Reflections: Hot Crab Summer

Emily Dombrowski, College of Charleston

Reflections: In my first and second posts, I discussed how horseshoe crabs are very important to human health and ecosystems and how I evaluated relative levels of stress in horseshoe crabs. Over the course of 10 weeks, I had the opportunity to explore this project and determine findings. My research questions and methods brought me to the following findings: 

• As temperature increases…
  • Horseshoe crab heart rate increases
  • Healthy immune cells increase
• In younger crabs, as temperature increases…
  • Horseshoe crab amebocyte (blood cell) density increases

Some of my results were expected. We hypothesized that as temperature increased, so would horseshoe crab heart rate. This can indicate higher responses tp stress. The results about blood cells and immune cells were interesting. These indicate that as temperature increases, crabs may be having an immune response. More studies will be needed to confirm these findings, but now we will have a baseline for continued research about horseshoe crabs, age, temperature, and stress!

Gus Snyder, Jody Beers, and I collecting horseshoe crabs on the beach

Overall, this project taught me a lot about the behind the scenes aspects of research. I had thought about all of the work that goes into maintaining research organisms, but I never considered how much time is required to take care of animals. I would spend hours each week feeding and taking care of the horseshoe crabs, and I had 26 pages of notes about water quality that we needed to maintain the crabs. This information was barely mentioned in any presentation or manuscript. The time that I put into this project gives me a lot more respect for other scientific studies with bigger sample sizes and organisms that require more care. 

Additionally, this summer showed me how scientists work together to get their results. My fellow lab members were instrumental to my research process. I needed help collecting horseshoe crabs, gathering data, and practicing my presentation skills. I also had the opportunity to help my peers in the REU program with their projects. I really enjoyed the collaborative nature of our projects. It makes me more confident to continue with science and enthusiastic to see all the work that goes into my peers’ studies.

Acknowledgements:

I would like to thank all members of the Beers Lab: Dr. Jody Beers, Jessica Daly, Augustus Snyder, and Jacob Cashour. Special thanks to Dr. Daniel Sasson and the Department of Natural Resources for collaboration during this project, and to Dr. Robert Podolsky for overseeing the 2021 Fort Johnson REU. This research was supported by the Fort Johnson REU Program, NSF. DBI-1757899.

Algae Microbiome: Health Inspection

Olivia Suarez, College of Charleston

The Approach: In my previous post I discussed the functions of algae, such as oxygen production and water purification, and how the seven species chosen this study are facing the threat of urbanization. Because they are of such importance in an ecosystem, this summer I investigated whether species with different characteristics would differ in features of their microbiome.

The seven macroalgae species in this study were collected from ‘Ewa Beach, Hawai’i on May 22-26, 2021 during morning low tide by Gabbie Kuba, Drs. Heather Spalding, and Heather Fullerton. After washing to remove debris and loosely associated epiphytes and bacteria, they were stored in RNAlater and shipped on dry ice to the College of Charleston. 

Completed gel electrophoresis imaging.

I extracted each algal sample using the MPBio FastDNA Spin Kit for soil. These DNA samples were tested with PCR to confirm bacteria presence on the algae, and DNA concentrations were measured using Nanodrop. The DNA concentrations will be used as a proxy for bacterial abundance in each species’ microbiome. In particular, we predict that the most morphologically complex species, Asparagopsis taxiformis, will have relatively high bacterial abundance per unit weight given its relatively high surface area. We also predict that the invasive species Acanthophora spicifera and Avrainvillea sp. will show relatively high bacterial abundance given their ability to survive in new and disturbed locations.

Acknowledgments

Special thanks to Dr. Heather Fullerton for her guidance on this project as well as Dr. Heather Spalding and Gabrielle Kuba for collecting these species in Hawai’i. This project is supported by the Fort Johnson REU Program, NSF DBI-1757899 and NFWF.

Serving up Seaweeds: The Recipe

Sophie Spiegel, College of Charleston

The Approach: In my previous post, I explained that seaweed could be a culinary solution to climate change. It can be sustainably farmed and an alternative food source to current food systems, which are in decline. To see if Seaweed farming is a possible future venture for Charleston, it is first necessary to ask: Is the seaweed in Charleston safe to eat?

This time, I will be taking you through my recipe to the experiment to help me cook up some final results that are sure to be chef’s kiss. 

Ingredients: Like any great recipe, the preparation process begins with gathering the key ingredients. The same goes for my experimental design, although instead of a fancy brioche and truffle oil, the key to my project was a selection of species and sample sites. I chose three species as my experimental subjects because of their abundance during the summer months. I decided to look at the species Gracilaria tikvahiae because it is native to the southeastern United States. Additionally, there is already a lot of interest in the aquaculture industry surrounding Gracilaria tikvahiae in the United States because it can be used as a food grade agar added to food products as a thickening agent. The second species I looked at is Agarophyton vermiculophyllum. Although it is not commonly eaten in the United States, it belongs to the same family as the popularly eaten species commonly called ogo in Hawaii. Furthermore, because it is considered an invasive species, meaning that it came from other parts of the world and it has the potential to outcompete native species and take over, farming and harvesting this species could be used as an invasive management strategy. Finally, I looked at the seaweed Ulva spp. I used this type of seaweed as a water quality indicator. Seaweeds are fixed to one place in the water; they can absorb the nutrient conditions in the water column over days to weeks and therefore work better at telling the nutrient quality in a particular area than regular water sampling. Regular discrete water sampling is only suitable for measuring one specific moment because water fluctuates with changing tides. Ulva spp. is very thin and flat, so it can filter through many nutrients in the water column, making it a good water quality indicator. 

• 

  Agarophyton vermiculophyllum

• 

  Ulva spp.

• 

  Gracilaria tikvahiae

The other key ingredient to my experiment design was site selection. I had a variety of environments, such as sandy mudflats, jetties, and boat docks, based on the preferred habitat of the species of focus. I also chose sites based on the Department of Health and Environmental Control Shellfish Monitoring Program. In the state of South Carolina, there are currently no regulations for growing seaweed. The Shellfish monitoring program takes water samples along the coast of South Carolina and establishes if waterways are approved or prohibited for oyster farming and shrimping. Shellfish such as oysters absorb similar nutrients from the waterways as seaweeds. My thought process was if these areas are safe enough to grow shellfish, shouldn’t these areas also be safe enough to grow seaweed? I called the approved sites “clean” and compared them to prohibited or “dirty” sites to see if there was a difference in the seaweed found at these sites. 

Map of South Carolina and the seven sample sites color-coded based on “clean” vs “dirty”. Inset map shows the 4 “dirty” sites in Charleston Harbor that are located within the the10B Shellfish Monitoring Program station (shaded).

Cooking Measurements: Another critical part of a recipe is attention to the cooking measurements to get the tastiest outcome. Choosing the correct measures will help me to best answer my question. For this project, I will be focusing on three different measurements. I will be looking at δN15, percent nitrogen, and heavy metals concentrations compared to regulatory limits. δN15 helps to understand the main source of nutrients in seaweed tissue, whether natural, fertilizer, or wastewater. Percent nitrogen measures how much of the nutrients are stored within seaweed tissue. A high percentage of nitrogen and high δN15 value suggests seaweed could be receiving its nutrients from wastewater. If seaweeds receive nutrients from wastewater, the seaweed could be exposed to fecal matter, pathogens, and heavy metals, making it unsafe for human consumption. Measuring heavy metals compared with regulatory limits will determine if there are safe levels of trace metals; otherwise, the seaweeds could be toxic to humans if consumed. 

So now that the recipe is complete, we have to wait to eat the final dish! Literally, in this case because I have to make sure the seaweed is safe to eat! 

Acknowledgements: This research is supported by the College of Charleston and funded by the Fort Johnson REU program, through NSF DBI-1757899. The study is being facilitated at the Grice Marine Laboratory. Special thanks to my mentor Dr. Heather Spalding for her guidance, as well Dr. Bob Podolsky, Elle Pestorius and other staff, faculty, and students at Grice Marine Laboratory.

A Pinch Packs a Punch

Zoe Munson, College of Charleston

The Approach: In my previous post, I outlined how the issue of heightened carbon dioxide concentrations in our environment may pose threats to calcifying organisms, or organisms who need certain minerals from their surroundings to maintain a calcium carbonate-based shell, skeleton, or other bodily structure. Specifically, my interest is piqued by how these environmental trends can affect a crab’s muscular and exo-skeletal strength, which relies on their ability to recalcify a protective layer around their body directly following a molt. So how does one determine how well a crab regains its strength after a molt in different environmental conditions?

Crab collection using a seine net. Photo taken by Dr. Robert Podolsky

I began my experimental design with periodic collections of ornate crabs, Callinectes ornatus, using a seine net off of Fort Johnson, South Carolina in the Charleston harbor in order to have a steady income of readily molting specimens. Once the crabs are identified, I take initial measurements of the size of their body and claws in millimeters, the force of their pinch on a reading pad in newtons, and the stiffness of the exoskeleton cuticle of their non-pinching claw using a force gauge that is connected to analyzing software. By taking initial recordings of the collected crabs’ performance in these aspects before they molt, I am able to determine a baseline that can be used to compare the progression of their growth then following a molt.

Crab pinching measurement using the force reading pad. Photo taken by Dr. Robert Podolsky

Once a crab molts, they are placed in one of two holding containers that are connected to a controlled flow of gas; one at 400ppm carbon dioxide, or the estimated current level of carbon dioxide found in the ocean, and the other being at 1000ppm carbon dioxide, or 2.5 times the estimated current level. I then keep a close eye on them and repeat measurements of their pinching force twice a day for each day after they are found molted, between 9-10 AM and 5-6 PM. I also repeat the process of measuring their cuticle under the force gauge once a day during the afternoon check-in time to track how far along they are in their post-molt growth process.

High and Low carbon dioxide concentration containers. Photo taken by me

By comparing the rate and magnitude that a crab’s strength and shell hardness progresses after a molt when placed in higher and lower levels of carbon dioxide, I am able to see the extent to which environmental conditions can have an impact on the functions in a crab’s life. Does our changing planet pose threats in the future for the health of coastal organisms?

Acknowledgements: This study is made possible by the Biology Department at the College of Charleston and funded by NSF through the Fort Johnson REU Summer Program. It is being facilitated at Grice Marine Laboratory in Fort Johnson, South Carolina. Special thanks to faculty mentor Dr. Podolsky and other building faculty/staff/students who contribute aid to the research or the collection of crabs.

Scouting Out the Competition

Elle Pestorius, College of Charleston

The Approach: In my previous blog post, I discussed the general problem of resilience in invasive species, and the main question of how invasive algae compare to native algae. First, we need to understand the importance of finding the differences in physiological characteristics of these algae species. The algal physiology being studied is looking to try to figure out how the invasive algae functions in order to adapt and thrive within such a variation of habitats, and whether it outcompetes the native species. These variables will help to figure out who copes better within different environments. The specific physiological functions I am examining are photosynthetic efficiency and nutrient storage. So, how do these species of algae compete with one another?

Photosynthetic efficiency is very important to study because photosynthesis is a necessary function that allows the alga to produce energy for themselves to survive. I am looking to see which species, the invasive Agarophyton vermiculophyllum or natives Gracilaria tikvahiae and Ulva spp. found, can maximize productivity with the least amount of expenses of energy sources. This process uses a PAM (pulse amplitude modulation) fluorometer to measure photosynthetic variables, like light level maximal, rate, yield, and maximum capacity, of photosynthesis. A light pulse is given that mimics the sunlight conditions in which that specific alga was found, and data is collected from the alga’s response which is determined by the absorption of this light. Overall, this data shows which species is most photosynthetically efficient.

Algae also absorb nutrients from the water that allow them to grow. They can either utilize them right away for rapid growth or store them for future use. The storage of nutrients can be found within the alga’s tissue content. To determine their ability to store nutrients the algae samples are dried, ground, and shipped off to the Biochemical Stable Isotope facility at University of Hawaii at Manoa. By looking at the percentage of nitrogen found within the tissues of the different species, I can figure out who stores nutrients the best for future uses like growth, survival, and adapting to changing environments.

If we can figure out if the invasive algae have any special abilities that allow them to succeed and dominate the waters, we may be able to help find solutions to protect local biodiversity and prevent the spread and initial introductions of invasive species in the future. In order to conduct this research, there are many techniques and methods that lead up to analyzing data and answering these important questions.

Acknowledgements

Special thanks to my mentor Dr. Heather Spalding and lab mate Sophie Spiegel, as well as Dr. Podolsky and the rest of the REU interns. This research is funded by the Fort Johnson REU program, through NSF DBI-1757899.

What time is it? Mud Time Woo!

Charles Taibi, College of Charleston

The approach: In my previous post I discussed the importance of the salt marshes for coastal cities and seafood connoisseurs like myself. The ability for salt marshes to support the life that lives within hides in the base of the entire system, the mud, and what grows in the mud, the marsh grass. To understand how these two parts of the marsh go together I will be analyzing the mud in one part of my study and then the examining the physical properties and profile of the marsh. Hopefully by the end of this summer, I will be able to find a link between these two components of the marsh.

This summer I am doing an entire swath of analyses, so for timely purposes I will simply list them in this article. Of the listed analyses we can break them into two separate groups: Soil content analysis and marsh profile. Within the soil analysis we are looking at Ammonia, water content, organic content, reduction potential, salinity, pH, bulk density, and particle size. For the marsh profile we will be considering elevation, distance from creek, marsh grass plant density and marsh grass height. After reading that list you may be thinking: ‘how on earth are you going to do these analyses over the course of one summer? Well, I am getting a lot of help from the other REUs as well as the professors I am working with on this project.

pH probe used for measuring pH of the mud as well as the reduction potential

Quadrats used for sampling plant density and plant height.

Though I will not go through the grueling details of each and every analysis, I will still run you through what a typical day looks like in the marsh. I begin by selecting the marshes I will visit. Then I collect all my materials to do my in-situ observations, a transect, quadrats, ORP redox probe, meter stick and cups to collect mud. In the field I first lay down a transect spanning the marsh from the creek to the middle of the marsh, inland. I then collect samples of mud at each ends as well as record plant height and plant density. While this is all going on, I am recording the reduction potential of the soil as well as the pH using the redox probe. Once I am finished recording all the data for that specific marsh I collect all my materials and head out to the next marsh for more mud.

The mud collected in the field is then dried at 50^oC for a week and then analyzed for one of the many factors listed above. Though my time in the marsh was brief, I have built respect for the muddy arenas and the complexity that is within them  

Acknowledgements: I would like to thank my mentors Dr. Erik Sotka, Dr. Theodore Them, and Dr. Scott Harris for their continual support and guidance throughout this project. I would also like to thank my fellow REU interns for their help in the field and lab. Thank you the NSF for providing me with the funding required to carry out this research.

Tadpoles, Tadpoles, and Even More Tadpoles

Regan Honeycutt, College of Charleston

One of the southern toads collected for this research. This toad is named Jenny.

The approach: In my previous post, I spoke of a new threat to amphibians: salt. We know that amphibians aren’t doing so well with the sudden environmental changes that come with climate change. We also know that amphibians are an indicator species meaning that their health represents the health of the entirety of their ecosystem because of their extreme sensitivity to change. This means that what amphibians are experiencing now will most likely affect other organisms in the future. I am studying how the exposure to salt during early life stages impacts amphibians later in life, almost like how childhood trauma affects adult humans. By answering this question, we can better understand what we can do to care for these animals in case removing them from the harmful environment isn’t enough.

But, how am I trying to answer it? I am raising hundreds of cute little toads! My lab mentor, Dr. Allison Welch, lab mate, Aubrey Anthony, and I collected 7 pairs of southern toads that laid eggs in our lab. We ended up using 3 clutches, or groups of eggs, in the experiment. Each clutch can have anywhere from 2,500 to 4,000 eggs each! I took 600 eggs from each of the 3 clutches and separated them into 3 different solutions. One is a control solution that has a salinity similar to a freshwater pond. The other two are saltier with 4 parts per thousand salt (PPT) and 6 parts per thousand salt (PPT). This means for every 1000 water molecules there are 4 or 6 salt molecules respectively. For reference, ocean water is about 28-30 parts per thousand salt (PPT), so the water that I put these eggs in isn’t that salty at all! 

Some of the southern toad eggs laid in the lab. This species of frog lays eggs in long strings with a clear jelly connecting them all. Photograph by Regan Honeycutt

Once the eggs hatched into tiny tadpoles we moved them to larger containers, but we switched the solutions around! Some of the tadpoles continued in the same solution (went from control to control or from 4 PPT to 4 PPT). Others went from control solution to 4 PPT to simulate an environment that gets saltier over time, and finally some went from 4 PPT to control solution to simulate an environment that gets less salty over time. 

One of the tadpoles in the experiement

As of right now, I am simply raising these animals and monitoring their growth, but once they grow their legs and become fully fledged toads we will measure the differences in sizes, leg length, jump length, and number of jumps between the groups of toads based on what solutions they were exposed to. We can equate their ability to move with how well they would do in the wild. Movement is extremely important in these animals’ lives as it is required to escape predators and catch food. If exposure to salt early in life impacts a toad’s ability to move in the future, we would then know that simply removing the animal from the harmful situation will not solve the overarching problem.

I would like to thank my mentor Dr. Allison Welch and my lab mate Aubrey Anthony along with Dr. Bob Podolsky and the rest of the REU interns. This research was supported by the Fort Johnson REU Program, NSF DBI-1757899.

Let’s Rumble: Sinking Our Teeth Into Science

Christian Simmons, College of Charleston

The approach: In my Previous Post, I discussed two top predators of coral reefs, sharks and moray eels, and how they maintain the order and balance of reef fish populations. Even though these two animals live in the same environment, or neighborhood, they do not like to share food with each other and as a result fight. Sharks have triangle shaped teeth with rough saw-like edges and small tooth-like scales. Moray eels have long, slender, and curved teeth and loose and slimy skin. With different physiological traits, the question posed is who is better at biting and fighting? This post will sink our teeth into the kind of data that I collected and how the various comparisons made will help me determine which top predator is the underwater biting fighting champion. 

Spotted Moray Eel (Gymnothorax moringa) with skin sample removed

Starting off, you’re probably wondering how I get an Atlantic Sharpnose Shark (Rhizoprionodon terraenovae), a Spotted Moray Eel (Gymnothorax moringa), and a Purplemouth Moray Eel (Gymnothorax vicinus) to fight each other. Well, while it would make for sensational tv, the animals used in my project are dead specimens. Although not alive, having actual skin samples is very important to testing which animal can resist being bitten more. The reason for this is because skin is a composite material, meaning it is made up of more than one thing. Skin has three layers, epidermis, dermis, and hypodermis (in order from outside to inside). The middle layer, the dermis, is made up of collagen and elastin that allows skin to be stretched greatly. When a shark bites a moray eel or vice versa, that animal has to produce enough force, or have enough strength in its jaw, to puncture through all three layers. 

Metal ASTM (American Society for Testing and Materials) Probe

Now that we’ve covered that, here’s how I do it. With the skin samples secured over a hole, I can then use a metal probe, or stake, attached to a force gauge to measure the amount of strength it takes to puncture through. As the probe is lowered into the sample I am able to record the data on a computer and view how much force it takes and how far the skin stretches before it breaks. I repeat this process for all of the shark and moray skins and then compare them to come to a conclusion. 

Your next question may be, how do these comparisons help me come to a conclusion? Simply put, the more force it takes for the probe to puncture the skin and the further the distance that the skin stretches, the more resistant the skin is. Using this logic, I can look at the data for each animal and determine which one has the highest values for force and distance. Making comparisons are important in understanding the role, behaviors, and interactions of these animals. More importantly, soon these comparisons will help me to crown the ultimate underwater biting fighting champion.

Acknowledgments

I would like to thank my research advisor, Dr. Andrew Clark, and lab partner, Melanie Fischer, for your assistance on this project. Thank you to the National Science Foundation, College of Charleston, Grice Marine Lab, and South Carolina Department of Natural Resources. This research was supported by the Fort Johnson REU Program, NSF DBI-1757899.

How to listen to a crab’s heart, and other super useful skills

By Emily Dombrowski, College of Charleston

The Approach: In my Previous Post, I discussed how horseshoe crabs play a vital role in human medicine and how the human harvesting industry affects crab populations. Similarly, I talked about the importance of reducing stress in the blood taking procedure while maximizing the quality of blood. I will be researching this by analyzing crab blood, heart rate, age, and sex. So, to start, how do you tell how old a horseshoe crab is? How do you take it’s heart rate, and look at it’s blood? 

In order to get information about crabs and temperature, we gradually reduced or increased tank temperatures and held the crabs at the desired temperature for a week before taking any measurements.

For our study, we are interested in how different age groups of horseshoe crabs react to blood draws. In order to assess this, I used shell darkness, the amount of mucus, and the amount of bacterial degradation to the shell to group crabs into “older” and “younger” categories. 

Horseshoe crab with infared heart rate monitor attached to pericardial membrane (Photo credit: Emily Dombrowski, 2021)

Unlike us, horseshoe crabs have a hard, outer carapace that doesn’t expose veins. In order to draw blood from crabs, we must “fold” them in half over a hand made mount. This exposes a small area called the pericardial membrane that connects to a vast network of veins and arteries. It’s like a central train station that all trains arrive at before being shuttled around a city. Instead of hunting for a small vein, we can draw blood directly from this membrane. 

Similarly, this is the location where we can take a crab’s heart rate. We do so by attaching a small infrared heart rate monitor to the crab with mounting putty. When there is a large amount of blood in a vessel, a different amount of light is reflected back to the monitor than when a small amount of blood is present. Using these differences and various calculations, the monitor can calculate heart rhythms and beats per minute from the crab. 

Infrared heart rate trace from horseshoe crab (Photo credit: Emily Dombrowski, 2021)

Using these strategies, along with counting relative amounts of blood cells in horseshoe crab blood, we will be able to look at how age, sex, and temperature affects stress and blood quality in horseshoe crabs. Heart rate values will give us a way to quantify stress, while counting blood cells will give us information about overall blood quality. Age and sex will then be used to look at how crabs held in hot and cold tanks are potentially reacting differently, and how we can alter horseshoe crab care procedures to better suit these crabs. These different metrics will help us answer our central research questions.

Acknowledgements:

I would like to thank all members of the Beers Lab: Dr. Jody Beers, Jessica Daly, Augustus Snyder, and Jacob Cashour. Special thanks to Dr. Daniel Sasson and the Department of Natural Resources for collaboration during this project, and to Dr. Robert Podolsky for overseeing the 2021 Fort Johnson REU. This research was supported by the Fort Johnson REU Program, NSF. DBI-1757899.

Methods to the Sponge Madness

Jake Kuenzli, College of Charleston

The approach: In my previous post, I spoke about the possible environmental consequences that could arise if sponges were consuming and/or modifying DMSP. But the question still remains, how could one see what is flowing through the holes of a sponge. To test this, four species of sponges were collected around the NE section of Looe Key Preservation area in the Florida Keys. These were Iotrochota birotulata, Callyspongia aruleata, Aplysina cauliformis, and Ircinia felix. The sponges were then hung from string with chip-clips and placed into tanks (pictured below). A tank without a sponge that had a string, and a chip-clip was included to act as the control. All tanks were subjected to running water of around the same pressure, except around the times of sampling. At collection periods, the pumps would be turned off to allow the water in the tanks to settle. Water samples were also collected around a depth of 25ft at the sampling site to determine the amount of DMSP present in the environment where the sponges were collected. With this set-up, I was able to run different experiments to look at DMSP, DMS/VOC, and tissue samples. The experiments were carried out over three days, each with a new sponge specimen.

Experimental set-up (Photo credit: Jake Kuenzli)

Gravity filtration rig (Photo credit: Dr. Peter Lee)

DMSP Protocol: To determine if this chemical compound was being affected by the sponges, I needed to see if the seawater chemistry in the tanks was being altered. First, a large amber bottle was used to collect 150 mL of water from each tank. This would be how the total and dissolved samples were gathered. 10 mL of the water were placed into scintillation vials for the total samples, as they were not filtered. For the dissolved samples, 50 mL of water was run through a gravity filtration unit (pictured to the right) and 10 mL of the filtered water was placed into other scintillation vials. Both of these were treated with 100 μL of 50% sulfuric acid and then refrigerated.

DMS/VOC Protocol: DMS (dimethylsulfide) and VOCs (volatile organic compounds) are other products that sponges could be producing. To see if these were present, the 40 mL left over from the filtration units were placed into amber vials. These were then frozen to make sure the organic compounds were preserved.

A. cauliformis being sectioned (Photo credit: Dr. Chris Freeman

Tissue Sampling Protocol: At the end of each day, each species of sponge was dissected (pictured to the left). There was the full section (outside and core of sponge), the outside layer, and the core of the sponge. Each section was placed into cryovials and frozen using liquid nitrogen. These would be used to see what was present in the bacteria living inside the sponges.

Acknowledgements: I would like to thank my mentors Dr. Peter Lee and Dr. Chris Freeman for their continued guidance, as well as Grice Marine Lab, Hollings Marine Lab, and Mote Marine Lab for allowing me to carry out my research. This research was supported by the Fort Johnson REU Program, NSF DBI-1757899.