoinium size comparison (1) 16 Figure 10. Images of histological slides (6) 17 Figure 11. Images of histological slides (7) 17 Figure 12. Images of histological slides (8) 18 Figure 13. Box plot of Symbidoinium size comparison (2) 19 Figure 14. Box plot of Symbiodinium size variance (1) 20 Figure 15. Images of histological slides (9) 21 Figure 16. Images of histological slides (10) 22 Figure 17. Images of histological slides (11) 22 Figure 18. Images of histological slides (12) 23 Figure 19. Images of histological slides (13) 24 Figure 20. Images of histological slides (14) 25 Figure 21. Box plot of Symbidoinium size comparison (3) 25 Figure 22. Box plot of Symbiodinium size variance (2) 27

INTRODUCTION

Anthropogenic Effects on Coral Reefs

It is abundantly clear that anthropogenic effects are changing the nature of coral reefs (Hughes et al 2003, Bellwood et al 2004), and that as humans continue to impact the global climate, the negative effects of these changes on coral reefs will increase (Hughes et al 2003, Hughes et al 2017). There are a number of different factors that can influence these effects, including stress effects from sedimentation and pollution (ISRS 2004), temperature (Glynn 1990, Winter et al 1998), high levels of solar radiation (Lesser et al

1990, Gleason and Wellington 1993, Brown 1997), ocean acidification (Anthony et al

2008), and nitrogen enrichment (Wiedenmann et al 2012, Vega-Thurber et al 2014).

One of the most well-known coral stress indicators is that of coral bleaching, or the expulsion of symbiotic dinoflagellate algae known as Symbiodinium normally maintained in coral polyp tissues (Brown 1997). This loss of Symbiodinium results in the coral tissue becoming pale or transparent, causing the skeleton to become visible and resulting in a white color. The first observed and published mass bleaching event of a wide range of coral reefs simultaneously occurred in Gulf of Chiriquí, Panama in 1982 during the El Niño temperature anomaly (Glynn 1984). Since this event, there have been numerous local and widespread mass bleaching events worldwide, including three widespread mass bleaching events in 1998, 2002, and from 2016 to 2017, the first back- to-back mass bleaching event (Hughes et al 2017) on the Indo-Pacific Archipelago. Other major bleaching events include the 2005 Caribbean reef bleaching event (Eakin et al

2010), and the 1982 Eastern Pacific bleaching event (Harriot 1985, Glynn 1990). The

result is clear: bleaching events are becoming increasingly common, and understanding these events is essential for reef protection and recovery (Hughes et al 2003).

The bleaching mechanism lies in the symbiotic relationship between

Symbiodinium and the coral animal, and understanding this relationship is essential to the coral bleaching mechanism. With a more complete understanding of this mechanism, human conservation policy can be accurately shaped to protect reefs from anthropogenic- caused stress.

Symbiodinium and the Adaptive Bleaching Hypothesis

Symbiosis, or “the living together of unlike organisms” (Douglas 1994) has been a long-observed relationship in biology. First used to describe the relationships among lichens in the late 1800’s (Trappe 2005), there are several required factors for a relationship to be classified as symbiosis. First, at least one organism must play a significant role in the life cycle of the second, and second, the organisms are in sustained and close contact throughout their lifetimes (Lewis 1973).

Symbiodinium are a genus of dinoflagellates found maintaining symbiotic relationships with several marine invertebrate phyla including Cnidaria, Mollusca,

Platyhelminthes and Porifera (Stat et al 2006). The most well-known relationship of these groups is that between Symbiodinium and the Cnidarian Anthozoa.

The relationship between coral animals and Symbiodinium, known colloquially as zooxanthellae, is an essential relationship within the coral animal life cycle.

Symbiodinium provide essential nutrients to the coral animal through photosynthesis for cell nutrition, maintenance and reproduction (Muscatine, Porter, 1977), fixing up to four times as much carbon as coral lacking Symbiodinium in their tissues (Muscatine et al

1984). Maintaining symbiotic relationships with algae allow coral to function as primary producers as well as primary and secondary consumers (Muscatine, Porter 1977), resulting in the role of coral in reef ecosystems as a keystone species. The carbon fixed by Symbiodinium plays an especially important role in the complex calcium carbonate skeleton certain coral secrete that comprises the physical shape of the reef itself (Anthony et al 2008). These calcifying coral not only function as animals within the reef ecosystem, coral compose the shape of the reef itself literally through their skeletal structures

(Tambutté et al 2011). Furthermore, Symbiodinium play an important role in juvenile coral growth, fixing carbon as the coral polyp settles on the reef (Cantin et al 2008).

Symbiodinium have only recently become the focus of understanding the adaptive mechanisms of coral to stress.

Symbiodinium were first grouped into 22 different taxonomic groups of indeterminate level using restriction fragment length polymorphisms from the small ribosomal subunit RNA (Rowan and Powers 1991). Later these groups became the modern clade taxonomic classification of these algae, composing nine clades (A-I), and a number of subclades, with primary associations of clade C with coral. Symbiodinium are acquired in coral tissue through two ways: horizontally from the environment, or vertically from parental populations (Byler et al 2013). Originally thought to be specific to the host, Symbiodinium transmission is flexible both between species of coral and within species, with species maintaining multiple subclades of a single Symbiodinium clade within their tissues, and some maintaining even multiple clades (Little et al 2004).

It has been shown that different Symbiodinium clades adjust the metabolism and stress coping mechanisms for different aspects of the holobiont system. For example, hosts

harboring clade C Symbiodinium have an increased growth rate (Little et al 2004), whereas populations maintaining clade D have a higher tolerance of thermal stress

(Rowan 2004, Baker et al 2004).

Herein lies the key to coral bleaching and coral adaptation to stress conditions.

Individual clades of Symbiodinium have relatively narrow ranges of stress tolerance due to their thylakoid lipid-membrane composition (Tchernov 2004), and thus it is very difficult for populations of Symbiodinium within a single clade to survive alone with the holobiont system in changing environments. However, there are two mechanisms that allow corals to shift the dominant clades of Symbiodinium maintained within their tissues: switching, or completely expelling the symbionts to replace with others taken up horizontally, and shuffling, or rearranging existing populations within their tissues for a different symbiont clade to influence the coral’s adaption to the environment (Reich et al

2017). Bleaching was first hypothesized in 1993 and later refined as an adaptive mechanism through which coral could shuffle Symbiodinium within their tissues to increase tolerance to local conditions known as the Adaptive Bleaching Hypothesis

(Buddemeier and Foutin, 1993; Buddemeier et al 2004). Further research into this mechanism lead to the hope that corals would be able to adapt for the short term to human-influenced changes, such as increasing ocean temperatures up to a point

(Berkelmans, Oppen 2004), though it is increasingly clear that the complicated nature of

Symbiodinium taxonomy and nuanced understanding of thermal and stress tolerance requires reclassification from the current taxonomic clade system (Sampayo et al 2008).

Whether or not scientists can use this relationship to help protect and potentially recover reefs in future anthropogenic climate fluctuations is unknown.

The Interaction Among Nitrogen, Coral, and Symbiodinium

The interaction between coral reefs and nitrogen has been a subject of study since the 1970’s, when the observation was first made of the coral reef nutrient paradox. Reefs are located in areas of relatively low nutrient levels and yet are able to maintain high levels of productivity (Webb et al 1975), thus the mechanisms through which coral are able to efficiently use nitrogen are of major concern. Nitrogen is the primary limiting nutrient for marine systems within coral- reef systems (Thomas 1969), and it has been shown that coral’s response to the nitrogen cycle and internal nitrogen cycle are closely tied with their susceptibility to bleaching and disease (Weidenmann et al 2013, Vega-

Thurber et al 2014, Rädecker et al 2015).

In brief, the nitrogen cycle begins in the atmosphere as N2, nitrogen is fixed by archaeobacteria and eubacteria located in sediment in an anaerobic process called

- denitrification to NO3 or nitrate, which is then taken up by organisms, usually plants, and

+ + - converted to ammonium (NH4 ). Microbial processes convert NH4 to NH3, then NO2 ,

- and finally to NO3 , and the cycle can repeat. These converted forms of nitrogen,

= + + especially NO3 and NH4 , can be taken up by organisms and transformed back to NH4

+ or urea, which then sink down into the sediment and are converted again. NH4 can also be nitrified back into N2, where it is returned in the atmosphere (Falkowski 1997).

However, human impact on the nitrogen cycle through artificial fertilizers in

- modern agriculture and burning fossil fuels have added a second influx of NO3 into the system, where it is washed into the ocean, most notably manifesting as algae blooms and areas of anoxic or dead zones along the coast (Vitousek et al 1997, Canfield et al 2010).

There is clear evidence that these anoxic blooms are increasing with time, resulting in

decreasing diversity along coastal dead zones including among seagrass beds and coral reefs, lower fish and shellfish resources for human fisheries and aquaculture, increasing dominance of nuisance algae and increasing occurrences of toxic dinoflagellate blooms or “red tides” known to cause shellfish poisoning among humans (Vitousek et al 1997).

Tropical coral reefs are most often located in coastal or near-coast ecosystems and are thus subject to these anthropogenic influxes of nitrogen into the system. Because coral maintain symbiotic algae within their tissues, it is likely that this influx of nitrogen into the ocean is causing changes to these communities. Very little research has been done on the interactions between coral and the nitrogen cycle, though there is increasing evidence that nitrogen plays an important role in the dynamic of the coral holobiont and the bacterial and algal members of its symbiotic community (Siboni et al 2008,

Pogoureutz et al 2017)

Palythoa toxica and Study Objectives

Figure 1. A colony of Palythoa toxica (Walsh and Bowers 1971) from the University of Maine Aquaculture Research Collection used in this experiment. Photo credit to Allyson Redcay.

Palythoa toxica (Walsh and Bowers 1971) is a zoanthid hexacoral that produces the infamous palytoxin, one of the most deadly toxins within the natural world. This

species is not a reef-building sclearactinian coral. Instead these animals have relatively large anemone-like polyps, but maintain similar symbiotic algal associations as scleractinian coral. The species was first isolated in 1971 from Hawaii from a tropical coral referred to as limu-make-o-Hana, and was the first organism discovered to produce palytoxin (Walsh and Bowers, 1971). Since its discovery, palytoxin and its effects on humans have become the focus of research on this group (Moore and Scheuer, 1971;

Deeds and Schwartz 2010). Currently there is very little research on histological changes during prolonged stress periods, though ecological studies of Symbiodinium in zoanthid coral have been conducted (Kemp and Cook 2006; Burnett 2002) and indicate zoanthid coral exhibit similar bleaching mechanisms as scleractinians (Kemp and Cook, 2006).

Furthermore, P. toxica is relatively easy to obtain and culture, and was used for this experiment due to its accessibility and lack of research focusing on this particular symbiotic relationship.

The goal of this study is understanding the relationship between P. toxica and its associated Symbiodinium when exposed to elevated levels of nitrogen and temperature both separately and in combination using histological analysis. Symbiodinium were not genetically analyzed to determine clade. The hypothesis of the study was based on increased growth of dinoflagellates when exposed to elevated nutrient levels (Parkhill and Cembella 1999), and tests for a direct relationship between the level of nitrogen within the coral tissues and the size of the Symbiodinium cells. Temperature was added as a second factor to understand stress effects on coral in combination with nitrate stress, and to retest the effects of temperature on coral bleaching, especially for the relatively understudied zoanthid clade.

MATERIALS AND METHODS

System Design and Sampling

Palythoa toxica samples were acquired from a local aquarist in Orono, ME, and moved to the University of Maine Aquaculture Research Center where they were maintained in a 710 liter raceway (290 L raceway, 420 L bottom sump) under a Tek light

T5. The water was maintained around 26˚C, 32 ppt salinity, a pH of approximately 8, and a water exchange rate of 35 L/sec.

Figure 2. The system in which colonies of P. toxica were maintained before the experiment.

The samples comprised three genetically identical colonies of various size. 125 polyps were cloned from these samples using an Ecotech Marine Coral Propagation Kit, glued onto cement coral plugs using Ecotech coral glue, and maintained in the raceway for two weeks prior to the experiment. The design of the experiment comprised one control and four experimental treatments with one 4-L tank per treatment. 60 polyps were used for each experiment, in groups of 12 polyps per treatment.

Figure 3. A reconstruction of the experimental system setup.

Two experiments were run over the course of this project, both lasting 14 days, of two levels of elevated nitrate and two levels of elevated temperature (Table 1). Nitrate levels were elevated by adding powdered NaNO3. After completion of experiment one tanks were drained and cleaned using a water and bleach rinse before being randomly rearranged for experiment two. Experiment two consisted of the same experimental setup. Nitrate levels were readjusted for the second experiment, and the same temperature levels were used.

Treatment Experiment 1 Experiment 2 1 26˚C, 0 ppm 26˚C, 0 ppm 2 26˚C, 7 ppm 30˚C, 10 ppm

3 26˚C, 15 ppm 30˚C, 20 ppm 4 30˚C, 0 ppm 35˚C, 10 ppm 5 35˚C, 0 ppm 35˚C, 20 ppm

Table 1. Treatment temperature and nitrate levels for experiment 1 and experiment 2.

All tanks contained a fish tank heater of 50 or 100 watts to maintain temperature, and were maintained beneath two 50 watt lights, on a light schedule of 12 hours on, 12 hours off (Figure 3).

For both experiments two polyps were sampled from the control treatment on day one, and then from every treatment every four days. On day 14 all remaining polyps were sampled from each treatment. Histology fixation was conducted using a standardized procedure as follows: samples were placed in 10% neutral buffered formalin, placed in a standard tissue processor for 16.5 h, then embedded in paraffin. Five-µm thick sections were cut and stained with hematoxylin and eosin. Sampling protocol was designed using

IACUC Power Study protocol (University of Maine). Nitrogen-enrichment levels were based on two Australian reefs with higher levels of nitrate: Yongola Reef in the mid-

Great Barrier (Vridaka) as the low-enriched level, and Rottnest Island, in Western

Australia as the high-enrichment level (Thomson et al 2011). Nitrogen levels were tested every day for the first four days of each experiment using a LaMotte water quality testing kit and then every other day for the duration of the experiment. When nitrogen levels were lower than the required level, approximately 0.01 grams of NaNO3 was added to the

- tanks (increasing NO3 by approximately 2 ppm).

Symbiodinium Cell Size Analysis

Feret diameter was used as a quantitative measurement of the change of size in

Symbiodinium (See Appendix 1). Slides were photographed using an Olympus CX31 microscope with Infinity Capture software. Photographs were analyzed using ImageJ version 1.51s and Java1.8.0_151. Analysis was calibrated for 40x images using a Motis calibration for 40x, 100x and 400x magnification. The perimeters of 100 randomly

selected Symbiodinium were traced using the freehand trace function and measured for each polyp for a total of 200 Feret diameters for each treatment barring the samples from

Trial 1 Treatment 1 Day 1 (3 polyps), Treatment 3 Day 12 (3 polyps), Treatment 4 Day 8

(1 polyp), Treatment 5 Day 8 (3 polyps), and Trial 2 Treatment 1 Day 1 (1 polyp), as well as samples from the 14th day of the experiments (3-4 polyps). Due to stress, the polyps from the 12th day of Trial 2 for treatments 4 and 5 were too degraded for histology, and thus there are no data for those samples. Symbiodinium were treated as the individual

(N=1) for this process instead of polyps within the treatment.

Statistical Analysis

All statistical analysis was conducted in RStudio version 3.3.3 on a basis of 95% confidence. Data were organized by Experiment (1 or 2), treatment (control, high/low nitrate, high/low temperature), time of sampling (Day 4, 8, 12, or 14) and polyp. Each

Symbiodinium Feret measurement was assigned a number, and average Feret diameter was calculated by averaging the Feret maximum and minimum. One-way analysis of variance (ANOVA) was used to compare mean Feret of treatments (See Appendix 2).

Outliers were removed from each dataset and assumptions of the ANOVA were met by each set (Levene’s test for symmetry, p>0.05, Shapiro-Wilk test for normality, p>0.05).

Two ANOVA analyses were conducted. The first ANOVA used treatments as the explanatory variable, and the second used day of sampling as the explanatory variable. A

Tukey HSD post-hoc was used for both ANOVA results for individual mean comparison.

RESULTS

Experiment One

Figure 4. Control Day 4, from left to right, top to bottom: 400x, 100x and 40x images of Polyp II.

Figure 5. Images of control polyps throughout different sampling times of the experiment. Green arrows indicate healthy Symbiodinium cells.

To establish a concept of a ‘healthy’ polyp as a basis of comparison for the experiment, samples of the control polyp were taken at every sampling step of the

experiment. There are several factors that characterize a healthy polyp for this experiment, as can be demonstrated by Figures 4 and 5. Healthy polyps contain no faded tissue; in other words, polyp tissue is tinted darker shades than bleached counterparts.

Healthy Symbiodinium are in general maintained in one of two places: either along the tentacles near the top of the polyp (Figure 4, 400x, Figure 5, Day 12, 400x) or maintained in gastrodermis in the mesenteries of the polyp (Figure 5, Day 14 400x Polyps 1 and 3).

Small numbers of Symbiodinium could also be found maintained in vesicles within the polyp epidermis (Figure 5, Day 14 Polyp 3). Finally, there is no sign of Symbiodinium lysis; in other words, all cells of Symbiodinium are whole and contain plastids (Figure 4,

400x, Figure 5, all 400x images show relatively healthy Symbiodinium). Not every control polyp displayed these characteristics; bleaching occurred in the control treatment of the second experiment on the 14th day.

Effects of Elevated Nitrogen

Figure 6. Images of polyps taken from Treatments 2 and 3 (elevated Nitrogen treatments) at 40x throughout the experiment. Polyps in treatment 3 days 4, 8 and 14 exhibit moderate bleaching.

Figure 7. Images of polyps taken from Treatments 2 and 3 (elevated Nitrogen treatments) at 400x throughout the experiment.

Within elevated nitrogen treatments, there was a major difference between these treatments and the control treatments. Full bleaching of coral tissue did not occur; however, there was a trend of gastrointestinal pockets of Symbiodinium among the

tentacles and mesenteries being expelled into the coelenteron of the polyp (Figure 7).

Near the end of the experiments, some of the Symbiodinium had lysed within Treatment 3

(Figure 7; Treatment 3, Day 14).

Figure 8. A comparison of Symbiodinium on the last day of the experiment for the control (treatment 1) and elevated nitrogen treatments (2 and 3) at 400x.

There appears to be a trend with time between different levels of elevated nitrate.

Small levels of nitrate may stimulate growth of Symbiodinium (Figure 8) to an unhealthy degree. Higher levels of nitrate cause coral tissue to bleach partially, but retain those

Symbiodinium within their system.

Figure 9. Box plots comparing Symbiodinium Feret diameter among Control, Treatment 2 and Treatment 3. Matching colors represent insignificant differences (ANOVA, P>0.05). In the case of Day 12, Treatment 2, the mean was significantly different from Treatment 1, but not from Treatment 3.

Elevated nitrate affected Symbiodinium Feret diameter, however, how nitrate affected the size of the algae cells varies between time and treatment (Figure 9). For the first half of the experiment, there was a clear trend of increasing size with increased nitrate levels within the water. As the experiment continued however, the average Feret diameter of treatment 3 decreased from almost 10 µm in diameter on Day 4 at the highest to as low as 9 µm on Day 12. Within nitrate treatments, Symbiodinium Feret diameter remains between 8 and 10 µm. For both treatment 2 and 3, diameter increased in the first half of the experiment and appears to decrease in the second half.

Effects of Elevated Temperature

Figure 10. Images of polyps taken from Treatments 4 and 5 (elevated temperature treatments) at 40x throughout the experiment. Treatment 4 on day 14, and treatment 5 on day 4 exhibit symptoms associated with bleaching.

Figure 11. Images of polyps taken from Treatments 4 and 5 (elevated temperature treatments) at 400x throughout the experiment.

Elevated temperature in general caused minor bleaching (Figure 10, Treatment 4

Day 14, Treatment 5 Day 4 and Day 14). In general, similar to nitrogen treatments, there was a trend of Symbiodinium being expelled from the polyp tissues to coagulate in spaces

among tentacles and within the coelenteron of the coral (Figure 11, Treatment 4 Day 14,

Treatment 5 Day 4). Some breakdown of gastrointestinal tissue occurred in particularly unhealthy polyps (Figure 10, Treatment 4 Day 14, Figure 9, Treatment 4), however the major trend remained change in Symbiodinium location, moving out of gastrointestinal lining into spaces inside the polyp (Figure 10). Bleaching remained minimal, with only one polyp in treatment 4 and one polyp in treatment 5 displaying signs of moderate bleaching.

Figure 12. A comparison of Symbiodinium on the last day of the experiment for the control and elevated temperature treatments at 40x.

Figure 13. Box plots comparing Symbiodinium Feret diameter between Control, Treatment 4 and Treatment 5. Matching colors represent insignificantly different samples (ANOVA, P>0.05).

The pattern of Symbiodinium diameter size change in elevated temperature treatments was more random than elevated nitrate treatments but displayed a similar pattern. In general, treatments 4 and 5 had significantly different means than the control

(Figure 13). In the case of the beginning of the experiment, both treatment 4 and 5 had larger Feret diameters on average, with treatment 4 being the largest of the three.

However, by the end of the experiment, the average Feret diameter of treatments 4 and 5 had decreased. By Day 12, there was no significant difference between treatment 4 and the control. By Day 14, there was no significant difference between treatment 5 and the control, and treatment 4 had decreased significantly to below the average Feret diameter of the control.

Variance Among Treatments

Figure 14. Box plots of average Feret diameter within each individual treatment throughout the experiment. Matching colors represent insignificantly different samples (ANOVA, P>0.05). In the case of multiply colored samples, a significant difference occurred for at least one treatment comparison. For treatment 2, Day 12, Feret diameter was significantly different from Day 14 but equivalent to Days 4 and 8. In Treatment 4, Day 14 was not significantly different from either Day 4 or Day 12, but both Day 4 and Day 12 varied from each other. Finally in Treatment 5, Day 8 and 14 were significantly different than Day 12, but not Day 4.

There was significant variance among individual tank Feret diameters as time passed during the experiment. Among treatments 1, 2 and 3, treatment Feret diameter remained consistent for several days at a time but still varied at different points of the experiment. Treatments 4 and 5 varied to a higher extent, with Treatment 4 containing the most variation among Feret diameters within the experiment.

Experiment Two

Figure 15. Images of the control polyps throughout experiment 2 and 40x and 400x.

Control polyps of experiment 2 matched the healthy characteristics from experiment 1 with little change. Healthy Symbiodinium were maintained between tentacles, among mesentarial tissues and within vesicles along the epidermis of the polyp.

Bleaching occurred within the control tank on Day 14.

Effects of Combined Elevated Nitrogen and Temperature

Figure 16. Images of polyps from treatments 3 (low temperature, high nitrate) and 4 (high temperature, low nitrate) at 40x.

Figure 17. Images of polyps from treatments 3 (low temperature, high nitrate) and 4 (high temperature, low nitrate) at 400x.

Within varying combined levels of stress, there was a clear trend that higher temperatures in combination with nitrate stress of any level resulted in increased bleaching (Figure 16, Figure 17). High elevated nitrate levels and lower elevated

temperature levels did not cause bleaching to occur in treatment 3 (whereas moderate and some severe bleaching occurred in highest nitrate levels in experiment 1). The most severe cases of bleaching occurred on Day 12 within treatments 4 and 5 as samples were too damaged to be histologically fixed. Within treatment 3, however, high temperature in combination with stress resulted not only in more frequent bleaching, but more severe bleaching when it did occur, though there was no bleaching on the final day of experiment 2 in either treatment 3 or 4. Bleaching generally resulted in Symbiodinium still being maintained within the coral coelenteron. Fading of coral tissues was more obvious within experiment 2 (Figure 17 Treatment 4 Days 4 and 8).

Figure 18. Images of polyps from treatments 2 (low temperature, low nitrate) and 5 (high temperature, high nitrate) at 40x.

Figure 19. Images of polyps from treatments 2 (low temperature, low nitrate) and 5 (high temperature, high nitrate) at 400x.

Treatment 2 contained no bleached samples, generally maintaining relatively healthy tissues and Symbiodinium compared to control treatments. Treatment 5 contained the most occurrences of bleaching, with bleaching occurring severely in every sample taken (Figure 19, Figure 20, Treatment 5), and the samples from Day 12 were too damaged to even histologically fix. Movement of Symbiodinium bleaching is most obvious within these treatments. Tissues are completely pale, Symbiodinium are found exclusively in either the coelenteron or between tentacles, and contain several instances of what may be lysed cell material among surviving Symbiodinium (Figure 19, Treatment

5, Day 14, Figure 19).

Figure 20. A comparison of Symbiodinium between all four treatments of experiment 2 at 400x on the final day of the experiment.

Figure 21. Box plots comparing average Feret diameter within each individual treatment throughout the experiment. Matching colors represent insignificantly different samples (ANOVA, P>0.05). In the case of Treatment 4, Day 4, Feret was significantly different from treatment 5, but not treatment 1. Within Day 8, treatments 2, 3 and 5 were significantly different than treatment 1, but not treatment 4. Within Day 14, treatments 2, 3 and 4 were significantly different than treatment 5, but not from treatment 1.

Within experiment 2 Symbiodinium Feret diameter varied generally differently than the control treatment (with the exception of Day 14) with no other discernible pattern. It seems with multiple stressors on the polyp, the Feret diameter generally increases in size, with the major exception of samples from Day 14.

Variance Among Treatments

Figure 22. Box plots comparing average Feret diameter within each individual treatment throughout the experiment. Matching colors represent insignificantly different samples (ANOVA, P>0.05).

In general, Feret diameter varied strongly within each treatment more so than in experiment 1. In the case of all experimental treatment, there is strong variation throughout the experiment with nearly every average value varying significantly from the previous sample with the most variation occurring in treatments 2 and 3.

DISCUSSION

Nitrogen Effects on P. toxica

There is a general trend of size increase among elevated nitrate treatments in the

Feret diameters of Symbiodinium; however, this trend is inconsistent throughout the experiment. Furthermore, there is a trend of variation within each system. There is significant variation within the control treatment with a general trend of Feret size increase; however these conditions were kept constant. This is potentially a sign that when P. toxica is maintained at constant condition, healthy Symbiodinium will grow, causing the increase in size (Figure 14, Figure 22). Both treatments displayed a similar trend: near the beginning of the experiment, size was elevated directly with size, though by the end of the experiment both treatments dropped to either an insignificantly different size to the control, or significantly lower. This could mean that with elevated nitrogen there is a promotion of growth due to these excess nutrients, then a collapse of the system, resulting in the cell lysis displayed in both nitrate treatments and an overall decrease in size.

Size trends within each treatment varied significantly as well. The control treatment varied significantly, increasing with time during the experiment. Originally it was thought that if conditions remained constant and healthy, Symbiodinium size would not be affected. However, this could support the idea that a healthy coral displays

Symbiodinium growth. Meanwhile, Treatment 2 remained consistent, with a slight insignificant decrease. Treatment 3 varied significantly, decreasing and then increasing.

Nitrate uptake in Symbiodinium has been studied in the past, and there is a consensus of Symbiodinium nitrogen uptake in forms of ammonium and nitrate, with a preference for ammonium (D’Elia et. al. 1983). Furthermore, sudden increases in nitrogen cause an increase in uptake of nitrogen in Symbiodinium and storage as uric crystals (Kopp et al 2013). This could be an explanation of the sudden increase and subsequent decrease in size of the Symbiodinium, however, within this experiment the timeline of Symbiodinium increase and decrease within treatments 2 and 3 occurs more slowly than formation of uric crystals, and these crystals were not examined within the cell. Corals generally display a nutrient limitation on their symbionts, disrupted by organic carbon (Wang and Douglas 1998), and the effects of nutrient stimulation on corals is potentially helpful in the short term, promoting growth in the symbionts, but eventually leading to a collapse in growth, especially with compounded stressors. There is also evidence that the coral microbiome plays a large role in the nitrogen cycle within corals (Pogourez et al 2017), so there is good reason to believe that there is more happening within this pattern of Symbiodinium growth and collapse than what this study examined. Recent studies have begun to emphasize the importance of nitrogen as a bleaching trigger (Wiedenmann et al 2012).

Bleaching occurred for the samples enriched in nitrate with a total of eleven cases of bleaching out of the total of twenty one samples, about 50%. However, the bleaching that did occur was moderate in general, with most Symbiodinium colonies remaining in the coelentron of the coral. Only by the very end of the experiment in the highest nitrate treatment did severe bleaching occur in Day 12 and Day 14. Therefore, according to the

results of this experiment, elevated nitrate triggers the bleaching mechanism of corals to some extent.

Temperature Effects on P. toxica

The effect of temperature on the Feret diameter of Symbiodinium has a similar trend to the nitrogen enriched treatments. At the beginning of the experiment, there is a trend of increasing Symbiodinium size in the elevated temperature treatments. By Day 8, treatment 4 (held at 30˚C) had the highest Feret diameter, before crashing in size and falling to a significantly smaller diameter than the control treatment. Treatment 5 had a similar trend of increasing in size and decreasing in size, though these changes were less extreme. By the end of the experiment Treatment 5 had fallen back to an insignificantly different size than the control.

These could be following a similar pattern of the nitrogen enrichment experiment.

It has been shown in Pocillopora coral that slight temperature elevation can increase the size of Symbiodinium for short periods of time (Edmunds et al 2005), however this effect ceased at 27˚C. There was significant variation in the temperature of the two treatment tanks during the experiment, so this may have contributed to the slight increase in size and eventual decrease. Furthermore, this effect was only noted for short-term exposure

(24 hours), whereas this experiment incubated coral polyps for 14 days.

Within treatments, similar to the nitrogen enriched treatments, there was a trend of variance through time. Symbiodinium size in both treatments increased significantly compared to the control (for treatment 4 this occurs on Day 8; for treatment 5 this occurred on Day 12), before a collapse, resulting in sizes significantly lower than the control.

There is very little literature on the dynamics of how Symbiodinium change size during temperature elevation, as this method for measuring Symbiodinium size change was a novel method developed from coral reproduction methods (Waller, Tyler 2005).

However, it has been shown that Symbiodinium photosynthesis decreases in response to elevated temperature (Goulet et al 2005), though this response varies depending on the coral acclimation to temperature (Kemp 2006). This experiment on these zoanthids are the first case of a more extended length study on elevated temperature on Symbiodinium size, and it is clear that there is a linkage between length of time in treatment and

Symbiodinium size.

There is an indication that temperature is a bleaching trigger in P. toxica, with the highest temperature treatment resulting in six cases of bleaching, the most bleaching occurring at the end of the experiment, linking time to bleaching as well. Prolonged exposure to elevated temperature is the most consistent trigger of coral bleaching and has been studied numerous times, mostly in scleractinian corals (Glynn and D’Elia 1990,

Brown 1997, Hughes et al 2017) but in zoanthids as well (Kemp et al, 2006). This experiment serves further confirmation of the bleaching response in P. toxica.

Combined Stress Effects

Nitrate and temperature enrichment in combination mostly continue the pattern in

Symbiodinium size displayed in the first experiment. In the most stressed conditions in treatment 5, there is no significant difference from the control. Treatment 4 displayed the pattern of Symbiodinium increase in size and decrease in size, which again supports the idea that low amounts of nitrogen stimulate Symbiodinium growth. However, for the most

part there is not much significant difference in Symbiodinium sizes between treatments within experiment 2.

It is clear that combined stressors within an environment cause further damage to systems than singular stressors alone (Paine et al 1998, Hughes et al 1999). There has been a movement in coral research to understand multiple stressors on coral in combination, especially in combination with temperature (Coles and Jokiel, 1978, Porter et al 1999, Wiedenmann et al 2012). It has been shown that elevated nitrogen increases coral’s susceptibility to bleaching (Widenmann et al 2012). There has been some debate in the past as to whether or not elevated nitrogen is unhealthy for coral reefs, with studies that support very different ideas about nitrogen and coral. One study supports the idea that low amounts of nitrogen stimulate coral growth (Atkinson et al 1995). Other studies show that elevated nitrogen does not have a negative effect on coral growth (Szmant

2002, Bongiorni et al 2003). More recently it was shown that elevated nitrogen increases bleaching severity and disease susceptibility (Vega-Thurber et al 2014).

According to the results of this study, however, there is a clear detrimental effect of combined nitrate and temperature stress on coral. The instances of bleaching within the second experiment outnumbered the instances of bleaching in experiment one, with fourteen instances of bleaching across all treatments, as opposed to the eleven cases of bleaching in experiment one. Furthermore, the vast majority of bleaching in experiment one was moderate, with only a slight fading in zoanthid tissue color. Symbiodinium cells were maintained within the coelenteron of the polyp rather than being expelled from the polyp completely. Within the second experiment however, nearly every case of bleaching was severe, with high degradation of coral tissues (to the point where samples were

unable to be histologically fixed). Finally, Symbiodinium were for the most part lysed, and zoanthid tissue was nearly clear (Figure 16, 18), especially in high temperature treatments. According to the results of this experiment, then, there is evidence that elevated nitrate in combination with temperature is a bleaching trigger within Palythoa toxica.

Conclusions

Overall, the results of this experiment confirm the effect of elevated temperature as a bleaching trigger for P. toxica and provide evidence that elevated nitrate is a potential bleaching trigger in prolonged exposure. Furthermore, there is evidence that combined elevated nitrate and temperature compound bleaching mechanisms, with the highest amount of bleaching occurring in the most stressed treatments.

A pattern of Symbiodinium size changes emerges within this experiment as well.

When exposed to low levels of nitrate and moderate temperature elevation, there is a trend of Symbiodinium Feret diameter increase. With time, however, there is an eventual collapse of growth, resulting in Symbiodinium diameter decrease and an increase of algal mass potentially from cell lysis within bleached Symbiodinium remaining between tentacles and within the coelenteron.

However, these conclusions come with major caveats. There is no replication within this experiment between coral polyps with only two genetically identical polyps sampled from the same water treatment. In future experiments, it will be necessary to replicate treatments, polyps and genetic individuals to provide for good replication.

Furthermore, there was a significant amount of variation within treatments, so further water quality maintenance methods must be developed, as coral prove very delicate in

care (as demonstrated by the bleaching that occurred in the control treatment near the end of experiment 2). That being said, this experiment does provide preliminary evidence of

Symbiodinium growth dynamics when exposed to prolonged periods of nitrate and temperature stress. It is likely that prolonged stress results in different dynamics of coral adaptation to stressed conditions, and this project is part of that effort to understand sublethal prolonged chronic stress conditions. Future experiments building on the knowledge gained from this project are essential to understanding the true dynamic between nitrogen, temperature and Symbiodinium.

REFERENCES

Anthony, K. R. N., Kline, D. I., Diaz-Pulido, G., Dove, S., & Hoegh-Guldberg, O. (2008). Ocean acidification causes bleaching and productivity loss in coral reef builders. Proceedings of the National Academy of Sciences of the United States of America, 105(45), 17442–6. https://doi.org/10.1073/pnas.0804478105 Atkinson, M.J., Carlson, B. & Crow, G.L. Coral growth in high-nutrient, low-pH seawater: a case study of corals cultured at the Waikiki Aquarium, Honolulu, Hawaii. Coral Reefs (1995) 14: 215. https://doi.org/10.1007/BF00334344 Baird, A. H., Bhagooli, R., Ralph, P. J., & Takahashi, S. (2008). Coral bleaching: the role of the host. Trends in Ecology and Evolution, 24(1), 16–20. https://doi.org/10.1016/j.tree.2008.09.005 Baker, A. C. (2003). Flexibility and Specificity in Coral-Algal Symbiosis: Diversity, Ecology, and Biogeography of Symbiodinium. Source: Annual Review of Ecology, Evolution, and Systematics Annu. Rev. Ecol. Evol. Syst, 34(34). https://doi.org/10.1146/132417 Baker, A. C., Starger, C. J., McClanahan, T. R., & Glynn, P. W. (2004). Corals’ adaptive response to climate change: Shifting to new algal symbionts may safeguard devastated reefs from extinction. Nature. https://doi.org/10.1038/430741a Bellwood, D. R., Hughes, T. P., Folke, C., & Nyström, M. (2004). Confronting the coral reef crisis. Nature, 429(6994), 827–833. https://doi.org/10.1038/nature02691 Bellwood, D. R., Hughes, T. P., Bellwood, D. R., Card, M., Connolly, S. R., Folke, C.,Roughgarden, J. (2001). Regional-Scale Assembly Rules and Biodiversity of Coral Reefs. Science, 292(5521), 1532–1535. https://doi.org/10.1126/science.1058635 Berkelmans, R., & van Oppen, M. J. H. (2006). The role of zooxanthellae in the thermal tolerance of corals: a “nugget of hope” for coral reefs in an era of climate change. Proceedings. Biological Sciences, 273(1599), 2305–12. https://doi.org/10.1098/rspb.2006.3567 Bongiorni, L., Shafir, S., Angel, D., & Rinkevich, B. (2003). Survival, growth and gonad development of two hermatypic corals subjected to in situ fish-farm nutrient enrichment. Marine Ecology Progress Series, 253, 137-144. doi:10.3354/meps253137 Brown, B. E. (1997). Coral bleaching: causes and consequences. Coral Reefs, 16, 129– 138. Buddemeier RW, & Fautin DG. (1993). Coral Bleaching as an Adaptive Mechanism. BioScience, 43(5), 320–326. Buddemeier, R. W., Baker, A. C., Fautin, D. G., & Jacobs, J. R. (2004). The Adaptive Hypothesis of Bleaching. Coral Health and Disease, 427-444. doi:10.1007/978-3- 662-06414-6_24

Burnett, W. J. (2002). Longitudinal variation in algal symbionts (zooxanthellae) from the Indian Ocean zoanthid Palythoa caesia. Marine Ecology Progress Series. https://doi.org/10.3354/meps234105 Byler, K. A., Carmi-Veal, M., Fine, M., & Goulet, T. L. (2013). Multiple symbiont acquisition strategies as an adaptive mechanism in the coral Stylophora pistillata. PloS One, 8(3), e59596. https://doi.org/10.1371/journal.pone.0059596 Coles, S.L. & Jokiel, P.L. (1978). Synergistic effects of temperature, salinity, and light on the hermatypic coral Montipora verrucosa. Marine Biology, 49: 187. https://doiorg.prxy4.ursus.maine.edu/10.1007/BF00391130 Deeds, J., & Schwartz, M. (2010). Human risk associated with palytoxin exposure. Toxicon, 56(2), 150-162. https://doi.org/doi:10.1016/j.toxicon.2009.05.035 D’Elia, C. F., Domotor, S. L., & Webb, K. L. (1983). Nutrient uptake kinetics of freshly isolated zooxanthellae. Marine Biology, 75, 157–167. Douglas, Angela (1994), Symbiotic interactions, Oxford University Press, ISBN 0-19- 854294-1 Eakin, C. M., Morgan, J. A., Heron, S. F., Smith, T. B., Liu, G., Alvarez-Filip, L., … Yusuf, Y.(2010). Caribbean Corals in Crisis: Record Thermal Stress, Bleaching, and Mortality in 2005. PLoS ONE, 5(11), e13969. https://doi.org/10.1371/journal.pone.0013969 Edmunds, P. J., Gates, R. D., Leggat, W., Hoegh-Guldberg, O., & Allen-Regua, L. (2005). The effect of temperature on the size and population density of dinoflagellates in larvae of the reef coral Porites astreoides. Invertebrate Biology, 124(3), 185–193. Fabricius, K. E. (2005). Effects of terrestrial runoff on the ecology of corals and coral reefs: review and synthesis. Marine Pollution Bulletin, 50, 125-146. https://doi.org/10.1016/j.marpolbul.2004.11.028 Falkowski, P. G. (1997). Evolution of the nitrogen cycle and its influence on the biological sequestration of CO2 in the ocean. Nature, 387, 272-275. https://doi.org/10.1038/387272a0 Gleason, D. F., & Wellington, G. M. (1993). Ultraviolet radiation and coral bleaching. Nature, 365(6449), 836–838. https://doi.org/10.1038/365836a0 Glynn, P. W., & D ’croz, L. (1990). Experimental evidence for high temperature stress as the cause of El Nifio-coincident coral mortality. Coral Reefs, 8, 181–191. Goreau, T. J., & Hayes, R. L. (1994). Coral Bleaching and Ocean “Hot Spots.” Ambio, 23, 176–180. Goulet, T. L., Cook, C. B., & Goulet, D. (2005). Effect of short-term exposure to elevated temperatures and light levels on photosynthesis of different host–symbiont combinations in the Aiptasia pallida Symbiodinium symbiosis. Limnol. Oceanogr, 50(5), 1490–1498.

Harriott, V. J. (1985). Mortality rates of scleractinian corals before and during a mass bleaching event. Marine Ecology, 21, 81–88. Hughes, T. P., Baird, A. H., Bellwood, D. R., Card, M., Connolly, S. R., Folke, C., … Marshall, P. (2003). Climate Change, Human Impacts, and the Resilience of Coral Reefs. Science, 301, 929–933. Hughes, T. P., & Connell, J. H. (1999). Multiple stressors on coral reefs: A long-term perspective. Limnol. Oceanogr, 44(2), 932–940. Hughes, T. P., Kerry, J. T., Álvarez-Noriega, M., Álvarez-Romero, J. G., Anderson, K. D., Baird, A. H., … Wilson, S. K. (2017). Global warming and recurrent mass bleaching of corals. Nature, 543(7645), 373–377. https://doi.org/10.1038/nature21707 ISRS (2004) The effects of terrestrial runoff of sediments, nutrients and other pollutants on coral reefs. Briefing Paper 3, International Society for Reef Studies, pp: 18 Kopp, C., Pernice, M., Domart-Coulon, I., Djediat, C., Spangenberg, J. E., Alexander, D. T. L., … Meibom, A. (2003). Highly Dynamic Cellular-Level Response of Symbiotic Coral to a Sudden Increase in Environmental Nitrogen. mBio, 4(3): 1- 9. https://doi.org/10.1128/mBio.00052-13 Little, A. F., van Oppen, M. J., & Willis, B. L. (2004). Flexibility in Algal Endosymbioses Shapes Growth in Reef Corals. Science, 304, 1492–1494. Lesser, M. P., Stochaj, W. R., Tapley, D. W., & Shick, J. M. (1990). Bleaching in coral reef anthozoans: effects of irradiance, ultraviolet radiation, and temperature on the activities of protective enzymes against active oxygen. Coral Reefs, 8, 225–232. Moore, R., & Scheuer, P. (1971). Palytoxin a new marine toxin from a coelenterate. Science, 172(3982), 495-498. https://doi.org/doi:10.1126/science.172.3982.495 Muscatine L, & Porter JW. (1977). Reef Corals: Mutualistic symbioses adapted to nutrient-poor environments. BioScience, 27(7), 454–460. Muscatine L, Falkowski PG, Porter JW, & Dubinsky Z. (1984). Fate of photosynthetic fixed carbon in light and shade-adapted colonies of the symbiotic coral Stylophora pistillata. Proc. R. Soc. Lond., 222, 181–202. Neal E. Cantin, Madeleine J. H. van Oppen, Bette L. Willis, J. C. M. &, & Andrew P. Negri. (2009). Juvenile corals can acquire more carbon from high performance algal symbionts. Coral Reefs, 28, 405–414. Paine, R. R., Tegner, M. J., & Johnson, E. A. (1998). Compounded Perturbations Yield Ecological Surprises. Ecosystems, 1, 535–545. Parkhill, J.-P., & Cembella, A. D. (1999). Effects of salinity, light and inorganic nitrogen on growth and toxigenicity of the marine dinoflagellate Alexandrium tamarense from northeastern Canada. Journal of Plankton Research, 21(5), 939–955.

Pogoreutz, C., Rädecker, N., Cárdenas, A., Gärdes, A., Voolstra, C. R., & Wild, C. (2017). Sugar enrichment provides evidence for a role of nitrogen fixation in coral bleaching. Global Change Biology, 23(9), 3838–3848. https://doi.org/10.1111/gcb.13695 Porter, J. W., Lewis, S. K., & Porter, K. G. (1999). The effect of multiple stressors on the Florida Keys coral reef ecosystem: A landscape hypothesis and a physiological test. Limnol. Oceanogr, 44(2), 941–949. Rädecker, N., Pogoreutz, C., Voolstra, C., Wiedemann, J., & Wild, C. (2015). Nitrogen cycling in corals: the key to understanding holobiont function. Trends in Microbiology, 23(8), 490–497. Reich, H. G., Robertson, D. L., & Goodbody-Gringley, G. (2017). Do the shuffle: Changes in Symbiodinium consortia throughout juvenile coral development. PLOS ONE, 12(2), e0171768. https://doi.org/10.1371/journal.pone.0171768 Reimer, J. D. (2008). Implications for different diversity levels of Symbiodinium spp. (Dinophyceae, Suessiales) within closely related hosts: zoanthids (Cnidaria: Hexacorallia: Anthozoa) as a case study. Galaxea, 10, 3–13. Rowan, R., & Powers, D. (1991). A Olecular Genetic Classification of Zooxanthellae and the Evolution of Animal-Algal Symbioses. Science, 251, 1348–1351. Rowan, R. (2004). Coral bleaching: Thermal adaptation in reef coral symbionts. Nature, 430, 742. https://doi.org/10.1038/430742a Sampayo, E. M., Ridgway, T., Bongaerts, P., & Hoegh-Guldberg, O. (2008). Bleaching susceptibility and mortality of corals are determined by fine-scale differences in symbiont type. Proceedings of the National Academy of Sciences of the United States of America, 105(30), 10444–9. https://doi.org/10.1073/pnas.0708049105 Stat M, Carter D, & Hoegh-Guldberg O. (2006). The evolutionary history of Symbiodinium and scleractinian hosts— Symbiosis, diversity, and the effect of climate change. Perspectives in Plant Ecology, Evolution and Systematics, 8, 23– 43. Stat, M., Morris, E., & Gates, R. D. (2008). Functional diversity in coral-dinoflagellate symbiosis. Proceedings of the National Academy of Sciences of the United States of America, 105(27), 9256–61. https://doi.org/10.1073/pnas.0801328105 Szmant, A. M. (2002). Nutrient Enrichment on Coral Reefs: Is it A Major Cause of Reef Decline. Estuaries, 25(4b), 743–766. Tambutté, S., Holcomb, M., & Ferrier-Pagés, Stéphanie Reynaud, Eric Tambutté, Didier Zoccola, D. A. (2011). Coral biomineralization: From the gene to the environment. Journal of Experimental Marine Biology and Ecology, 408, 58–78. Tchernov, D., Gorbunov, M. Y., de Vargas, C., Narayan Yadav, S., Milligan, A. J., Haggblom, M., & Falkowski, P. G. (2004). Membrane lipids of symbiotic algae are diagnostic of sensitivity to thermal bleaching in corals. Proceedings of the National Academy of Sciences, 101(37), 13531–13535. https://doi.org/10.1073/pnas.0402907101

Trappe, J. M. (2005). A.B. Frank and mycorrhizae: the challenge to evolutionary and ecologic theory. Mycorrhiaz, 15, 277–281. University of Maine. (n.d.). How to do a Power Analysis - UMaine Research - University of Maine. Retrieved March 12, 2018, from https://umaine.edu/research/faculty/research-compliance/institutional-animal- care-and-use-committee-iacuc/protocol-review-form/how-to-do-a-power-analysis/ Vega Thurber, R. L., Burkepile, D. E., Fuchs, C., Shantz, A. A., Mcminds, R., & Zaneveld, J. R. (2014). Chronic nutrient enrichment increases prevalence and severity of coral disease and bleaching. Global Change Biology, 20(2). https://doi.org/10.1111/gcb.12450 Vitousek, P. M., Aber, J. D., Howarth, R. W., Likens, G. E., Matson, P. A., Schindler, D.W., … Tilman, D. G. (1997). Human alteration of the global nitrogen cycle: Sources and consequences. Ecological Applications, 7(3), 737–750. Wang, J., & Douglas, A. E. (1998). Nitrogen recycling or nitrogen conservation in an alga-invertebrate symbiosis. The Journal of Experimental Biology, 201, 2445– 2453. Waller, R. G., & Tyler, P. A. (2005). The reproductive biology of two deep-water, reef- building scleractinians from the NE Atlantic Ocean. Coral Reefs, 24, 514–522. https://doi.org/10.1007/s00338-005-0501-7 Walsh, G. E., & Bowers, R. L. (1971). A review of Hawaiian zoanthids with descriptions of three new species. Zoological Journal Linnean Society, 161–180. Wiedenmann, J., D ’angelo, C., Smith, E. G., Hunt, A. N., Legiret, F.-E., Postle, A. D., & Achterberg, E. P. (2012). Nutrient enrichment can increase the susceptibility of reef corals to bleaching. Nature Climate Change, 3. https://doi.org/10.1038/NCLIMATE1661 Winter, A., Appeldoorn, R. S., Bruckner, A., Williams, E. H., & Goenaga, C. (1998). Sea surface temperatures and coral reef bleaching off La Parguera, Puerto Rico (northeastern Caribbean Sea). Coral Reefs, 17, 377–382.

APPENDICES

APPENDIX 1: IMAGEJ FERET DIAMETER CALCULATIONS

Feret diameter, Feret’s diameter, or the caliper diameter of an object is a measurement used for a non-circular object. It is the measurement between two furthest points on the surface of the object. ImageJ was used to calculate this measurement, and in this case, it was used to measure Symbiodinium.

Figure 1. A Hypothetical Symbiodinium

The perimeter of the Symbiodinium was used by the program to calculate Feret diameter. The maximum and minimum Feret diameter is calculated by ImageJ by rotating the object and drawing diameters every 2˚ of rotation. The coordinates of the two furthest points are recorded, and the smallest diameter, the Feret minimum, is located. The angle on which these two lines intersect is the Feret angle (Figure 2). These two Feret measurements were then averaged for each Symbiodiniu, which were used for all size analysis.

Figure 2. The Feret measurements imageJ calculates.

However, ImageJ’s calculation of Feret diameter is not the true Feret measurement, it is a very close approximation. Because the program measures by rotation, there is a slight chance it can miss the true furthest points.

In order to find the true Feret maximum of a small object, in this case a

Symbiodinium, the perimeter of the object must be drawn. Because of the noncircular nature of Symbiodinium, there are places where the cell has concave imperfections, however the only points in the Symbiodinium that matter are convex imperfections, as the calculation is for Feret maximum, and the maximum point will not occur at a concave point. A perimeter can be traced where only the convex points of the cell shape touch the perimeter, and all of these measurements can be compared until the Feret maximum is calculated (Figure 4). This object is called the convex hull.

Figure 4. The convex hull of the hypothetical Symbiodinium.

However, this method is tedious for near-circular objects. In just this simple example, the number of pairs a program would have to remember would be 7 choose 2, or 21. In an object with several hundred convex points, for example, 150, a program would have to measure 150 choose 2, or 11175.

An applicable theorem states that “the diameter of a convex figure is the greatest distance between parallel lines of support” [Theorem 4.18, Preparata and Shamos,

Computational Geometry, 1985] reduces the number of pairs a program would have to measure. A pair of points in an object that follows this theorem is called antipodal vertices (Figure 3).

Figure 4. All of the antipodal vertex pairs of the hypothetical Symbiodinium.

The number of antipodal pairs the program would now have to measure is reduced to four pairs, a much easier number of distances to measure and store. A similar method is used to find the Feret minimum with concave points. Hypothetically, if there was a small number of Symbiodinium, this method could be used to measure the actual

Feret diameter of each Symbiodinium cell.

That being said, there are a lot of data in this project, and that data was measured under ImageJ’s assumptions of Feret diameter approximation, and is assumed to be

“close enough”.

References:

Eddins, S. (2017). Feret Diameter: Introduction. Retrieved from https://blogs.mathworks.com/steve/2017/09/29/feret-diameter-introduction/

Ferriera, T., & Rasband, W. (2012). ImageJ. Retrieved from https://imagej.nih.gov/ij/docs/guide/146.html

APPEDIX 2: R SCRIPT FOR STATISTICAL ANALYSIS

##ANOVA SET I

#Trial 1

#fetching data #transfer all cvs from folder to desktop #note: Feret diameter was misspelled as ‘ferret’ in data column organization #thus ‘ferret’ was used during this program script in place of ‘feret’ setwd("~/Desktop") T1D4 <- read.csv("T1D4.csv", header = TRUE) T1D8 <- read.csv("T1D8.csv", header = TRUE) T1D12 <- read.csv("T1D12.csv", header = TRUE) T1D14 <- read.csv("T1D14.csv", header = TRUE)

#make some boxplots library(ggplot2) ggplot(T1D4, aes(x = factor(Tank), y = Ferret)) + geom_boxplot() ggplot(T1D8, aes(x = factor(Tank), y = Ferret)) + geom_boxplot() ggplot(T1D12, aes(x = factor(Tank), y = Ferret)) + geom_boxplot() ggplot(T1D14, aes(x = factor(Tank), y = Ferret)) + geom_boxplot()

#Trial 2

#fetching setwd("~/Desktop") T2D4 <- read.csv("T2D4.csv", header = TRUE) T2D8 <- read.csv("T2D8.csv", header = TRUE) T2D12 <- read.csv("T2D12.csv", header = TRUE) T2D14 <- read.csv("T2D14.csv", header = TRUE)

#more box plots library(ggplot2) ggplot(T2D4, aes(x = factor(Tank), y = Ferret)) + geom_boxplot() ggplot(T2D8, aes(x = factor(Tank), y = Ferret)) + geom_boxplot() ggplot(T2D12, aes(x = factor(Tank), y = Ferret)) + geom_boxplot() ggplot(T2D14, aes(x = factor(Tank), y = Ferret)) + geom_boxplot()

#removed a bunch of outliers in excel to normalize

#test for symmetry library(lawstat) levene.test(T1D4$Ferret, T1D4$Tank) levene.test(T1D8$Ferret, T1D8$Tank)

levene.test(T1D12$Ferret, T1D12$Tank) levene.test(T1D14$Ferret, T1D14$Tank) levene.test(T2D4$Ferret, T2D4$Tank) levene.test(T2D8$Ferret, T2D8$Tank) levene.test(T2D12$Ferret, T2D12$Tank) levene.test(T2D14$Ferret, T2D14$Tank)

#test for normality shapiro.test(T1D4$Ferret [T1D4$Tank=="A"]) shapiro.test(T1D4$Ferret [T1D4$Tank=="B"]) shapiro.test(T1D4$Ferret [T1D4$Tank=="C"]) shapiro.test(T1D4$Ferret [T1D4$Tank=="D"]) shapiro.test(T1D4$Ferret [T1D4$Tank=="E"]) shapiro.test(T1D8$Ferret [T1D8$Tank=="A"]) shapiro.test(T1D8$Ferret [T1D8$Tank=="B"]) shapiro.test(T1D8$Ferret [T1D8$Tank=="C"]) shapiro.test(T1D8$Ferret [T1D8$Tank=="D"]) shapiro.test(T1D8$Ferret [T1D8$Tank=="E"]) shapiro.test(T1D12$Ferret [T1D12$Tank=="A"]) shapiro.test(T1D12$Ferret [T1D12$Tank=="B"]) shapiro.test(T1D12$Ferret [T1D12$Tank=="C"]) shapiro.test(T1D12$Ferret [T1D12$Tank=="D"]) shapiro.test(T1D12$Ferret [T1D12$Tank=="E"]) shapiro.test(T1D14$Ferret [T1D14$Tank=="A"]) shapiro.test(T1D14$Ferret [T1D14$Tank=="B"]) shapiro.test(T1D14$Ferret [T1D14$Tank=="C"]) shapiro.test(T1D14$Ferret [T1D14$Tank=="D"]) shapiro.test(T1D14$Ferret [T1D14$Tank=="E"]) shapiro.test(T2D4$Ferret [T2D4$Tank=="A"]) shapiro.test(T2D4$Ferret [T2D4$Tank=="B"]) shapiro.test(T2D4$Ferret [T2D4$Tank=="C"]) shapiro.test(T2D4$Ferret [T2D4$Tank=="D"]) shapiro.test(T2D4$Ferret [T2D4$Tank=="E"]) shapiro.test(T2D8$Ferret [T2D8$Tank=="A"]) shapiro.test(T2D8$Ferret [T2D8$Tank=="B"]) shapiro.test(T2D8$Ferret [T2D8$Tank=="C"]) shapiro.test(T2D8$Ferret [T2D8$Tank=="D"]) shapiro.test(T2D8$Ferret [T2D8$Tank=="E"]) shapiro.test(T2D12$Ferret [T2D12$Tank=="A"]) shapiro.test(T2D12$Ferret [T2D12$Tank=="B"]) shapiro.test(T2D12$Ferret [T2D12$Tank=="C"]) shapiro.test(T2D14$Ferret [T2D14$Tank=="A"]) shapiro.test(T2D14$Ferret [T2D14$Tank=="B"]) shapiro.test(T2D14$Ferret [T2D14$Tank=="C"]) shapiro.test(T2D14$Ferret [T2D14$Tank=="D"]) shapiro.test(T2D14$Ferret [T2D14$Tank=="E"])

#boom

#anova time anova.results <- aov(Ferret ~ Tank, data=T1D4) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T1D8)

summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T1D12) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T1D14) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T2D4) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T2D8) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T2D12) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Tank, data=T2D14) summary(anova.results) TukeyHSD(anova.results)

##ANOVA SET II

#bring in the data setwd("~/Desktop") T1 <- read.csv("T1.csv", header = TRUE) T2 <- read.csv("T2.csv", header = TRUE) T3 <- read.csv("T3.csv", header = TRUE) T4 <- read.csv("T4.csv", header = TRUE) T5 <- read.csv("T5.csv", header = TRUE)

#all the data is already normally distributed and symmetrical so anova time anova.results <- aov(Ferret ~ Day, data=T1) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Day, data=T2) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Day, data=T3) summary(anova.results) TukeyHSD(anova.results) anova.results <- aov(Ferret ~ Day, data=T4) summary(anova.results) TukeyHSD(anova.results)

anova.results <- aov(Ferret ~ Day, data=T5) summary(anova.results) TukeyHSD(anova.results)

##now for the final figures

#Experiment 1: Nitrogen Figures setwd("~/Desktop") ND4 <- read.csv("ND4.csv", header = TRUE) ND8 <- read.csv("ND8.csv", header = TRUE) ND12 <- read.csv("ND12.csv", header = TRUE) ND14 <- read.csv("ND14.csv", header = TRUE) library(ggplot2) ggplot(ND4, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot() ggplot(ND8, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot() ggplot(ND12, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot() ggplot(ND14, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot()

#Experiment 1: Temperature setwd("~/Desktop") TD4 <- read.csv("TD4.csv", header = TRUE) TD8 <- read.csv("TD8.csv", header = TRUE) TD12 <- read.csv("TD12.csv", header = TRUE) TD14 <- read.csv("TD14.csv", header = TRUE) library(ggplot2) ggplot(TD4, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot() ggplot(TD8, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot() ggplot(TD12, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot() ggplot(TD14, aes(x=factor(Treatment), y=Feret)) + theme_classic() + geom_boxplot()

#end script

AUTHOR’S BIOGRAPHY

Molly Westbrook grew up in Ithaca, NY with a huge passion for the natural sciences. They went on to attend the University of Maine, majoring in Marine Science with concentrations in Aquaculture and Marine Biology, and minoring in Mathematics.

Molly’s done several things while at UMaine, including co-founding the UMaine Coral

Club and acting as Treasurer for the first year, DJing at WMEB 91.9 fm, performing as a singer-songwriter in the greater Orono area, studying abroad at James Cook University in

Australia, working in many labs, analyzing many datasets, and playing many board games. They had to write a lot of bios when they were a stage manager throughout middle school and high school, and ten years later they’re still not great at it.