Undergraduate Bachelor of Science, Elasmobranch Research Team, Florida Southern College
Advisors: Dr. Gabriel Langford and Dr. Allison Durland-Donahou
When most people think of sharks, they picture a scene out of Jaws with a big Great White shark in clear blue oceans. However after this summer when I think of a shark, I think of my little friends: the juvenile or neonate bull sharks and cownose rays we worked with during our research. This research was done to begin bridging gaps in knowledge of brackish waters and the roles they play in elasmobranch development. Hillsborough Bay, FL, is our study site and serves as a nursery ground for bull sharks, cownose rays, and other elasmobranch species. The FSC Elasmobranch field team is led by Dr. Gabriel Langford and Dr. Allison Durland-Donahou, while the lab team is led by Dr. Melanie Langford. Using 100 and 300-ft gill nets to catch our specimens, we are able to tag them with PIT tags, dart tags, and acoustic tags, and collect fin clip samples and skin swabs. Another tool our team uses is acoustic receivers that are able to collect data from our acoustically tagged elasmobranchs, which are placed in eight different locations throughout Hillsborough Bay that coincide with our sampling locations (Figure 1).
Figure 1: Locations of the eight receivers our team has placed in Hillsborough Bay, Florida. Each location is intentionally placed in areas that we have examined our target species and have successfully sampled them as well.
Our research has not gone without its hard moments, like most cases of field work you can not predict everything. In 2021, a lethal red tide event occurred in Tampa’s waters that damaged seagrass beds and more, along with deeply impacting the Southern and Atlantic stingray populations that were found in Hillsborough Bay. This past summer, we saw a return in both species and even happened to collect a few. Another struggle was the warmer summers that Tampa has seen over the past few years. The way we were able to combat that this summer was placing all of our collected animals into a bin with a water pump that was either placed in their mouth or the bin (Figure 2). This pumped in new water which was cooler than the surface of the boat and on extra hot days we put ice into the bin to give the animals a cooler experience. We found that this increased the success of the animals swimming away sooner since they wouldn’t have to recover from the heat once placed back into the bay.
Figure 2: A bull shark placed into a bin with a pump in its mouth during a workup. The pump has two functions: pump oxygenated water over the shark’s gills, and bring in fresh water from the bay that will help to cool off the shark while on the boat.
This research is intended to continue for many more years and hopefully continue to provide information about the nursery ground and habits of bull shark pupping. The receivers will hopefully continue to inform our team on how long the tagged elasmobranchs are staying within the safe home waters of Hillsborough Bay before moving on to the big open ocean.
It’s funny how life works out. I started at Nova Southeastern University in 2020 to earn my B.S. in Marine Biology thinking it would just be about science – long hours in the lab, diving into data (and sometimes the ocean), and learning how life thrives in one of the most complex environments on Earth.
During my third year, I was offered a spot in the Fisheries and Avian Ecology Lab, led by Dr. David Kerstetter. For the next two years I researched fish physiology and how fish survive constantly changing environmental conditions, though, in the end I learned more about how fish DON’T survive constantly changing conditions. More importantly, these past two years, now going on three, I’ve unexpectedly found a second family. We’re a weird bunch for sure, my lab mates and I, but anyone would have to be at least a little weird to willingly dig through fish and bird guts on a professional level. Somehow, we all fit together perfectly, though our backgrounds separate us, our passion for the science bring us together.
After only one semester as a master’s student, it’s been made very clear to me that grad school is no walk in the park. Between coursework, lab work, and the pressure to strive for more, I’ve had my fair share of stressful moments – long nights when I felt my progress was moving backwards and days when I questioned why I was doing this at all. Those were the times when my lab family came through for me.
What makes this lab special isn’t just our shared passion for marine science; it’s that we care about each other beyond the walls of the lab. We’re more than just researchers, we’re friends who check in on each other, notice when someone’s had too much coffee and not enough sleep, and know when to push someone to take a break. Grad school can feel isolating at times, but having a lab family like mine makes all the difference. Sure, we have our squabbles, some real and some caused by sleepless delirium, but at the end of the day, we know we’ve got each other’s backs.
As I write this I’m sitting in our lab’s conference room, surrounded by friendly faces, and am only an hour from our start-of-the-semester lab get together at a local brewery where we’ll enjoy time with each other as friends separate from the lab.
My name is Angela Barreto and I am currently pursuing my master’s degree in marine science at Nova Southeastern University. I graduated with my bachelor’s degree in marine science and a minor in political science at Coastal Carolina University. At Nova Southeastern University I am apart of Dave Kerstetters Fisheries and Avian Ecology Lab. In this lab I am conducting research regarding trace metals on stingrays in the Indian River Lagoon. The Indian River Lagoon is a shallow lagoon in which is heavily affected due to human populations. In this study we plan to look at naturally occurring toxic nonessential metals/metalloids such as mercury, cadmium, arsenic, lead, silver, as well as essential metals/metalloids such as selenium, iron, copper, chromium, zinc, manganese, and nickel. Uptake of radioisotopic metals in oviparous elasmobranchs demonstrated that cartilaginous fish may be more susceptible to toxic metal accumulation because of higher rates of metal uptake (Jeffree et al. 2006b). To be able to understand if these stingrays are displaying toxic levels of these trace metals, we will take a closer look by dissection of the liver and brain. We are also interested as to whether or not there is a concentration of toxic metals involved with maternal offloading specifically due to the uterine villi, and histotroph more understood as the mixture of nutrients and substances in which assist to support the embryo. Looking at off-loading from the mother to pups stems from the idea that the egg case of oviparous sharks may act as a reservoir for radioisotopic metals, and therefore a potential source of radiation exposure to developing embryos (Jeffree et al. 2006a, 2006b; Jeffree, Oberhansli, and Teyssie 2008). With this study we aim to answer any questions we may have as well as be able to draw conclusions and comparisons regarding sex and weight. Additional research on elasmobranchs and trace metals is valuable because there is possibility for there to be behavioral alterations, emaciation, cerebral lesions, and impaired gonadal development. Specifically shown in spiny dogfish to be capable of dramatically inhibiting DNA. Toxic levels could lead to significant declines in male shark fertility. Toxic effects of Hg have also been observed in spiny dogfish rectal glands in which mercuric chlorine can inhibit chloride secretion (Silva, Epstein, and Solomon 1992; Kinne-Saffron and Kinn 2001; Ratner et al. 2006). Thank you for your consideration as this grant can further our understanding and insight as research progresses and as this study takes shape.
Movement strategies, or syndromes, are suites of correlated movement traits shaped by an individual’s responses to environmental cues and cognitive abilities (Abrahms et al., 2017; Spiegel et al., 2017). The movement strategies selected by animals often shift spatiotemporally along gradients of environmental heterogeneity and predictability which has broad implications for species interactions, population dynamics, ecological niches, and ecosystem function (Riotte-Lambert & Matthiopoulos, 2020; Teitelbaum & Mueller, 2019). Most research on movement strategy selection has focused on terrestrial ecosystems, leaving aquatic environments comparatively underexplored. This gap is primarily due to the technological challenges of tracking animal movements underwater. Those of you working with acoustic telemetry data, particularly from non-gridded gate or curtain-style arrays, know that the resolution of a tagged fish’s movements can often be coarse.
Gridded VPS-configured arrays provide finer resolution movement data, making them well-suited for classifying animal movement states using methods like Hidden Markov Models (HMMs) or State-Space Models (SSMs). These models rely on the more continuous tracking of movements to assess characteristics such as step lengths and turn angles (Whoriskey et al., 2022). In contrast, due to the lack of tracking continuity offered by gate or curtain designed arrays, HMM’s or SSM’s are challenging to apply to tracking data from non-gridded gate and/or curtain arrays. If you’re working with data collected from such arrays and want to classify the movement strategies of your fish, here is a potential method, complete with linked example code and data. While this approach is particularly suited for riverine environments, it may also apply to open systems where fish can leave and return to the array but require a little more fine-tuning and thought.
Step 1: Data Wrangling
Start by filtering and cleaning your raw detection data. Remove false detections and exclude fish that don’t meet your minimal threshold criteria. Next, transform the raw detection data into pseudo-continuous data. Detection data from non-gridded arrays often include lengthy intervals between detections, which can span days or weeks. Pseudo-continuous data bridges these gaps by assuming the fish’s location between detections, based on its last known location. For example, if Snook 5445 was last detected in River Zone A at 10:00 a.m. and detected again in the same zone 24 hours later, we can assume it remained in Zone A during that time. However, this assumption is only supported by knowledge of the species’ behavior, range testing of the array, and by taking a deep dive into the data. The pseudo-continuous tracking data will help us calculate movement metrics that are more accurate to how the fish actually moved than if we had just used the raw data alone. This step will take some serious thought and consideration as to how to best apply to your own movement data, which again, means that it’s important range test and evaluating the efficiency of your array. If you’re working in an open system, you may want to consider adding time cut-offs to your coding for pseudo-continuous data. For example, the two systems that I’m conducting my dissertation work in have bays that open to Florida Bay making that portion of the array “open”. The way we designed the receiver array makes it challenging to determine whether a tagged fish has left the system and entered Florida Bay or if it still remains in our focal system outside of the detection range of a receiver. To account for this uncertainty, I decided to add a threshold cut-off to the pseudo-continuous data so that if a fish was detected in a bay, then not heard from again for at least 7 days, I assumed the fish had left the system. A detection within the bay following the 7 day threshold would then began a new event and start to the pseudo-continuous dataset again.
Step 2: Calculate Movement Metrics
Because movement strategies are characterized by suites of correlated traits, the next step is to calculate movement metrics that will characterize the traits of your fish’s movement. For non-gridded acoustic telemetry data, metrics like residence time, net squared displacement, and mean time-to-return are most applicable and similar to those used in higher-resolution satellite tag studies such as Abrahms et al. (2017). For my work, I’ve focused movement metrics on residence time and movement frequency (somewhat of a proxy for net displacement). These metrics were quantified by first calculating movement events. A movement events occurred when a fish was first detected in a zone within my array and the residence time was the duration of time that movement event lasted before the fish swam into a different zone, starting a new movement event and residence time duration. This is also the point where you will need to define your temporal window (e.g., weekly, monthly, or seasonally) interest before summarizing the metrics. The movement metrics I have ended up focusing on are the average, max, and min residence times of an individual’s movement events, the variance of all of that individual’s residence times, the total number of movement events an individual made, and the proportion of system it used.
Step 3: Clustering
Now you should have summarized movement metrics for each of your fish over the temporal window of your choosing. After summarizing movement metrics for each fish, use clustering methods to classify distinct movement strategies. The best method of clustering and optimal number of clusters is determined by the silhouette coefficients and gap statistics. If done properly, each cluster will represent a distinct movement strategy. For my data, k-means clustering with three groups consistently provided the best results across various temporal windows (season and month; Figure 1). However, the choice of clustering method and number of clusters will depend on your fish’s movement patterns, selected metrics, and array design.
Figure 1. Scatterplot of movement metric summaries based on PCA-defined axes with the movement strategy groupings from k-means clustering (ncluster_1 = 204, ncluster_2 = 65, ncluster_3 = 57). Each point is the summary of one fish’s movements during a quarter season.
Step 4: Name Your Clusters
After you’ve run cluster analysis, you’ll pull the cluster groupings and join them back to your movement metric dataframe. I’m sure there are a lot more intricate, and likely better ways to determine what makes each of your movement clusters distinct such as simulation (again see Abrahms et al., 2017) or perhaps generalized linear models, but my method was to average the movement metric values by clustered group, z-score the data, then create a heat plot using geom_tile to see how each group’s movement metrics differed. Because the data are now z-scored, positive values indicate above average, negative, below average, and 0 represents average for that metric. If things are still not very clear, it may help to also join your cluster groups back to your pseudo-continuous dataframe and create abacus plots or movement history plots.
Figure 2. Z-scored movement metric summaries used for a-posteriori classification of k-mean movement strategy groupings. Group 1 was classified as the Transient movement strategy, group 2 as Mixed, and group 3 as the Resident movement strategy.
Conclusion
This method for classifying fish movement strategies from non-gridded acoustic telemetry data is far from perfect but has yielded exciting results when paired with environmental data and stable isotope analysis. I hope that this post maybe piques some interest or better yet, ideas on how to improve methods of classifying the movement strategies of fish using the course resolution tracking data that we often collect from non-gridded acoustic arrays. If you’re interested in trying it out, check out the example code and data provided at: https://github.com/CodyEggenberger/Move_Strat_Classification.
MS Student, Tropical Aquaculture Laboratory, University of Florida
Advisor: Matthew DiMaggio
When discussing the topic of aquaculture, most people would think of vibrant ornamentals or large saltwater food fish. However, many are unaware of the diversity of aquaculture science and the utilization of fish as model organisms. In many cases, lab animals are thought of as rats or other small rodents. However, there are various species of fish that have become extremely useful as model organisms in the fields of genetics and medical science. The Zebra Danio (Danio rerio) has been one of the most prominent lab cultured research organisms, since the 1980’s. Their development is rapid and large quantities of these freshwater fish can be maintained in a small spatial footprint, when compared to rodents. Zebra Danio share a wide range of molecular and genetic similarities to a high percentage of vertebrates, including humans. More fish have been phased in as model organisms since then, with each species having its own unique life history and physiology, which can be leveraged for specific research goals.
Since graduating from Roger Williams University in 2022, I have been able to gain first-hand experience working with fish as model organisms from the Environmental Protection Agency (EPA), in Narragansett, Rhode Island. The Mummichog (Fundulus heteroclitus), also known as the Atlantic Killifish, is the model organism of interest at the EPA, as its life history is similar to the Zebra Danio, in terms of its rapid development, clear chorion, and low maintenance culturing properties. The major difference between utilizing Mummichogs in research is that they are native to brackish waters of the east coast rather than freshwater, like Zebra Danio. Mummichogs are an important baitfish in marine and estuarine environments, meaning that their abundance and distribution directly affect higher trophic level predators and the stability of food webs. The EPA utilizes uses Mummichogs to conduct embryonic PFAS (perfluoroalkyl and polyfluoroalkyl substances) exposure assays as a method of understanding the biological effects of these forever chemicals on organismal development. The work I contributed to featured growth, survival, heart rate, swim bladder measurement, and behavioral endpoints as a method of quantifying PFAS induced effects.
Mummichogs – Environmental Protection Agency in Narragansett, Rhode Island
Since my time at the EPA, I have decided to move south and pursue a master’s degree at the University of Florida’s Tropical Aquaculture Lab. Here I will be working with Mummichogs, Zebra Danio, and Green Spotted Pufferfish (Dichotomyctere nigroviridis). Rather than focusing on PFAS effects, my current work focuses on SREB receptors (Super-Conserved Receptors Expressed in Brain) and their potential ligands, in relation to fish reproduction. It is my hope to uncover more information about how these receptors operate and how their proposed ligands might be useful as potential spawning inducers. This master’s position will allow me to learn more about molecular biology and develop new skills. I hope to learn how the fusion of molecular biology and aquaculture science can benefit the industry and lead to innovation.
The Green Spotted Pufferfish has recently become a more prominent model organism as it possesses one of the smallest vertebrate genomes, making it a desirable research organism for genetics work. These fish are also native to brackish habitats, but they have no large aquaculture producers, meaning wild caught populations are the main supply for both the research and aquarium industry. University of Florida has bred these fish in captivity using an ovarian lavage technique, but there are still many hurdles that stand in the way of a large-scale aquaculture production effort.
Green Spotted Pufferfish – University of Florida Tropical Aquaculture Lab in Ruskin, Florida
The aquaculture industry is constantly expanding, which paves way for increased innovation and collaboration with different fields. The fusion of both molecular biology and aquaculture science paves way for a great deal of research opportunity. I hope to learn more about these species and gain a better understanding of the intricacies involved with fish reproduction on a molecular scale. The aquaculture industry will only grow in importance in the coming years as its utilized to meet the demands of both the food-fish industry as well as the aquarium trade.
My name is Mia Braun, and I am a senior at Florida Southern College, majoring in Marine Biology. My passion and appreciation for the ocean began the day I put snorkeling gear on. This quickly extended to scuba and my life was forever changed. After each dive trip, I grew fonder and fonder of life below the ocean’s surface. These experiences ultimately led me to Florida Southern. Throughout my time at FSC, I have gained more incredible experiences and knowledge than I could have ever imagined.
Last March, I had the opportunity to take a trip to Roatán, Honduras with a tropical ecology class. In Honduras, we expanded our classroom knowledge of coral health and restoration, as well as dipped our toes in the water with scientific diving techniques. Through a series of presentations given during our time there, we were provided valuable information on a variety of topics. Those topics included: coral reef monitoring methods, coral species identification, coral bleaching assessments, coral restoration/outplanting methods, along with the importance and significance of each. It was here that I truly learned the desperate need for conservation and restoration of reefs worldwide. As with all science, there are many different considerations, assessments and precautions required to best protect these vital ecosystems – I gained new insight into these factors and how fundamentally important they are.
Some of the most riveting moments for me were cleaning the coral nursery trees and outplanting coral fragments onto the reef. While on scuba, we were given the opportunity to scrub and clean algae off the PVC pipes containing Acropora palmata and Acropora cervicornis fragments. Using steel bristle brushes we were able to successfully remove the majority of the algae and reduce competition for the growing Acropora fragments in the nursery (Figure 1). Additionally, we got the chance to outplant coral fragments onto the reef using marine epoxy. This unique and impactful procedure also required extensive evaluation to minimize disturbances and maximize survival and establishment. Furthermore, we performed and enhanced coral identification skills, coral health assessments, and other research methods. These events undoubtedly impacted my admiration for not only coral restoration but also coral ecosystems, and marine science as a whole.
Figure 1: Me contributing to the maintenance of the coral nursery at the Roatan Institute for Marine Sciences.
Figure 2: Coral fragments that we outplanted.
Figure 3: Coral identification exercise indicating a Boulder Star Coral.
MS Student, Tropical Aquaculture Laboratory, University of Florida
Advisor: Jeffrey Hill
My name is Meghan Eaton, and I am in my first year of my master’s degree at the University of Florida’s Tropical Aquaculture Lab (TAL) in Ruskin, FL. I am studying the cold tolerance of freshwater ornamental fish under Dr. Jeff Hill and Dr. Quenton Tuckett. I obtained a BS in Marine Science from the University of Delaware in 2021. My research is focused on nonnative species and their potential distribution.
I use a recirculating system (nicknamed the Mastodon) to manipulate the temperature of individual tanks. The original Mastodon was donated by the Florida Wildlife Conservation Commission to the TAL in 2012. This system allowed researchers at the TAL to evaluate the thermal tolerance of fishes, including the chronic lethal minimum temperature (CLmin). Chronic trials consist of lowering or raising the temperature of the water slowly, often 1°C a day. There are three major endpoints that are recorded during a trial- cessation of feeding, loss of equilibrium, and death.
The original Mastodon system consisted of an external header tank that was chilled to 14°C. The chilled water was gravity fed to eight 190-L tanks. Each tank was equipped with a 500-W electric heater to maintain the temperature in each tank individually. Temperature changes were controlled manually and had to be fine-tuned. The first chronic lethal minimum trial at the TAL was conducted on juvenile Arapaima Arapaima gigas and found a CLmin of 16°C, making it one of the most cold sensitive freshwater species for which there are CLmin data. Chronic lethal minimum has been estimated using this system on a variety of species, including lionfish species Pterois, Dendrochirus, and Parapterois and catfish species Goonch Bagarius spp., Redtail Catfish Phractocephalus hemioliopterus, and Tiger Sorubim Pseudoplatystoma tigrinum.
The original Mastodon that was donated to the UF/IFAS TAL by the Florida Wildlife Conservation Commission.
The Mastodon was upgraded in 2020. The system now system consists of 18 58-L tanks and 12 152-L tanks. Each tank is equipped with a heater, aerator, and two temperature probes. Temperature changes are controlled electronically (using Nucleus software) and can be monitored remotely (using a Sensaphone probe). This new system allows for more fish to be ran at the same time and provides temperature data for the entire trial.
The current Mastodon that was built in 2020.
The Nucleus system that monitors and controls the temperatures.
I have conducted three trials since August 2024 and have measured the chronic lethal minimum temperature of 15 tropical freshwater ornamental fish species. I plan on estimating CLmin for a total of 50 species by June 2025. Some examples of fish I have estimated CLmin for include a variety of rainbowfish (e.g. Turquoise Rainbowfish Melanotaenia lacustris; Neon Dwarf Rainbowfish M. praecox)and cichlids (e.g. Venustus Cichlid Nimbochromis venustus; Red Zebra Maylandia estherae).
A group of rainbowfishes in one of the tanks.
Collecting the CLmin for these ornamental fishes will be used to predict the risk of invasion in Florida and throughout the conterminous USA. General warming trends have allowed species to expand their range. However, cold fronts and low temperatures continue to cause fish kills as temperatures drop below tolerance thresholds. Understanding the CLmin can predict how far a species can spread and can inform the community of when to cover their ponds.
Earth Systems Science PhD Student, Florida International University
Advisor: Dr. Jennifer Rehage
Have you ever put a lot of time into creating a nice map for your publication or presentation only for it to be spoiled by a poor quality basemap? This was the norm in my early days of making maps while working on my undergraduate habitat suitability project. Hi, my name is Cameron Atkinson, and I am a first year PhD student at Florida International University. I am working under the guidance of Dr. Jennifer Rehage and alongside many talented graduate students and lab staff in the Coastal Fisheries Research lab. My dissertation research will assess the movement and habitat use of the Common Snook (Centropomus undecimalis) across various spatial scales.
I was fortunate as an undergraduate student to have access to a GIS lab on campus where I could conduct my own research project. Along with my undergraduate research mentor, I came up with an idea to model habitat suitability for the Eastern Oyster (Crassostrea virginica) and Red Drum (Sciaenops ocellatus) throughout the coastal estuaries of Georgia. The primary way I communicated the results of my models were through maps. While the results of my models were interesting the maps themselves left something to be desired. This was due to the dark, grainy basemaps that were available through the GIS software. I was not yet aware of an alternative existed.
During my master’s work, I stumbled upon the EarthExplorer program by USGS which houses a database of high quality aerial imagery (along with many other datasets) for the entire U.S. Accessing these images are quite simple and only involves a few steps. First, draw a polygon around your study area by simply clicking on the interactive map on the web page. Next, click the “Data Sets” selection to progress to a menu of all data available within your study area. Here, select the aerial imagery tab to drop down all of the imagery options. While there are several options to choose from, I typically select the NAIP option for the best quality imagery (Figure 1). Once selecting the NAIP tab, click on “Results” to access the individual image frames. Here, you will see each individual image frame listed and dated. You can also view the footprint of single image frames to determine which files you will need to download (Figure 2). If you are downloading a single frame, use the “Downloads Option” selection. However, if you are downloading multiple frames I recommend using the “Bulk Downloads” selection. Prior to downloading you will need to create an account and/or log in.
Each file will be ~1-1.5 gigabytes each so it might take a little bit of time to download your frames, especially if you need several frames to cover your study area. The files are georeferenced so they will automatically populate to the correct location once they are imported into the mapping software you are using. When you are using multiple files to cover your study area, I recommend using a mosaic function to merge each frame together for a crisper image quality. For example, I use ArcGIS Pro and this process just requires you to select all the files, click the imagery drop-down, and select the mosaic option. From here you will have high-quality basemap imagery to build quality maps for your papers and presentations.
Figure 1. Demonstration of the selection of the NAIP imagery option with the pre-drawn study area (red polygon) on the right. Side note: Under the “Aerial Photo Single Frames” tabs are historical aerials that often span back from the 2000s to the1950s by the decade. These are often neat to see how your site has changed over time.
Figure 2. Demonstration of the individual image frame selections with the frame footprint (blue rectangle) displayed within the study area (red polygon). The green arrow points out the “Downloads Option” selection for single downloads and the orange arrow points out the “Add to Bulk Download”.
Marine Biology and Environmental Studies Undergraduate, Florida Southern College
Advisor: Dr. Gabriel Langford
My name is Brady Huber and I am currently finishing up my senior year at Florida Southern College where I am majoring in Marine Biology and Environmental Studies. For as long as I can remember, I have wanted to work with sharks in any capacity that I could. This past summer, I had the honor to work alongside Dr. Gabriel Langford and Dr. Allison Durland-Donahou on an elasmobranch spatial ecology project in the Alafia River Estuary (ARE). The goal of the project is to understand how different age classes of bull sharks (Carcharhinus leucas) and cownose rays (Rhinoptera bonasus) utilize the ARE. We do so through the deployment of acoustic receivers throughout the ARE and surrounding Hillsborough Bay, as well as the surgical implantation of acoustic tags in the bull sharks and cownose rays. Although this was an incredibly rewarding experience and a true childhood dream come true, it wouldn’t be science without some setbacks. The first of many occurred on the very first day of the summer when an angry mama bull shark ripped through our 300-foot gill net, causing so much damage that it had to be shortened to a 280-foot gill net. This was discouraging because we, as the interns, were in charge of repairing the net, but it was also exciting to see that the mamas were in the water and ready to pup. The next major setback occurred when our boat refused to start one day, and we almost became stranded in the middle of the ARE. Luckily our brave captain, Dr. Durland, was able to get the boat back up and running, but it definitely made for a shorter day in the field. The last major setback was even trying to get out into the field with the rough weather we had this summer. We have a smaller boat that is not equipped to handle high winds and rough seas and; therefore, we were limited in our time out in the field. Although they were not a “major” setback, I would like to give an honorable mention to the many catfish that became entangled in our net. The slime they excreted and the extent to which they were tangled made them quite the nuisance to remove. Even with all the trials and tribulations, I had an amazing experience on the boat, and I would not trade it for anything.
Me (Brady) with a pufferfish that got caught in our net. Credit: FSC Elasmobranch Lab
Me (Center), Dr. Durland (Left), and Mia (Right) hold tightly onto a juvenile bull shark during a work-up. Credit: FSC Elasmobranch Lab
The FSC Elasmobranch Crew of 2024. Credit: FSC Elasmobranch Lab
Have you ever counted the rings of a tree stump to estimate its age? This is the basis of my thesis research, but with fish! My name is Jordan Zulli, and I am a graduate researcher in the Fisheries and Avian Ecology Lab at Nova Southeastern University. Using fisheries age and growth methods, my goal is to define the life history of two sympatric stingray species found in the Indian River Lagoon, Florida.
The Indian River Lagoon (IRL) is an estuary ecosystem located along the east coast of Florida. The two study species – Hypanus sabinus (Atlantic stingray) and Hypanus say (bluntnose stingray) – are commonly found in this ecosystem, yet there is still much to learn about their age, growth, and reproductive patterns.
There are multiple bony structures that can be used in conjunction to age teleost fishes: fin spines, otoliths, and opercular bones, most commonly. Now, I know what you’re thinking… stingrays don’t have bones. While you are absolutely correct that elasmobranchs are cartilaginous fishes, they still possess a calcified vertebrae that presents a banding pattern on each sectioned centra, thus providing growth data. Each ring, or “annuli”, represents a year of growth, and the opaque and translucent bands occur in summer and winter, respectively (Goldman 2005).
Let’s talk more about the two study species. H. say and H. sabinus are both considered to be relatively understudied in terms of elasmobranch life history data. As you could have guessed, age/growth studies are very common in teleost fishes rather than elasmobranchs, given that they have more bony structures to be analyzed, some of which can be performed with non-lethal methods. It’s a bit trickier with elasmobranchs that only possess one hard part requiring lethal methods, which can be problematic for species with slow maturity and low fecundity. But without this information, how can we make management decisions to conserve struggling populations? If performed correctly, these studies can provide valuable data and increase our knowledge of the life history of not only the study species, but similar species that are also data deficient.
Figure 1: Measuring the disk width of a male H. sabinus before release. Photo from author.
From the lagoon…
It wouldn’t be a graduate thesis project without some field work! Thanks to a collaboration with Florida Fish and Wildlife Conservation Commission’s (FWC) Fisheries Independent Monitoring Program, inshore sampling is conducted for specimen collection. We deploy a 183-meter beach seine net and manually pull it towards the shore, drawing all fauna in the area into the bag of the net (and yes, it is as physically exhausting as it sounds). The goal is to collect two stingrays of each sex for both species every month. This ensures the consistency of the age and growth data while staying in line with IACUC protocols. Additional stingrays are tagged and released. All other nonstingray specimens are measured, recorded, and released.
Figure 2: Sampling with a 183-meter beach seine net with FWC in the IRL. Photo from author.
To the lab!
Here at the Fisheries and Avian Ecology Lab, we are known for our dissections. And that’s exactly what we do with the collected stingrays. All morphometric measurements are recorded, including total length, disk length, and disk width. Reproductive stages are then analyzed and recorded. This additional data provides further insight into the maturity and proposed reproductive cycles of each species. Finally, the moment you’ve been waiting for! The vertebral column is removed extending from the occiput to the pelvic girdle. To remain homogenous, around 10-15 of the largest centra are removed from the thoracic region. The centra are then ready to be mounted and sectioned via an Isomet-type low-speed diamond wheel saw. These sections can then be affixed to a glass slide and analyzed under a stereo microscope.
The work I do is unbelievably rewarding and knowing that the effort I put into this project has the potential to positively alter the trajectory of two ecologically important species makes the long field days and late lab nights worth it.
Figure 3: Six sections from individual centra; the “bowtie” shapes are ready to be analyzed for growth bands. Photo from author.
Figure 4: Translucent and opaque growth bands shown through stereo microscope imaging. Photo from author.
Citations:
Goldman, K. (2005). Age and growth of elasmobranch fishes. In Management techniques for elasmobranch fisheries (Musick, J. A., Bonfil, R.). Food and Agriculture Organization of the United Nations. chapter6.indd (fao.org)
Reefs to Rivers: Florida Fisheries Science 