Unlocking the mysteries of shark reproduction

By Kat Mowle, UNF MS Student

If you grew up watching Jaws, there’s a good chance you fear sharks when you go swimming in the ocean. In reality, the vast majority of sharks are completely harmless, and you are more likely to be struck by lightning than die from a fatal shark attack. If you think about it, sharks actually have more of a reason to fear us than we have to fear them, because many shark populations are in decline due to factors like overfishing, pollution, and loss of habitat. Shark and ray species are more vulnerable to overexploitation than bony fish are because sharks tend to mature at a later age, produce fewer offspring, and grow slower. Additionally, mating events for some shark and ray species may occur only every two to three years. All of these factors combined mean that sharks and rays are particularly vulnerable to overexploitation, and it is absolutely critical that we work to understand the biology and life history of these organisms so we can better manage their populations (Hoenig and Gruber, 1990; Stevens et al., 2000).

Traditionally, understanding the reproductive cycles of sharks has required lethal sampling in order to examine changes in their reproductive tracts throughout the year. This approach is unsustainable for threatened species, and cannot be used for endangered species (Hammerschlag and Sulikowski, 2011). In more recent years, scientists have moved to developing non-lethal methods for assessing reproduction of sharks and rays, which is what my advisor, Dr. Jim Gelsleichter, works on at the University of North Florida. Ultrasonography can be used to determine pregnancy, just like with humans. Scientists are also able to measure the concentrations of reproductive hormones in the blood of sharks to determine when various reproductive events occur. For male sharks, high plasma concentrations of testosterone occur during the peak time of sperm production, indicating when a male is capable of mating. For females, high concentrations of estrogen (E2) correlate with the development of eggs in their ova, indicating when a female is ready for fertilization and pregnancy (Awruch et al., 2008; Awruch, 2013; Sulikowski et al., 2007).

While these methods have proven to be useful for many species, analyzing only E2 does not always provide researchers with the full picture of a species’ reproduction. As mentioned earlier, some shark species only reproduce every two or three years. How often an individual reproduces is referred to as reproductive periodicity, with females that reproduce every year termed annual, every two years biennial, and every three years triennial. For females of species with more unusual patterns of reproductive periodicity, examining levels of E2 in their plasma doesn’t always give the full picture of their reproductive cycle. This is where my research comes in. My master’s research focuses on measuring vitellogenin (Vtg) in the plasma of elasmobranchs, specifically focusing on the bonnethead shark, Sphyrna tiburo (Figure 1).

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Figure 1. Bonnethead shark (Sphyrna tiburo) specimen

Vtg is a protein that is produced by the liver during the time of follicular development, when female sharks have new ova developing in their ovaries. Thus, determining when Vtg is produced is a good indicator of when follicular development occurs. If researchers are able to couple measurements of plasma Vtg with a nonlethal determination of pregnancy (such as ultrasonography), then a new nonlethal method for determining reproductive periodicity can be developed. For example, if a female has Vtg present in her plasma during pregnancy, that indicates the female will have eggs ready to be fertilized after she gives birth, and thus she will likely give birth again the next year. So this female would be an annual reproducer. On the other hand, if a female does not have Vtg present during pregnancy, she will likely need to take time off to grow eggs before she is ready to mate again; such a female would likely be a biennial or triennial reproducer. These measurements do depend on the length of a female’s gestation period, as the bonnethead shark is an annual reproducer, but does not produce Vtg during pregnancy. However, this species has a fairly short gestation period of only 4 to 5 months, giving females time to develop new eggs and still be able to mate and give birth every year.

My research focuses specifically on measuring Vtg in the plasma of female bonnethead sharks and characterizing the process in this species. In order to do this, I collect blood samples from female bonnethead sharks out in the field (Figure 2). I then centrifuge the blood to separate the plasma and analyze the plasma for the presence of vitellogenin using a process called Western blotting or immunoblotting. This process doesn’t give us quantitative results, but is a good method for determining whether or not Vtg is present in the plasma of a female bonnethead shark.

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Figure 2. Obtaining a blood sample from a mature female bonnethead shark.

We ultimately were able to detect both Vtg itself and a likely component protein (lipovitellin, or Lv) in the plasma of female S. tiburo. Since it is known that Vtg breaks down fairly quickly if plasma is not stored immediately or not stored with a protease inhibitor, we chose to classify females with only the Lv component protein in their plasma as also likely producing Vtg (Figure 3). It was determined that the proteins being detected were likely Vtg (~200 kD) and Lv (~70 kD) based on what had been observed for another elasmobranch species (Perez and Callard, 1992).

Figure 3

Figure 3. Immunoblots showing the detection of Vtg and Lv within the plasma of female S. tiburo. a) Detection of Vtg within plasma of females sampled in March (Lanes 2-8) and early May (lane 12). b) Detection of Lv within the plasma of mature female S. tiburo. The Lv protein is outlined in the boxes.

Looking at both the detection of Vtg and Lv as evidence of Vtg production occurring, it was determined that the highest numbers of females were producing Vtg in March and April, which matches previous studies on this species (Parsons, 1993). The protein was also produced in May for one individual. What was interesting was we found evidence of the protein’s production beginning as early as August for some individuals, with the protein continuing to be found in the plasma from August to December (Figure 4).

Figure 4

Figure 4. Proportion of mature S.tiburo females that were determined to have either vitellogenin or the putative lipovitellin protein present within their plasma during each month.

This finding suggests that female bonnethead sharks begin developing new eggs within their ovaries immediately after they give birth; production is not limited to the spring time period, which was suggested by previous studies that focused on when the highest number of large yolky eggs were observable in the ovaries. The variations we observed in when Vtg was produced are likely due to variations between populations. It has been observed, for example, that females in populations in South Florida give birth earlier than those in North Florida, which would explain why Vtg production is observed in August for females in South Florida but not until October for females captured in South Carolina (Lombardi-Carlson et al., 2003).

As I have been conducting my master’s thesis research, we have also been testing whether we can measure Vtg in the plasma of other elasmobranch species. We focused on the bonnethead because our antibody against Vtg was developed specifically for this species. But, as noted, measurement of Vtg would be particularly useful for clarifying the reproductive periodicity of other species, such as the blacknose shark (Carcharhinus acronotus), which seems to be capable of both annual and biennial reproduction in the Atlantic (Driggers et al., 2004). Measuring the concentrations of E2 in the plasma of this species does not effectively answer questions about the reproductive status of females of this species; researchers are unable to determine if a female is resting or reproductively active.

So far we have confirmed that we are able to measure Vtg (or at least component proteins of the larger Vtg protein) in the plasma of eight other elasmobranch species. These results indicate that the methods I have developed in my study will be useful for studying reproduction of other species. Particularly, these methods will be a good nonlethal method for characterizing reproductive periodicity. Having a good understanding of exactly how often a species reproduces is critical to management of a population, as the population growth depends on how many females are actually contributing to the next generation in a given year. Thus, the methods developed specifically throughout my master’s thesis will hopefully continue to be used and will act as an ideal new nonlethal method for determining reproductive periodicity, providing crucial information about a species’ reproduction to managers.

UNF Shark Biology Facebook Page

 

References

Awruch CA, SD Frusher, NW Pankhurst, and JD Stevens. 2008. Non-lethal assessment of                reproductive characteristics for management and conservation of sharks. Marine              Ecology Progress Series 355: 277-285.

Awruch CA. 2013. Reproductive endocrinology in chondrichthyans: the present and the                future. General and Comparative Endocrinology 192: 60-70.

Driggers WB, DA Oakley, G Ulrich, JK Carlson, BJ Cullum, and JM Dean. 2004.                                    Reproductive biology of Carcharhinus acronotus in the coastal waters of South                    Carolina. Journal of Fish Biology 64(6): 1540-1551.

Hammerschlag N and J Sulikowski. 2011. Killing for conservation: the need for                                alternatives to lethal sampling of apex predatory sharks. Endangered Species                      Research 14: 135-140.

Hoenig JM and SH Gruber. 1990. Life-history patterns in the elasmobranchs:                                    implications  for fisheries management. In Elasmobranchs as Living Resources:                 Advances in the Biology, Ecology, Systematics, and the Status of the Fisheries.                       Proceedings of the Second United States–Japan Workshop East–West Center,                         Honolulu, Hawaii, 9–14 December 1987, pp. 1–16. Ed. by H. L. Pratt, S. H. Gruber,                 and T. Taniuchi. NOAA Technical Report NMFS 90. 518 pp.

Lombardi-Carlson LA, E Cortés, GR Parsons, and CA Manire. 2003. Latitudinal variation              in life-history traits of bonnethead sharks, Sphyrna tiburo, (Carcharhiniformes:                   Sphyrnidae) from the eastern Gulf of Mexico. Marine and Freshwater Research,                 54(7): 875-883.

Perez LE and IP Callard. 1992. Identification of vitellogenin in the little skate (Raja                       erinacea). Comparative Biochemistry and Physiology 103B(3): 699-705.

Parsons, GR. 1993. Geographic variation in reproduction between two populations of the           bonnethead shark, Sphyrna tiburo. In The reproduction and development of sharks,             skates, rays and ratfishes (p. 25-35). Dordrecht, Springer.

Stevens JD, R Bonfil, NK Dulvy, and PA Walker. 2000. The effects of fishing on sharks,                   rays, and chimaeras (chondrichthyans), and the implications for marine                               ecosystems.  ICES Journal of Marine Science 57(3): 476-494.

Sulikowski JA, WB Driggers III, GW Ingram Jr, J Kneebone, DE Ferguson, and PC Tsang.               2007. Profiling plasma steroid hormones: a non-lethal approach for the study of                 skate reproductive biology and its potential use in conservation management.                     Environmental Biology of Fishes 80(2-3): 285-292.

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Field Surgery as a Five Foot “Tall” Woman

By Beth Bowers, FAU PhD Student

BoatSurgery

Often we enter into graduate school with a broad idea of what we might be doing. If you’re anything like me, you received a swift slap to the face when you realized that YOU were solely responsible for figuring out the logistics of your project. Logistics are always the enemy. The only certainty that grad school affords you is that Plan A will be a miserable failure time after time. Then, there are the logistics of being a small woman in the field of fisheries. Whether we like it or not, we live in a man’s world where all of the equipment was designed for those with upper body strength, so you’d better get buff if you want to run with the boys or… you could get clever.

After being accepted at Florida Atlantic University, I soon found out that my doctoral dissertation, which consists of studying the migratory pattern of the blacktip shark, also includes recruiting volunteers, managing training records, completing IACUC amendments, changing the power steering on the boat, performing a surgery while leaning off the side of a boat while Sunday Palm Beach boaters and paddle boarders take pictures…but, I’m getting off topic. Here, I will offer a solution to those eager, useful yet underestimated, a little shorter than average, women in the field of fisheries/shark research.

Many job postings will state that you must be able to lift 70 lbs. or some other amount of weight that would require ant-like strength from your tiny body. When trying to appear as equally useful as your male counterpart, it helps to utilize the pully system whenever possible. For instance, when drum lining, utilize the gunwale as leverage, as long as there are no dents or scuffs in the fiberglass that can cause the rope to fray. Don’t be too proud to let the 6’6” man beside you help you. Be useful in other ways. Find something else to do and let him use his natural-born leverage to pull in the line. When long lining, let the boat do most of the work. Don’t try to pull the boat toward the line, pull in the slack of the line as it comes to you. This advice applies to the men on the boat as well but, we’re not addressing them at the moment. Build up your grip strength! There is no better way to show your worth on a research vessel than to unclip 60 tuna clips without faltering.

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Mike McCallister, the not so hypothetical 6’6″ man beside you

Now, you’ve caught a shark that’s longer than you are tall, what do you do? Do you stand back and let the men secure it? Absolutely not. You find clever ways of navigating around your “shortcomings.” Always have someone else holding the tuna clip in case the shark gets away from you. Prepare a tail rope by making a loop out of one of the deck lines that can be quickly tightened. Understand physics. A shark has evolved to remain horizontal in the water column. In the water, pectoral and dorsal fins behave as lifting surfaces. Out of the water, these lifting surfaces are useless so keep the shark in the water. To reach the tail, the head of the shark must be lower than the caudal fin, like a see-saw. Walk the shark like a dog on a leash alongside the vessel. Ask a colleague to use a little force and to persuade the shark to swim in a circle. This will help to keep the shark swimming rather than becoming a vertical dead weight. When the shark gets close to the boat, have them shorten the length of the gangion but do NOT pull the head above the water’s surface. As your colleague shortens the length of the gangion, the shark will swim or glide alongside the vessel.

At this time, you should have your looped deck line in hand and the loop should be big enough to fit around the entire height of the heterocercal caudal fin. As the shark’s tail end approaches, reach into the water from the stern of the boat, grab the caudal fin, and slip the deck line over the tail. Your colleague should keep the gangion taut to prevent the shark from turning around and biting you. Once the line is tight around the caudal peduncle, tie off the tail rope to the stern cleat of the boat, leaving enough slack for the shark to remain halfway under the water’s surface. This not only frees your hands from having to hold the shark, it keeps your crew at the stern of the boat, the only place where you can reach the water. You should be able to release the tail rope quickly in case the hook rips out of the mouth or a crimp on the gangion fails. This CAN and WILL happen. If you have ever had a timid undergraduate help you with repairing broken gangions, you already know this. Use an additional deck line as a mid-body rope. Most people can lean off the side and reach around the body of the shark to feed the tightening end of the deck line into the loop; I cannot…unless I feel like going for a dip in the ocean. Instead, I make a large loop in the deck line and feed it over the free end of the gangion. I then slip the loop over the head and pectoral fins of the shark, position the line anterior to the dorsal fin, and posterior to the pectoral fins.

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Securing a tail rope on a blacktip shark

Once you have the shark secured, prepare your surgery equipment. Thread the monofilament suture into the end of a reverse cutting suture needle. In my experience, a tapered edge suture needle will dull halfway through the procedure. Fill a syringe with lidocaine and inject it into the musculature on the anterior and posterior sides of the planned incision site. This is a good time to let the crew take measurements, a DNA fin clip, tissue samples, and insert an external dart tag. Don’t be a hero. Let them do it. Your back will take enough of a beating during the surgery. When you are ready for surgery, assign someone to play nurse and someone to help hold up the shark. You should also ask someone to be on watch for wakes. You’ll understand why I say that when you almost flip into the shark’s mouth the first time a wake passes. The nurse should be ready with the hemostats and suture needle. The shark holder should spare his/her back as much as possible by adjusting the mid rope and standing on the slack of the rope that is in the boat. You are again employing the pully system; good for you. This allows the holder to stand upright for most of the procedure and prevents him/her from grunting and whining in your face while you hang off the side of the boat and play doctor. Let your “nurse” pass you the next instrument and take the used one out of your hand. It helps if you teach your crew to pass you the instruments in the orientation in which you will use them, just as a nurse does to a non-shark surgeon.

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Boatside inversion

Lean over the side of the boat…hope you’re not feeling nauseous because you are basically inverted. Get ready for a hamstring workout. Make an incision midway between the pelvic fins and pectoral fins along the ventral midline. If you make the incision just large enough to fit the transmitter, then you only need one suture to close the incision. Your hamstrings will thank you. Use the blunt end of the scalpel handle to ensure that you have penetrated the abdominal cavity; through the skin, muscle, and peritoneum. Double check that the transmitter ID number has been recorded. You WILL forget to do this. If anyone who works with acoustic tags tells you they have never forgotten to record the ID code, he/she is a filthy liar and is never to be trusted again. You can forget once per day and figure out the code by process of elimination back at the lab, but not twice. Insert the transmitter into the incision.

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Hemostats holding a suture needle

Use the hemostats to insert the suture needle into the incision and back out through the side closest to you. Do so by moving your wrist in a scooping motion towards your body. Don’t poke yourself in the finger with the dirty suture needle as it exits the incision margin. When pulling the suture needle through, be careful not to allow the suture thread to slip out of the incision – starting over at this phase is not ideal. You can ensure that it will not slip out by clipping a pair of hemostats onto the free end of the suture monofilament before pulling the suture needle through. Close the incision with a surgeon’s knot. View a helpful video here: https://www.youtube.com/watch?v=Av2gp-3mKwE. At this point, your hamstrings feel like they’re going to rip through your skin. Flip the shark into the upright position, dorsal side up. Remove the hook from the mouth. Synchronize the loosening of the mid rope and the tail rope. Hold the dorsal fin until the shark awakes. Release the shark by allowing it to swim through the large loops you have created in the deck lines. Make sure the loops stay large until the caudal fin is free from entanglement with each deck line. God help you if you have the shark by the tail only; I have no advice for that situation.

Congratulations, you have just tagged a shark that is bigger than you are. Now stretch it out and suck it up because it’s peak fishing season and you’ll have nine more to do today!

Facebook: FAU Shark Migration          Instagram, Twitter: @sharkmigration

 

Smile for the Camera Fishes!

By Carissa Gervasi, FIU PhD Student

Gervasi_Happy Bonnethead

A happy Bonnethead posing for the camera!

When I began my PhD at Florida International University in August 2016, my advisor immediately threw a newly funded 3-year project at me to take the lead on. My task? Figure out a way to assess fish species diversity and abundance in a protected area using GoPro cameras. My first thought was wait, we are using GoPros to conduct scientific research?? Seemed a little sketchy to me…

After poring through the literature, I realized that underwater video survey methods are becoming more and more common, as they are minimally invasive, and don’t harm marine life or the environment. They also have the added bonus of catching the behavior of fish as they are swimming through their natural environment. Pretty cool!

Baited Remote Underwater Video Systems (BRUVS for short) are often used in coral reef environments to determine species richness and abundance without having to send divers down to count fish, an expensive and time consuming endeavor. I was tasked with taking this new method to the shallow, coastal waters of Florida Bay to assess how the closure of the Crocodile Sanctuary has impacted fish abundance and diversity. This area has been closed to the public since 1980!! That’s a long time without humans, and we were bound to see some changes to the fish communities.

One thing I never expected I would have to do for my PhD was become an engineer. I was tasked with constructing several BRUV systems that were small and lightweight, so we could take them on our tiny boats out into the super shallow waters of the bay. I was also working with a set budget, and had to be inventive about the materials used. My lab mates and I drafted a design, and after much trial and error, created our BRUVS!

BRUV-frame-

The anatomy of BRUVS

We deploy our BRUVS twice a year inside and outside the protected area using a stratified random survey design. They are quick and easy to deploy; we simply fill the bait bag, turn the cameras on, drop the frame in the water and leave! We let the cameras sit on the bottom and record data for 90 minutes each deployment.

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Me deploying a BRUV frame into the water

This project has been so unique and a lot of fun! We have a small army of undergraduate interns working in our lab to watch the thousands of hours of video data we have collected, and we have seen some really cool stuff! We have recently completed our second year of sampling, and are starting to analyze the data. So far we have seen that there are a lot more sharks in the protected area than outside, which is a very interesting result! There are many important fish species that use coastal areas within Florida Bay as nursery habitat, including recreationally targeted species like snook and tarpon. Our research will hopefully help guide management actions within Everglades National Park, and help conserve the species that call this special bay home.

Check out our Best of BRUVS YouTube video to see some of the coolest fish we have seen so far on our videos! https://www.youtube.com/watch?v=8ZYKXk-HWOs&t=2s

Also, check out our lab on social media!

Facebook: Rehage’s Coastal Fisheries Lab

Instagram: @fiu_fisheries_lab

All about that bait: Pinfish population dynamics in the eastern Gulf of Mexico

By Meaghan Faletti, USF MS Student

If you’ve ever gone fishing in the Gulf of Mexico – or anywhere around Florida, for that matter – you’ve probably used Pinfish (Lagodon rhomboides) as bait. They work well because many predators consume them as part of their natural diet. This includes nearshore species such as Tarpon, Snook, and Redfish, as well as offshore species such as groupers and snappers. In addition to being used in the recreational fishery, a commercial market exists for Pinfish and extracted over 100,000 pounds of Pinfish from Florida waters in 2016 (NMFS 2018). Furthermore, recreational and commercial landings in the Atlantic and Gulf coasts of the United States show a general increasing trend since 2000 (NMFS 2014). Despite their importance to fisheries, a formal stock assessment has not been conducted for Pinfish.

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Figure 1. Pinfish (Lagodon rhomboides)

With Florida’s ever-growing fishing industries, efforts to protect these important food sources are needed to sustain higher trophic-level species that are targeted by the recreational and commercial fisheries, and maintain a natural ecological balance. Not only are Pinfish providing food for predatory fishes, but they also act as a major source of nutrient transfer from nearshore to offshore foodwebs when they migrate to spawn. The nitrogen contribution from Pinfish migration offshore is on the same order of magnitude as trichodesmium, a major nitrogen-fixing bacteria, and one of the most significant contributors of nitrogen to marine systems (Nelson et al. 2013).  Due to the importance of Pinfish and other forage (“bait”) fishes, the Florida Fish and Wildlife Conservation Commission (FWC) adopted a resolution in 2017 encouraging further research on these species. The Florida Forage Fish Coalition was then formed to fund this research, and the USF Fish Ecology Lab was chosen as one of the first recipients.

We chose to study the population dynamics of Pinfish in the eastern Gulf of Mexico (eGOM) due to our lab’s familiarity and previous work with this system (Chacin et al. 2016; Stallings et al. 2015) as well as the extensive monitoring previously conducted by FWC’s Fisheries Independent Monitoring program. We chose to focus on four eGOM estuaries that have been sampled monthly with standardized seining methods since 1998: Apalachicola Bay (AP), Cedar Key (CK), Tampa Bay (TB), and Charlotte Harbor (CH). We analyzed density, biomass, and instantaneous growth rates (basically – how many, how big, and how fast are they growing?), and conducted time series analyses to see if these parameters were in synchrony with one another on an intra- and inter-annual basis. We then conducted a Zero-Altered Negative Binomial (ZANB) Model – don’t worry about the name, all you need to know is that it is used to assess what environmental factors may be driving these population patterns.

Pinfish estuaries

Figure 2. Four eGOM estuaries of interest where FWC Fisheries Independent Monitoring has conducted seine surveys since 1998. AP: Apalachicola Bay, CK: Cedar Key, TB: Tampa Bay, CH: Charlotte Harbor

We found density and biomass to be highest in CH, followed by TB, AP, then CK. Our time series analysis indicated that AP & CK (two northernmost estuaries) are in sync with one another, and TB & CH (two southernmost estuaries) are in sync with one another. On an intra-annual basis, all four estuaries are in sync with one another, with densities peaking around March-April, and biomass peaking in April-May. This makes sense from a life history standpoint, as Pinfish recruit to seagrass in the spring, and after feeding for a period of time, we expect to see an increase in mass lagging slightly behind the recruitment period.

The ZANB model indicated that density and biomass are significantly related to submerged aquatic vegetation (SAV; e.g. seagrass) for all estuaries. This is likely because seagrass plays an important role for Pinfish by offering protection for juveniles (Chacin & Stallings 2016) and substrata for food as they grow older and begin foraging on epiphytic algae. Pinfish abundance is related to salinity and temperature, which also affect seagrass, so this could either be through direct or indirect effects.

interannual density

Figure 3. Mean inter-annual Pinfish A) density and B) biomass for 1998-2016. Light gray dotted line represents the overall mean.

annual density

Figure 4. Intra-annual Pinfish A) density and B) biomass for 1998-2016. Biomass peak lagged ~1month behind density peak.

GrowthRate

Figure 5. Pinfish growth plots. A) Instantaneous Growth Rate (IGR) for each estuary. B) Mean Pinfish standard length (SL) by month. Month 0 represents the average size of the smaller cohort present in month 12 of the previous year.

It is important to understand baseline population dynamics of fisheries species, including forage fish, so that we can better understand the effects of commercial and recreational fishing. This is especially true for species that do not have formal stock assessments such as Pinfish. This information has been shared at the Florida Chapter Meeting of the American Fisheries Society (Haines City, March 2018), the 2nd Florida Forage Fish Workshop (St. Petersburg, April 2018), and will be presented to the FWC Commissioners during a 2018 Commission Meeting. The project has also attracted a wealth of media coverage due to the importance of Pinfish and concerns from stakeholders about potential overharvest. We plan to publish a peer-reviewed article on these data in the summer of 2018 and intend for this information to be used to help better manage Pinfish populations.

 

References:

Chacin, D. H., T. S. Switzer, C. H. Ainsworth, and C. D. Stallings. 2016. Long-term analysis of spatio-temporal patterns in population dynamics and demography of juvenile Pinfish (Lagodon rhomboides). Estuarine Coastal and Shelf Science 183:52-61.

Nelson, J. A., C. D. Stallings, W. M. Landing, and J. Chanton. 2013. Biomass Transfer Subsidizes Nitrogen to Offshore Food Webs. Ecosystems 16:1130-1138.

NMFS 2018 Commercial landings species locator. https://www.st.nmfs.noaa.gov/pls/webpls/FT_HELP.SPECIES

NOAA NMFS Marine Recreational Information Program, http://www.st.nmfs.noaa.gov/; National Marine Fisheries Service, accessed 4 March 2014

Stallings, C. D., A. Mickle, J. A. Nelson, M. G. McManus, and C. C. Koenig. 2015. Faunal communities and habitat characteristics of the Big Bend seagrass meadows, 2009–2010. Ecology 96:304-304.

 

Acknowledgments:

Florida Forage Fish Research Program funded by the Florida Forage Fish Coalition (www.floridaforagefish.org). We thank the Florida Fish and Wildlife Research Institute Fisheries Independent Monitoring Program for field collections, and Ian Williams, Aleksandra Cison, and Kiara Barbarette for lab assistance. We also thank Ethan Goddard for assistance with the mass spectrometer.

 

Funding Sources:

Funding

“Don’t go into the light!” – Using Light Traps to Sample Ichthyoplankton

By Amanda Croteau, UF PhD Student

The Original Light Trap

Meroplankton are animals that spend a portion of their lives (larval and early life stages) as plankton. These organisms eventually grow larger and become part of the nekton (animals that are able to swim and move independently of water currents) or benthic communities. Ichthyoplankton are the eggs and larvae of fish. Eggs are passive and dispersed by currents. Initially most larval fish have no or minimal swimming ability. As they develop, they become active swimmers.

It is important to study meroplankton and ichthyoplankton because they are indicators of the spawning population of adults, and the survival or mortality of meroplankton have a direct effect on adult population numbers. Species composition at a given location depends on the spatial distribution and reproductive habits (periodicity, fecundity, etc.) of adults, growth and larval stage duration, and abiotic factors that affect transport (currents, tides, salinity, etc.). Mortality depends on many factors such as predation, disease, food availability, and habitat. Habitat is important because individuals who fail to make it to their correct settlement or juvenile habitat are unlikely to survive. In estuarine environments, freshwater and tidal cycles play key roles in species distribution.

Mangroves and salt marshes provide vital juvenile habitat for many inshore, nearshore, and offshore marine species. Florida’s coastal habitats have been severely impacted by coastal development, and Tampa Bay has lost over 44% of its mangrove and salt marsh habitats (Lewis et al. 1985). Robinson Preserve is one of the largest (197 hectare) mangrove and salt marsh restoration efforts in Tampa Bay. Robinson Preserve was originally a coastal wetland that was ditched and drained in the 1920s for agricultural use. In 2006, tidal flow was restored through connections with Perico Bayou, Palma Sola Bay, and the Manatee River. Restoration also involved the planting of native upland and salt marsh vegetation. However, no efforts were made to supplement the aquatic flora and fauna, rather it was expected that they would colonize the preserve from neighboring populations. Ichthyoplankton and meroplankton abundances were selected as one metric to evaluate the quality of the restored ecosystem as nursery habitat.

Meroplankton and ichthyoplankton can be sampled in a variety of ways including light traps, benthic sleds, Miller high-speed samplers, push nets, tow nets, and light traps. As with any sampling gear, each method has its pros and cons and gear selection should be informed by target taxa, gear bias, and site constraints. Light traps utilize organisms’ natural attraction to light (photopositive) as bait. Photopositive taxa approach and enter the trap and are then funneled into a collection chamber. Light traps also allow you to sample continuously over an entire night at multiple locations.  Robinson Preserve is shallow (generally <2 m), with complex habitat types and obstructions. It is also a no motor zone. Due to these study site constraints, light traps were selected as the most efficient gear to sample ichthyoplankton and meroplankton within the preserve. The light trap designed by Jones (2006) was redesigned for deployment from shore and scaled down for use in shallow, estuarine systems (Figure 1 and 2).

1

Figure 1. Design of modified light trap. The trap is powered by a battery located on-shore. Fish are attracted by the light source in the entrance chamber. The size of the openings, restrict the size of the organisms that can enter the chamber. Organisms are then funneled into the collection chamber where a mesh screen allows water to exit, but prevents organisms from escaping. Floats keep the trap vertical in the water column, and it is anchored in place. In shallow tidal systems, the depth changes due to incoming and outgoing tides must be considered when placing the trap.

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Figure 2. Light trap deployed along mangrove shoreline in Robinson Preserve.

Among the larger organisms (≥3 mm) collected, 18 major taxonomic groups have been identified to date. Overall community composition was dominated by isopods (19%), caridean shrimp (18.2%), fish (15.7%), and parasitic copepods (13.1%), though species assemblages varied by site and season (Figure 3). The greater taxonomic richness in sites 1 and 3 is likely related to their locations. Both of these sites were located in areas with slower currents than in sites 2 and 4, which may have allowed some less mobile species to enter the light traps than in the latter two sites. Larval and settlement stage fish were collected in nearly every sample (Figure 4), including fish from at least 8 families. This is similar to the degree of diversity noted in other light trap studies in similar habitats (Hernandez and Shaw 2003; Strydom 2003). Juvenile mullet (likely Striped Mullet Mugil cephalus) were always present in winter samples, while juvenile clupeids (likely menhaden Brevoortia spp.) were present in the winter and spring, which corresponds well with their respective peak spawning periods within this region.

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Figure 3. Ichthyoplankton and meroplankton community composition for each site (1.DK in Mixing Zone, 2.B1 in Palma Sola Bay and Perico Bayou zone, 3.W in Upland Freshwater Drainage zone, and 4.PD in the Manatee River zone) by season.

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Figure 4. Ichthyoplankton community composition for each site (1.DK in Mixing Zone, 2.B1 in Palma Sola Bay and Perico Bayou zone, 3.W in Upland Freshwater Drainage zone, and 4.PD in the Manatee River zone) by season.

The use of a modified light trap in Robinson Preserve proved to be an effective method for sampling ichthyoplankton and meroplankton, as well as some other groups. Several fish parasites were collected in large numbers during this study. Juveniles of the parasitic isopod family Cymothidae were by far the most dominant form of isopod present in the light trap samples. Parasitic copepods of the genera Argulus and Caligus were also collected in large numbers. The high abundance of external fish parasites collected with this method may provide a new and efficient means of surveying such taxa in estuarine systems.

References:

Hernandez, F.J., and R. F. Shaw. 2003. Comparison of plankton net and light trap methodologies for sampling larval and juvenile fishes at offshore petroleum platforms and a coastal jetty off Louisiana. American Fisheries Society Symposium 36: 15-38.

Jones, D.L. 2006. Design, construction, and use of a new light trap for sampling larval coral reef fishes. NOAA Technical Memorandum NMFS-SEFSC-544.

Lewis, R. R., R. G. Gilmore, Jr., D. W. Crewz, and W. E. Odum. 1985. Mangrove habitat and fishery resources of Florida. Pages 281-336 in W. Seaman, Jr., editor. Florida aquatic habitat and fishery resources. Florida Chapter, American Fisheries Society, Kissimmee, Florida.

Strydom, N.A. 2003. An assessment of habitat use by larval fishes in a warm temperate estuarine creek using light traps. Estuaries 26(5): 1310-1318.

How to catch a Koi: A failed extraction adventure

By Allison Durland Donahue

UF PhD Student

What do you do when you receive a request from a concerned citizen asking for assistance rescuing her Koi from her neighbor’s pool? You excitedly accept the challenge and begin planning the best way to catch a Koi. You are the expert. You will know how to catch the darn fish when others have not been able to. Enter real life.

Last month, UF’s Aquatic Research Graduate Organization (ARGO) received a request to remove a Koi from a pool. This seemed like an excellent outreach adventure – catching a Koi and teaching citizens about fishing techniques, exotic species, and other fish things.

First question: How did the Koi get in the pool in the first place? Irma. During Irma, this area had major flooding (a creek became a lake). The Koi was moved over a mile with the flood waters. The owner assumed her prized Koi was lost, but her neighbor called with news that there was a Koi in his pool.

Next question: How many avid fisher people does it take to catch a Koi? More than three (plus a shellfish person). The four of us attempted to use our advanced degree trained minds and fishing expertise to design a plan to get the elusive Koi.

The challenge: An oddly shaped, 12-foot deep pool, a Koi that hangs out on the bottom, and three feet of water visibility.

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Analyzing the challenge

Extraction method, take one: Deploy a seine with the hopes that the Koi will startle into the net. The seine was stretched across the width of the pool and drug through the water column, catching nothing but leaves.

Extraction method, take two: Use the seine on the surface and an ingeniously crafted seine on the bottom to cover the entire water column. The owners had built their own seine out of bamboo poles and chicken wire. Combined, the two seines could cover the entire water column. Or so we thought.

Extraction method, take three: Tie a weight to the seine to (hopefully) ensure that the net is reaching and staying on the bottom. The trick is to make sure the net is not ever dragging the weight. One person stood on the opposite end of the pool and pulled the weight on the bottom of the pool as the other two pulled the seine. Alas, no fish.

Conclusion: Either that Koi is the smartest Koi ever or it was removed from the pool via natural methods (i.e. an eagle ate it).

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The only fish we caught

Even though the extraction failed, we were able to educate the owner about fish health requirements and various fishing methods. We left her with our last method idea: build a seine that is fourteen feet high with limited slack. And with that we hung our fishers’ heads in shame and swore at that elusive Koi.

President

Natalie Simon

Natalie

Natalie Simon is from New Jersey and received her BS in Marine Sciences from Stockton University. While working at Rutgers’s Haskin Shellfish Research Laboratory as a hatchery technician, she found her love for oysters. Not long after, Natalie moved to Gainesville to attend the University of Florida (UF) for a Master’s degree in Fisheries and Aquatic Sciences and has since stayed to continue her academic career for a PhD. Her research interests include cryogenics, germplasm preservation, and molluscan aquaculture.

 

Vice President

Allison Durland Donahou

Allison
Allison Durland Donahou is from Seattle, but ran away to warmer, sunnier weather ten years ago and has never looked back. She received her BA from the University of San Diego in Marine Biology and her MS from Nova Southeastern University in Marine Biology and Coastal Zone Management. While working with Alaskan fishing communities as a research assistant with NMFS, she discovered her interest in the human dimensions of fisheries. For her PhD, Allison is trying to tackle the challenge of managing invasive species, specifically examining the effects climate change will have on non-native fish distributions. When she finds “free” time, Allison loves partaking in water sports with her puppies and husband, as well as exploring what Gainesville restaurants have to offer.

Secretary

Beth Bowers

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Beth Bowers has been a member of the American Fisheries Society (AFS) and the Florida chapter since 2017. In addition to her Secretary/Treasurer appointment at the AFS Florida chapter, Beth serves as the Vice President of the College of Science Graduate Association at her university. She also serves on the Student Affairs Committee for the American Elasmobranch Society and is the Lab Supervisor of the Elasmobranch Research Laboratory at Florida Atlantic University. Beth is a doctoral candidate at FAU, where she studies the migratory behavior of the blacktip shark in the western Atlantic. She utilizes acoustic telemetry and mark recapture in her thesis work. Additionally, she contributes to a few collaborative projects, which involve accelerometry and characterizing the immunological repertoire of blacktip sharks.

University Liason

Lauren Kircher

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Lauren is from western New York and received her BS in Marine Biology from University of New Haven. Lauren participated in several fellowships at University of New Haven and University of Southern California, nurturing her love of research. Following her BS, Lauren started a Ph.D. in Integrative Biology at Florida Atlantic University. Her dissertation focuses on natural and anthropogenic environmental influences on the movement of a tropical sportfish (common snook) in St. Lucie estuary. Lauren’s research interests include fisheries, movement ecology, behavioral ecology, and physiology.