Seeing Red: Can hurricanes tell us how to bring back Red Drum (Sciaenops ocellatus) to Florida Bay?

By: Jonathan Rodemann, FIU PhD candidate

I was a bass fisherman as a kid. Fishing the lakes up in New Jersey was all I knew. Then I moved to Florida to attend University of Miami as an undergraduate student and was introduced to the wonders of coastal saltwater fishing. I heard stories about snook, tarpon, jacks, snapper, and other incredible sport species. However, one species caught my attention and has been an obsession ever since. Red drum (Sciaenops ocellatus), also known as redfish or just reds, is a coastal fish species ranging from the Gulf of Mexico all the way up to New Jersey (who knew I had them in my neck of the woods?!), but are most common in the southern portion of their range. In Florida, they are one of the most sought after sportfish, known for their “bullish” fights and delicious taste.

These traits ramped up interest in red drum fishing in Florida Bay through the mid-20th century. Fishing was good back then, with healthy populations and no regulations. By the early 1980’s, 40% of the recreational fishermen in Florida Bay were targeting red drum and there was a strong commercial harvest as well. It felt like the unlimited supply of reds would never end.

Then catches began to drop off. In a fashion that is too common nowadays, overfishing led to a large decline of red drum in Florida Bay. It got so bad that emergency seasonal closures of the fishery, as well as size and bag limits, were enacted in the late 1980’s. Redfish populations started to recover due to these regulations all around Florida, except for Florida Bay. The altered freshwater flow input into Florida Bay from the canalization of the Everglades has changed the salinity regime, reducing habitat quality throughout Florida Bay and causing widespread seagrass die-offs. With all these detrimental alterations to the habitat, the already low populations of red drum could not recover.

Coming to FIU, I knew I wanted to study the ecology of recreationally important fish species. So I jumped at the opportunity when a project opened up in the coastal fisheries lab run by Dr. Jennifer Rehage looking at how the 2015 seagrass die-off in Florida Bay has affected the movement and trophic ecology of grey snapper (Lutjanus griseus), spotted seatrout (Cynoscion nebulosus), and red drum. However, when describing my project to people who have fished Florida Bay, all I heard was “Good luck! You won’t find any redfish there!”. Worried about the project, I started to look at other species as options. But then I started to hear whisperings. Trickling down through the grapevine was information on an abundance of “puppy drum,” red drums under the size limit, in Florida Bay. Excitement built and then curiosity set in. What caused this huge recruitment pulse? I came to the only logical conclusion: Hurricane Irma.

Hurricane Irma passed directly over Florida Bay in early September of 2017, the first hurricane to do so since the mid 1900’s. While devastating for human populations in South Florida, Irma was a godsend for many fish species, including red drum. The category 4 hurricane dumped a tremendous amount of freshwater into the entire Everglades system, drastically reducing the normally high salinities in Florida Bay. This input of freshwater along with the timing (breeding season for redfish is August to December) created a perfect storm (pun intended!) for red drum recruitment.

Now in early 2019, we are seeing tons of 12-15 inch redfish, whose size is indicative of hatching right after Hurricane Irma. So what does this teach us about the red drum population of Florida Bay? It seems that high freshwater input will increase the number of spawning events and the survival rates of the offspring. Therefore, restoring freshwater flow into Florida Bay through programs such as CERP (Comprehensive Everglades Restoration Plan) has the potential to bring back red drum populations! I am very excited to start tracking these “Irma reds” to learn more about what makes them tick, hopefully leading to information that will help redfish thrive once again in Florida Bay.

The Cost of Fishery Collapse

By: Nick Fisch, PhD candidate, University of Florida

I’d like to tell a story about one of the most infamous fishery collapses in the modern era. The year was 1992, and the single largest unemployment event in Canada’s history was impending. Fisheries and Oceans Canada had just determined that the Atlantic Cod stocks off the coast of Newfoundland and Labrador could no longer sustain fishing pressure. Over five hundred years of exploitation had rendered the population to less than 10% of its historical levels. What followed is now history. A moratorium was imposed on the fishery along Canada’s east coast. Approximately 35,000 people from Newfoundland and Labrador were put out of work overnight, and hundreds of thousands more had their livelihoods negatively affected. Fishermen docked their boats, fish plants closed, local communities were devastated. What is often overlooked is that when a fishery collapses, not only do the people directly involved in the fishery suffer, but the entire community suffers. The money that was once made exporting fish, and subsequently used in downtown shops, grocers, and stores, is no longer available. In this particular case, the coastal community was an entire province. At first, the moratorium was to only last two years. As of today, twenty-six years later, the fishery has yet to fully re-open, and Newfoundland and Labrador has yet to fully recover.

Cod drying in Newfoundland (https://www.youtube.com/watch?v=K-nLURkGsac)

The reason the moratorium on Northern Cod was imposed suddenly, as opposed to a recovery plan involving more strict harvest quotas aimed at rebuilding the stock over time, was partly attributed to errors in stock assessment (Walters and Maguire, 1996). The assessment assumed fisher catch rates were proportional to stock abundance when in reality, the catch efficiency of fishers had been increasing over time with improvements in fishing gear. This caused declines in the stock that were “masked” by stable or increasing catch rates of fishers over time. By the time assessment scientists realized this, the stock had been depleted to less than 10% of its historical level and managers deemed an immediate moratorium necessary.

The interesting aspect to this case is that these assessment methods, whereby fisher catch-per-unit-effort (a metric of catch rate) is treated as a proportional index of abundance, were state-of-the-art at the time, and to some degree still are. To quote C. Walters and J. Maguire in their publication Lessons for stock assessment from the northern cod collapse; “The errors resulted from using assessment methods and population dynamics assumptions that were widely considered to be sound, and that still remain unquestioned by many fisheries scientists.” (Walters and Maguire, 1996). This remains an important reminder that we must constantly question our assumptions in any biological models from which management decisions may be based. In part, my research aims to do that by evaluating how assumptions we make regarding data collection affect results from stock assessments and ultimately fishery sustainability. I am currently in my first year as a PhD student in the University of Florida’s Program for Fisheries and Aquatic Sciences, supervised by Drs. Rob Ahrens and Ed Camp. I obtained my BS from UF in 2015 in Wildlife Ecology and Conservation with a minor in Statistics. I then went to Michigan State University for my MS in Fisheries and Wildlife. I finally made it back to Florida!

You might ask yourself, how does this story relate to Florida? Well, as many reading this blog may know, Florida contains some of the most valuable fishing industries in the country. Fishing industries that depend on sustainably managed fisheries, partly achieved using information from stock assessments. To keep this up (to keep Florida the “Fishing Capital of the World”), we must constantly evaluate our models to make sure they are accurate representations of reality.

Evaluating Invasive Risk of Alligator Gar (Atractosteus spatula) in Aquaculture in Florida

By: Lauren Lapham M.S Candidate, University of Florida

Figure 1. Atractosteus spatula (Source: FWC)

Alligator Gar, Atractosteus spatula, is imperiled across much of its native habitat; occurring in just a small portion of the Florida panhandle (Figure 2; Buckmeier 2008). In 2006, the Florida Fish and Wildlife Conservation Commission implemented a harvest closure on Alligator Gar, making it illegal to take or possess Alligator Gar without a permit (FWC). Research efforts are ongoing to determine Florida Alligator Gar abundance, population dynamics, movement, and habitat use (Wegener 2018). There is interest in Florida to culture Alligator Gar for food and out-of-state sale as ornamentals. Although Alligator Gar is imperiled and native to Florida, there is concern regarding aquaculture of the species due to potential deleterious genetic impacts resulting from interactions between native and cultured gar and the potential of cultured Alligator Gar establishing in peninsular Florida.

Figure 2. Map of Alligator Gar native range in the United States (Source: easternbrooktrout.org)

Before making decisions regarding commercial aquaculture, it is prudent to evaluate the risk of establishment and impact. Invasiveness risk will be evaluated through an extensive literature review and biological synopsis, risk screens, and a stakeholder-inclusive qualitative risk assessment. The biological synopsis is an extensive literature review of species biology, range, and factors related to potential invasiveness. Multiple independent assessors will use the Fish Invasiveness Screening Kit (FISK) v2 to provide a preliminary risk estimate of the species (Lawson et al. 2013, 2015). The stakeholder panel will consist of experts in state and federal agencies, academia, or other organizations and will further evaluate invasiveness risk using a qualitative risk assessment methodology (ANSTF 1996). Research and management recommendations arising from the risk-based process will be available to agencies and industry to support decision-making regarding the need for additional research or risk assessment, allowance of culture under basic Florida Aquaculture Best Management Practices (BMPs) or under specific risk-mitigating conditions, or a prohibition of aquaculture of Alligator Gar.

For any more information about this project, please contact Lauren at l.lapham@ufl.edu

C:\Users\llaph\Documents\AG FWC large.jpg
Figure 3. Alligator Gar from FWC sampling in Escambia River (Source: FWC)

References:

Aquatic Nuisance Species Task Force (ANSTF). 1996. Generic Analysis

Buckmeier, David. 2008. Life History and Status of Alligator Gar Atractosteus spatula, with Recommendations for Management. Heart of the Hills Fisheries Science Center; July 31, 2008. https://tpwd.texas.gov/publications/nonpwdpubs/media/gar_status_073108.pdf

FWC: https://myfwc.com/research/freshwater/sport-fishes/alligator-gar/

Lawson, L. L., J. E. Hill, S. Hardin, L. Vilizzi, and G. H. Copp. 2015. Evaluation of the Fish Invasiveness Screening Kit ( FISK v2 ) for peninsular Florida 6(4):413–422.

Lawson, L. L., J. E. Hill, L. Vilizzi, S. Hardin, and G. H. Copp. 2013. Revisions of the Fish Invasiveness Screening Kit ( FISK ) for its Application in Warmer Climatic Zones , with Particular Reference to Peninsular Florida 33(8).

USFWS: https://easternbrooktrout.org/groups/whitewater-to-bluewater/species-spotlight/alligator-gar/watershed-distribution-status-of-alligator-gar/image_view_fullscreen

Wegener, M. 2018. Alligator Gar Research in Pensacola Bay. Annual Project Report.

Fish Eyes: They’re not just for grossing out your siblings anymore!

Fish eye & dissection photos by Julie Branaman

By: Julie Vecchio, PhD candidate, College of Marine Science, University of South Florida

Random person on the street: Oh, you’re a marine biologist? What do you study? Sharks? Corals? Whales?

Me: Nope. Fish eyes.

RPOTS: ……oh……

Me: Actually, it’s quite interesting. The chemistry inside the eye-lenses records information about where the fish has gone and what it’s been doing throughout its whole life!

RPOTS: WOW! Really?

Me: YUP! We collect the fish using longlines or trawls or fishing poles. Then we bring them back to the lab, remove their eye-lenses, and…

Fish eye & dissection photos by Julie Branaman
Fish eye & dissection photos by Julie Branaman

RPOTS:

Image result for sick emoji

Me: Hang on! It’s actually super cool! See, throughout the life of the fish, and your life too, the eye-lens is constantly laying down new cells. But, all the DNA and mitochondria, and other organelles get in the way of the animal being able to see. So, they take them out, and just leave clear protein. That protein is made of the Carbon and Nitrogen that are available in the diet at the time that those cells formed. Once the cells have taken out all of the other “stuff,” it leaves the chemical signature of where they were at the time that lens layer was formed.

RPOTS: OH! So the chemistry in the eye-lens helps you discern where they were at any time during their life?

Me: Yes! But in order to know that, I have to peel the lenses apart. They are formed like an onion. The oldest proteins are on the inside (from when the fish was the youngest), and newer ones (as the fish grows) are deposited all around the older ones. The outer layer is the newest material, laid down within the past couple of weeks.

Fish eye & dissection photos by Julie Branaman
Fish eye & dissection photos by Julie Branaman

RPOTS: Wow! That must be a lot of very delicate work to peel those lenses.

Me: Yup. It sure is. The lenses start out about 4-5 cm in diameter, but as I peel, they get smaller and smaller. The smallest amount of protein we can process is about 0.5 mm in diameter, a tiny dot of tissue from when the fish was only about 1-2 months old. We are even able to get this information from fish that we catch at 5 or 10 years old.

RPTOS: So, you just know the chemistry after you peel the lenses?

Me: Well, not exactly. I then have to load each of those tiny samples into a machine called an Elemental Analyzer-Isotope Ratio Mass Spectrometer.

RPOTS: *glazes over*

Me: It burns up each one of those tiny eye-lens samples at 1000° and turns it into gas. It then counts the atoms in the gas and reports the amounts of heavy or light versions of the carbon and nitrogen isotopes.

https://www.marine.usf.edu/images/thermo-delta-xl-irms-continuous%20flow/IMG_9410.JPG

RPOTS: 1000°????

Me: Yup. Glad I’m not in there!

RPOTS: So, then you know EXACTLY where the fish were when they were tiny?

Me: Well, not EXACTLY where they were… But we have an idea of if they were offshore or inshore. If they were eating plankton or shrimp in the bottom. This is a new field, and we are still working on how to interpret a lot of the results that we get, but it’s cool anyway. This field has a lot of promise, and it’s just beginning. We are building the field as we go, here’s what we have so far.

https://www.sciencedirect.com/science/article/pii/S0278434313002781
https://www.sciencedirect.com/science/article/pii/S0278434313002781

By using these two schematics of the West Florida Shelf as background, we can put any carbon or nitrogen values we get into context. Low values of carbon indicate that the fish was far offshore. High values suggest that the fish was close to shore. Low values of nitrogen indicate that the fish was far to the south. High values indicate that the fish was far to the north.

Using these trends as our background, the plot on the left approximately translates into the map on the right. No matter where a fish was caught, there is a high likelihood that it spawned near where the fish icon is on the map for that species. Red Grouper spawn farthest to the south, Black Seabass farthest to the north and inshore. This mostly lines up with what we knew from decades of study, but adds a new dimension to our understanding. For example, there has not been evidence before that Red Snapper were spawning on the West Florida Shelf, but our results are suggesting that they do. There’s lots more cool data!

RPOTS: That IS pretty cool, but I need to go wash my cat, so…talk later?

Two Methods of Collecting Eggs from Marine Pelagic Spawning Species

By Grace Sowaske, University of Florida M.S Candidate

Melanurus Wrasse (Halichoeres melanurus)

My research focuses on species such as the Yellow Wrasse (Halichoeres chrysus), Melanurus Wrasse (Halichoeres melanurus), and the Pacific Blue Tang (Paracanthurus hepatus). All three of these species release their gametes into the water column in a spawning “rise” right around dusk. Collecting these eggs can be a challenge because of the instantaneous, wide dispersal of these eggs throughout the water column. However, the eggs are buoyant so collecting surface water is the most efficient way to obtain the embryos. Here at the Tropical Aquaculture Lab we use two methods in order to collect embryos. I will explain these methods which can easily be mimicked at other facilities who are wishing to begin work with pelagic spawning species.

We house some of our broodstock in harems of 6-17 fish per tank in 2,650L of saltwater. These tanks drain from the surface through an elbow into a smaller egg collection tank which is more of a specialized arrangement (Fig. 1). To collect the embryos, we use a 1 L bucket lined with 250um mesh (Fig. 2). This bucket is then set so that the overflow from the tank flows directly into the bucket before dusk and throughout the night (Fig. 2). The embryos of these species are around 600um and get caught on the inside of the bucket while the water flows through the screen. Algae and uneaten feed also make their way into this bucket. The following morning the eggs are then collected by washing the mesh sides down and concentrating the eggs on the side of the bucket without mesh. This is then poured into a 1L cup for further analysis and screening out of large particulates. (Fig. 5)

Figure 1. The 2,650 L tank is the larger of the two fiberglass tanks with the PVC elbow drain collecting surface water and exiting into the smaller egg collection tank
Figure 2. The egg collection bucket is set before dusk awaiting the evening’s spawn. This will then be detached from the elbow, rinsed down and poured into a 1 L collection cup.

The second method we use to collect eggs is an airlift collector. We use this type of collector in tanks which do not have a secondary collection area. This is the most readily adaptable design for many tank types. The collector (Fig. 3) is also lined with 250um mesh. The bucket is fitted with pool noodles so that it floats at the surface of the water. A PVC clip can be used to secure the collector to the edge of the tank or main standpipe (Fig 4 marked with a star). The collector has a standpipe in the middle, connected to an inlet pipe outside of the collector, forming a continuous U. A small, cylindrical air stone is placed in the bottom of the standpipe. This causes the water to enter via the outer part of the U. The water is then lifted to the top of the internal standpipe by the air bubbles where it overflows into the collection chamber (Fig 4). The eggs are then caught in the collection bucket while the water flows back out into the tank. These collectors are a little difficult sometimes as the inlet pipe must line up correctly with the surface of the water. You also are not collecting the entire water surface, so there is a chance that you are not collecting the entire spawn. Once the embryos are collected, the sides are washed down and poured into a 1 L cup for further analysis (Fig. 5).

Both methods allow for large particulates (uneaten food, algae clumps, detritus etc.) to make their way into the buckets, so further screening after collection needs to be done to clarify the sample. To do this I take the samples and pour them through a large mesh screen (2mm). The eggs are then concentrated through another 250um screen and poured into a small 200ml cup. I then look at the embryos under our dissecting microscope to enumerate them and stock them into larval experiments or rearing trials (Fig. 6). Research using these embryos will focus on optimizing parameters such as tank size, live feeds and larval stocking densities and algal species and densities used for greenwater larviculture in early life stages. These investigations will better define protocols which yield increased survival and growth and can then be transferred to commercial producers.

Figure 3. This is an airlift collector constructed from the bottom of a 5-gallon bucket. 250um mesh pool noodles, and middle pvc pipe.
Figure 4. This is the collector in action with surface water brought in from the pipe on the right via the air bubbles in the middle pipe. The water is sucked through the U and into the bucket, allowed to exit out the mesh sides while the eggs stay behind.
Figure 5. Collection cups with samples ready for further screening and clarification.
Figure 6.  Pacific Blue Tang embryos viewed with the dissecting microscope.

How Can I Help ? Three easy steps you can take every day to help the ocean

By Julie Vecchio, USF, Ph.D. candidate

If you love the ocean, you know it is in danger. There are many threats to the organisms and ecosystems within the ocean, but what can YOU, just one person, do about it? Here are a few suggestions that you can follow every day that can reduce your impact on the oceans.

1.BUY USED

Clothing, shoes, toys, and electronics are all good candidates for buying gently used. In the United States alone, 16.2 million tons of textile waste is generated every year. Less than a quarter of this waste is recycled. Buying used clothing and toys from a thrift store or consignment shop not only saves the materials, water, and electricity to produce the new product, it also saves a large amount of waste from going into landfills (1).

My take on it: I have a toddler who outgrows his clothes and toys about every 6 months! By buying his clothes and toys from a consignment shop, I am not only saving at least 50% on all the “stuff” I buy for him, but I’m also saving those items from being trashed by someone else.

 

2. REDUCE WATER CONSUMPTION

According to USGS each person in the U.S. uses between 100 and 180 gallons of water per day (2). All of this water goes into the municipal sewage stream (if you live in a city). This water is usually funneled toward wastewater treatment plants located on the edge of the nearest body of water then cleaned using several steps. About 90% of the chemicals and nutrients in the water are removed by this process (3), but 10% remains and is released into the environment, eventually making its way to the ocean. These chemicals can have a variety of effects on ocean ecosystems from anoxic zones to hormone disruption (4, 5).

You can reduce your water consumption by a few simple measures. First, turn the tap off while brushing your teeth, or washing your face and hands. Second, follow the “if it’s yellow, let it mellow” rule. Third, take shorter showers or don’t shower as often. Wash your hair less often. If you have an automatic sprinkler system, install a rain gauge to shut down the sprinklers when it has rained. Cities will often do this for you for free. All of these steps will help reduce freshwater consumption, but also reduce the amount of excess nutrients and chemicals that make their way to the ocean.

My take on it: I have spent several years of my life living on sailing or motorized vessels at sea. While many modern ships have desalinization plants to create fresh water, many of the vessels I have lived on did not. Whatever water we brought from the dock was the water we had, for drinking, bathing, cooking, etc. This experience has trained me to think twice before turning on the tap or leaving it running. For instance, I only wash my hair about once a month but I rinse it every time I get in the shower. We only run our sprinkler system when the grass is looking really brown, and I cringe each time it goes on.

 

3. BRING REUSABLE CONTAINERS

By now, the amount of trash (specifically plastics) that end up in the ocean is a problem that is well known to most people. Over 300 million tons of plastic waste was generated in 2015 globally (6). Most of this was single-use plastics or plastic packaging. Many restaurants are starting to ask patrons if they need a straw instead of dropping them off automatically. This is a great step. You can encourage restaurants to do this by asking for “no straw” or even talking to management about making this simple change in their procedures.  Some states and municipalities are even outlawing plastic bags all together (7).

bag

Photo from Shutterstock

My take on it: There are a few quick things I do (or try to do) to reduce my reliance on single-use plastics. First, I (almost) always use my travel mug. I brew my coffee at home in a French press and compost the grounds. If I’m out and I didn’t bring my travel mug with me, I think really hard about whether I need that cup of coffee. Most of the time I decide it’s not worth the trash. Second, I (almost) always bring my reusable bags to the grocery story. This saves somewhere between 7 and 9 plastic bags every week. Third, I pack my lunch every day for work. My lunch bag has a spoon and fork that live inside, I always eat leftovers from dinner the night before that I put in a reusable container, and I have reusable snack packs for my carrots and chips that I throw in the laundry. One thing I want to start doing is bringing my own doggie bag to restaurants. I go to certain restaurants where I know I will want to take some of my meal home with me. I am trying to remember to bring my own container so that I don’t take home a container that I’m just going to put in the trash.

Our best bet for a healthy ocean in the future is for each person to be mindful about their consumption, doing what they can when they can (8). If we all do our part, we can slow the environmental degradation that is threatening not only the ocean, but our livelihoods as well.

ocean.jpg

References

  1. https://www.epa.gov/sites/production/files/2016-11/documents/2014_pdf
  2. https://water.usgs.gov/edu/qa-home-percapita.html
  3. https://www3.epa.gov/npdes/pubs/primer.pdf
  4. http://www.marbef.org/wiki/Endocrine_disrupting_compounds_in_the_coastal_environment
  5. https://www.accessscience.com/content/anoxic-zones/037400
  6. https://wedocs.unep.org/bitstream/handle/20.500.11822/25496/singleUsePlastic_sustainability.pdf?isAllowed=y&sequence=1
  7. http://www.ncsl.org/research/environment-and-natural-resources/plastic-bag-legislation.aspx
  8. https://www.wisebread.com/17-cheap-and-awesome-reusable-replacements-for-disposable-products

 

Exploring Fisheries Aspects of Large-Scale Habitat Restoration in Tampa Bay

By Kailee Schulz, UF Master’s Student

 

Kaileebio

Estuaries form a link between marine and freshwater environments, harboring a rich assemblage of fish and plant species (Attrill and Rundle 2002). Because human population growth is typically highest near the coast and coastal freshwater environments, the loss and degradation of estuarine habitat is a major threat to resident species (Fitzhugh and Richter 2004, Vitousek et al. 1997, Kennish 1991). This is especially true for Tampa Bay, where a growing population of over two million reside within the ~5,700 km2 watershed (Greening et al. 2014, Rayer and Wang 2015). Habitat loss and degradation has led to an interest in large-scale restoration (Yates et al. 2011, Russell and Greening 2015). While improving environmental conditions through reductions in nutrient inputs are well documented, the benefits of restored, reconnected, and created habitats are still poorly understood, despite large initial investments in restoration efforts (Russell and Greening 2015).

The overall goal of my research is to understand how fish communities are utilizing restored habitats. Specifically, are restored sites functioning as suitable juvenile sportfish nurseries? I have two aims within these objectives: (1) describe the relationship between fish communities and habitat at three site types and (2) compare juvenile common snook (Centropomus undecimalis) growth and condition among habitats and sites.

Schulz2

Figure 2. The three restored (Cockroach Bay- restored, Rock Ponds, Terra Ceia), three impacted (Dug Creek, Newman Branch, E.G. Simmons), and three natural sites (Little Manatee, Cockroach Bay-natural, and Frog Creek) located within the Tampa Bay watershed.

To accomplish this, I sampled three impacted, three restored, and three natural sites quarterly (Fig. 2). An impacted site is a historically dredged canal or ditch that received minimal subsequent modification. A restored site is an area that has been physically and biologically modified to restore or create landscape characteristics that support aquatic communities. A natural site is an area with minimal physical and biological alteration to aquatic habitat. Beginning in March 2018, fishes were sampled quarterly at all 9 sites. 9.1 m and 40 m nylon seines were used for up to 9 and 3 samples per site, respectively (Fig. 3 and 4).

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Figure 3. The 9.1 meter net being pulled into the shoreline at Terra Ceia Restoration.

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Figure 4. The 40-meter net fully deployed and being pulled into the shore at Cockroach Bay natural site.

Collected fishes were identified to the species level using methods developed by Kells and Carpenter (2011). All sportfish, fishes of economic importance, and non-native species were counted, measured, and released. A subsample of common snook were retained for later analysis, with a maximum of 45 common snook per site kept during each quarter. These common snook were weighed and measured (SL, FL, and TL). The sagittal otoliths were removed and processed following protocols developed by VanderKooy (2009) (Fig 5). Juvenile snook age was estimated by counting daily growth rings along the sulcus beginning at the core. Two independent readers estimated age for each otolith, with the mean value used for analysis if both estimates were within 10%. Further, total lipid analysis was completed on the retained snook using the standard Folch extraction methods (Folch et al 1956). The age and of these juvenile snook will provide information on the functionality of the three site types. I also collected a variety of habitat parameters based on previous research by FWC’s Fisheries Independent Monitoring program (Table 1) and water quality which, when paired with the juvenile common snook condition, offer insight on the specific environmental conditions that provide functional juvenile sportfish nurseries.

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Figure 5. The ventral side of a juvenile common snook with its two otoliths exposed, the opaque, oval bones sitting within the brain cavity.

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Table 1. Habitat characteristics that are recorded at each seine pull. Unit of measurements with two variables include a species and the amount of space it covered the seined area. Recorded levels refers to the maximum number of parameter types that can be examined.

Thus far, I have caught 49,108 fish, a majority of which were collected at restored sites (Fig. 6). I found a significant difference in the growth rate between the snook caught at the three site types; restored, impacted, and natural. (Fig 7.) These preliminary data show that juvenile common snook grow faster at restored sites and natural sites compared to those at impacted sites. The next step is to evaluate snook body condition at the three site types. I will use the habitat characteristics and growth and condition of the juvenile snook to understand which specific habitat parameters are key in promoting successful nursery environments. The community structure will be evaluated at each site to assess which features promote a functional nursery habitat.

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Figure 6. The number of animals (fish, shrimp, and crab species) caught at three site types. This is standardized by the number of individuals caught per seined m2

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Figure 7. A comparison of juvenile common snook growth between three site types. There was a significant difference in mean growth rate between the three sites (F2,45 = 4.22, p = 0.021). Error bars represent SEM. Letters denote differences among site type as identified as TukeyHSD.

Habitat restoration is often conducted to benefit sportfish with many restorations aimed at improving nursery habitat (Lewis III 1992; Peters et al. 1998). This research will provide information on the parameters necessary in promoting juvenile sportfish success. Another goal for Tampa Bay restoration is enhancement of local diversity by creating and managing for habitat mosaics. To this end, I will compare fish community structure at sites with varying levels of habitat diversity. This research will be useful in future restoration projects and increase understanding of qualities that are important when designing and creating restoration projects. Habitat restoration is increasingly implemented as the human population continues to grow within the Tampa Bay watershed. Ultimately, my work will improve the effectiveness and utility of habitat restoration as it relates to fisheries resources.

 

References

Attrill MJ, Rundle SD (2002) Ecotone or ecocline: ecological boundaries in estuaries. Estuarine, Coastal and Shelf Science 55:929–936

Fitzhugh TW, Richter BD (2004) Quenching urban thirst: growing cities and their impacts on freshwater ecosystems. Bioscience 54:741–754

Folch, J. M. Less, and G.H. Sloane Stanley. 1956. A simple method for the isolation and purification of total lipids from animal tissues. Boston: Harvard University Press

Greening H, Janicki A, Sherwood ET, Pribble R, Johansson J (2014) Ecosystem responses to long-term nutrient management in an urban estuary: Tampa Bay, Florida, USA. Estuarine, Coastal and Shelf Science151:A1–A16

Kells V, Carpenter K (2011) A field guide to coastal fishes: from Maine to Texas. JHU Press, Baltimore, MD

Kennish MJ (1991) Ecology of estuaries: anthropogenic effects. CRC Press, Boca Raton, FL

Lewis RR III (1992) Coastal habitat restoration as a fishery management tool. Pages 169–173. In: Stroud RH (ed) Stemming the tide of coastal fish habitat loss. National Coalition for Marine Conservation Inc., Savannah, GA

Peters KM, Matheson RE Jr, Taylor RG (1998) Reproduction and early life history of common snook, Centropomus undecimalis (Bloch), in Florida Bulletin of Marine Science 62:509–529

Rayer S, Wang Y (2015) Pages 1–8. Projections of Florida population by county, 2015-2040, with estimates for 2014. University of Florida Bureau of Economic and Business Research Bulletin 171, Gainesville, FL

Russell M, Greening H (2015) Estimating benefits in a recovering estuary: Tampa Bay, Florida. Estuaries and Coasts 38:9–18

VanderKooy, S. 2009. A practical handbook for determining the ages of Gulf of Mexico fishes. Gulf States Marine Fisheries Commission Publication 167

Vitousek PM, Mooney HA, Lubchenco J, Melillo JM (1997) Human domination of Earth’s ecosystems. Science 277:494–499

Yates KK, Greening H, Morrison G (2011) Integrating science and resource management in Tampa Bay, Florida. Circular No. 1348. U.S. Geological Survey, Reston, VA

Coastal shark community assemblages

by Clark Morgan, University of North Florida, Master’s student

As animals grow and mature, their needs change. Throughout their life cycle, many animals relocate into different habitats to enhance their own survival, a process known as ontogenetic habitat shifts. Many shark species aggregate by size and life stage.  Smaller, younger sharks are found in “nursery” areas that have a high abundance of food resources and offer protection from predators (e.g. larger sharks). When a shark reaches sexual maturity, it often moves to another location where other like-minded (and bodied) sharks of the same species aggregate. These movements often correspond with changes in resource requirements for larger individuals. Multiple species often share resources, geographic areas, and prefer many of the same environmental conditions that deem a habitat suitable for use and consequently, complex communities are formed. Researching how these communities change in time and space can be a daunting task, but through the use of multiple methodologies, dynamic ecological questions can be answered for many scientific applications. Thus, understanding the relationship between sharks and their environment is crucial for sustainable management and conservation of shark populations (Simpfendorfer and Heupel, 2012).

Many coastal shark species of the southeastern United States are Carcharhinids, a family of fish known as the “requiem sharks.” A characteristic feature of this group is placental viviparity, in which pregnant females provide nutrients to their pups in uterovia placental connections before live birth. As a result, newborn pups have openings in their ventral surface that are essentially equivalent to a belly button in humans. These open umbilical scars heal quickly, but the size of the remaining wound can provide birthday estimations for these animals. Once completely closed, the presence of a healed umbilical scar is still useful for identifying an animal as a young-of-year (YOY), an important life-stage distinction.

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A partially healed umbilical opening on a neonatal sandbar shark (Carcharinus plumbeus)

A shark’s length is the most useful way to determine its life stage, a result of many lethal studies that measured the development of internal reproductive organs at different sizes. Male elasmobranchs possess external reproductive organs known as claspers that are used in the internal fertilization of a female. These structures harden via calcification as an individual matures, which allows for a quick and easy assessment of life stage by a researcher. Quantifying species-specific life stage abundances and the corresponding environmental parameters of their habitats provides the framework for understanding ecosystems. This is becoming increasingly important for coastal ecosystems as the negative results of anthropogenic disturbances such as pollution, increased human populations, and coastal development can result in habitat degradation (Pan et al. 2013).

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A mature male blacknose shark (Carcharhinus acronotus), with visibly large and developed claspers

Another useful method for assessing community interactions is known as Stable Isotope Analysis (SIA), which measures the naturally occurring ratios of heavy and light chemical elements found in animal tissue. Carbon (δ13C) and nitrogen (δ15N)are the most common isotopes measured. δ13C values trace the original base source of dietary carbon of a consumer, while δ15N values are indicative of relative trophic position (Peterson and Fry 1987, Post 2002).These ratios allow researchers to infer trophic levels, niche widths, and temporal foraging patterns, which provide deeper insight into how these communities may be competing for resources.To insure successful application of SIA, one must also consider the varying turnover time of different tissue types. Tissues that are less active in metabolic processing, such as muscle tissue, take longer for a change in diet to be reflected in a consumer’s isotopic signature (~1 year) while blood plasma provides dietary insight on a much shorter time scale (~2-3 months) (MacNeil et al. 2006; Matich et al. 2011). Comparing δ15N values of different tissue types from the same animal reveals temporal dietary changes which are common with ontogeny, while δ13C values indicate spatial dietary changes indicative of movements into areas of different carbon sources.

A combination of ecological factors like environmental characteristics, resource abundance and distribution, and the presence of other competing species influences nearshore habitat use by sharks (Knip et al. 2010).  Considering the known ecological importance of sharks, identifying influential factors on coastal shark habitat use is imperative to understand how shark species will respond to future changes in the environment (Heithaus et al. 2008, Pan et al. 2013, Yates et al. 2015).

References

Matich, P., Heithaus, M.R. & Layman, C.A. 2011. Contrasting Patterns Of Individual Specialization And Trophic Coupling In Two Marine Apex Predators. Journal Of Animal Ecology, 80, 295–304.

Post, D. M. 2002. Using Stable Isotopes To Estimate Trophic Position: Models, Methods, And Assumptions. Ecology 83:703–718.

Peterson, B. J., And B. Fry. 1987. Stable Isotopes In Ecosystem Studies. Annual Reviews In Ecological Systems 18:293–320.

Macneil, M. A., G. B. Skomal, And A. T. Fisk. 2006. Stable Isotopes From Multiple Tissues Reveal Diet Switching In Sharks. Marine Ecology Progress Series 302:199–206.

Heithaus, M.R., Frid, A., Wirsing A.J., Worm, B. 2008. Predicting ecological consequences of marine top predator declines. Trends in Ecology and Evolution 23: 202-210.

Knip, D. M., Heupel, M. R., and Simpfendorfer, C. A. (2010). Sharks in nearshore environments: models, importance, and consequences. Marine Ecology Progress Series 402, 1–11.

Pan, J., Marcoval M.A., Bazzini, S.M, Vallina, M.V., De Marco, S.G. 2013. Coastal Marine Biodiversity Challenges and Threats. Marine Ecologist in a Changing World. 43-67.

Yates, P. M., Heupel, M. R., Tobin, A. J., and Simpfendorfer, C. A. 2015. Ecological drivers of shark distributions along a tropical coastline. PLoSOne 10(4), e0121346.

Simpfendorfer, Colin A., and Heupel, Michelle R. (2012) Assessing habitat use and movement. In: Carrier, Jeffrey C., Musick, John A., and Heithaus, Michael R., (eds.) Biology of Sharks and Their Relatives. CRC Marine Biology Series. CRC Press, London, UK, pp. 579-601.

An assessment of Largemouth Bass fin rays and spines for use in non-lethal aging in Florida

By Summer Lindelien, MS student, University of Florida

Largemouth Bass (LMB) are a highly sought-after sport fish in the state of Florida. Many anglers fish for them recreationally, whereas others study them extensively. I happen to be both a trophy bass angler and a researcher. My love for fishing brought me into this field of study. I have always believed in conserving our bass fisheries for future generations, and when I saw the opportunity to attempt a methodology that has not been applied as frequently to warm-water fishes, I was eager and intrigued. Non-lethal aging of LMB in Florida has not been fully assessed, and it would benefit fisheries scientists and the public to know more about bass population structure (growth, mortality, and recruitment; Strickland and Middaugh 2015), especially when it can be difficult to find and capture a large number of trophy bass during field sampling, and killing fish is not an ideal option.

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Figure 1. A) Clipping a Largemouth Bass anal fin, B) dorsal fin rays thawing prior to being excised, and C) seven fin structures (pectoral rays 3-5, anal spine III, anal rays 3-5, pelvic spine I, pelvic rays 2-4, dorsal spines III-V, and dorsal rays 3-5) properly excised.

For my study, LMB (N= 686) were captured using daytime boat electrofishing on Rodman Reservoir. Sagitta otoliths as well as dorsal, pelvic, and anal fin spines, and pelvic, pectoral, anal, and dorsal fin rays were taken from individual fish (Figure 1). The bony structures were cleaned and stored for later processing. Otoliths were mounted to slides and sectioned with a low speed saw to 0.5 mm in width, and fin structures were mounted in two-part epoxy then differentially sectioned from 0.7 mm to 1.4 mm. These sections were permanently mounted to slides and aged under dissecting or compound microscopes (Figure 2).

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Figure 2. A) Largemouth Bass dorsal rays 3-5 excised and cleaned, prepped for the drying box, B) dorsal spines III-V imbedded in two-part epoxy, C) sectioning a fin structure with the low speed saw, and D) cross-sections (~0.7-1.4 mm) of several fin structures permanently mounted to slides.

Aging the otolith sections (Figure 3) was relatively simple compared to learning how to age each fin structure since fins all grow differently. After aging over 1,000 sections (Figure 4), I was able to identify which fin structure provided the most accurate (between otolith-based ages and fin structure-based ages) and precise (within-reader and between-reader ages) aging estimates by calculating the average percent error (APE; Beamish and Fournier 1981), coefficient of variation (CV; Chang 1982), percent agreement (PA; Sikstrom 1983), and Lin’s concordance correlation coefficient (ρc; Lin 1989; Lin et al. 2002; Lin et al. 2007). I used age biplots (Campana 2001; Figure 5), residual plots, and radii measurements (Murie et al. 2009; Figure 6) to understand age differences and potential reader biases. The dorsal fin spine (n = 122) provided the most precise (PA = 79%; CV = 4.0; ρc= 0.97) and accurate (PA = 77%; CV = 6.5; ρc= 0.98) ages relative to the other fin structures, and therefore was identified as the best potentially non-lethal aging structure.

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Figure 3. A) A 0.5 mm sagitta otolith section from a 10-yr old Largemouth Bass (LMB), B) a sagitta otolith section from an 11-yr old LMB, and C) an otolith section from a 12-yr old LMB.

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Figure 4. A) Dorsal spine section from a 9-yr old Largemouth Bass (LMB), white dots represent enumerated translucent bands, B) anal fin ray section from an age 4 LMB, white arrow represents the end of the first annulus which is a double band, and C) dorsal fin ray section from an age 4 LMB.

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Figure 5. Scatter plot comparison of age estimates obtained from Largemouth Bass sagitta otoliths versus dorsal fin spines for Reader 1. Diagonal line represents comparisons where otolith age = estimated dorsal spine age. Circle size represents sample size of a particular age combination relative to the largest subsample.

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Figure 6.  First and second Largemouth Bass dorsal spine annular radii (n = 114) as a function of age. Horizontal lines = median measurement for annulus 1 and 2. Analysis indicates overlap in young fish (i.e., 1 and 2-year olds). Position of first annulus was not consistent during aging of these young fish, leading to an overestimation of their ages.

I hope to further investigate these potentially non-lethal methods for aging LMB in additional waterbodies around the state of Florida to better understand dorsal spine growth. I also will be testing the survival of LMB after clipping their dorsal spines. I appreciate everything Florida Fish and Wildlife Conservation Commission (FWC) has done for me both as a student and as a biologist. I am grateful to AFS for providing this opportunity for me to share my research. Thank you for taking the time to read about my project, I hope you have plenty of questions. I am open to answering them! Feel free to contact me at summer.lindelien@ufl.edufor more information.

References

Beamish, R. J., and D. A. Fournier. 1981. A method for comparing the precision of a set of age determinations. Canadian Journal of Fisheries and Aquatic Sciences 38:982-983.

Campana, S. E. 2001. Accuracy, precision, and quality control in age determination, including a review of the use and abuse of age validation methods. Journal of Fish Biology 59:197-242.

Chang, W. Y. 1982. A statistical method for evaluating the reproducibility of age determination. Canadian Journal of Fisheries and Aquatic Sciences39:1208-1210.

Debicella, J. M. 2005. Accuracy and precision of fin-ray aging for gag (Mycteroperca microlepis) Master’s thesis, University of Florida, Gainesville, Florida.

Klein, Z. B., T. F Bonvechio, B. R. Bowen, and M. C. Quist. 2017. Precision and accuracy of age estimates obtained from anal fin spines, dorsal fin spines, and sagittal otoliths for known-age Largemouth Bass. Southeastern Naturalist 16:225-234.

Lin, L., A. S. Hedayat, B. Sinha, and M. Yang. 2002. Statistical methods in assessing agreement: models, issues and tools. Journal of the American Statistical Association 97:257-270.

Lin, L., A. S. Hedayat, and W. Wu. 2007. A unified approach for assessing agreement for continuous and categorical data. Journal of Biopharmaceutical Statistics 17:629-652.

Lin, L. I. 1989. A concordance correlation coefficient to evaluate reproducibility. Biometrics 45: 255-268.

Murie, D. J., D. C. Parkyn, C. C. Koenig, F. C. Coleman, J. Schull, and S. Frias-Torres. 2009. Evaluation of finrays as a non-lethal ageing method for protected Goliath Grouper Epinephelus itajara. Endangered Species Research 7:213-220.

Sikstrom, C. B. 1983. Otolith, pectoral fin ray, and scale age determinations for Arctic Grayling. Progressive Fish-Culturist 45:220-223.

Strickland, P. A., and C. R. Middaugh. 2015. Validation of annulus formation in Spotted Sucker otoliths. Journal of Fish and Wildlife Management 6:208-212.

 

 

My Summer Spent in Centipede Bay

by Cher Nicholson, University of Florida

My summer spent as an intern at Nature Coast Biological Station opened my eyes to the direct benefits that field work can bring to the environment. I learned that enhancement projects can be unique with respect to community involvement. The beginning stages of theenhancement project required permitting for the oyster reef location that involved both the Department of Environmental Protection (DEP) and the Army Corps of Engineers. In accordance with regulations, the chosenlocation for the artificial reef needed to be a specific distance from surrounding sea grasses, and it had to demonstrate adequate oyster recruitment capabilities. The site was chosen because it was within DEP’s standards, and experiments with tiles in Centipede Bay showed successful recruitment of oysters (Figure 1).  On deployment day, it was remarkable to see how many Hernando County volunteers contributed their time for prepping and deploying an artificial oyster reef (Figure 2).  After aiding in the construction of the artificial reef in April 2018, I was given the responsibility of monthly monitoring. I chose to measure salinity, temperature, pH, dissolved oxygen, reef area, and oyster recruitment, based on two oyster restoration manuals (Baggett et al. 2015), (Baggett et al. 2014). In addition to monitoring the reef, I was given the opportunity to go outside of my job description as an intern by conducting my own study!

 

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Figure 1. Location for the deployment of the oyster reef in Hernando County, FL

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Figure 2. Aerial view of reef deployment day with volunteer unloading bagged oyster shell from volunteer boats in Centipede Bay, FL

The main question for my study was based on the unique shape of the reef and if there was a difference in flow intensity in each subsection. I wanted to understand if a difference in flow intensity at seven unique sites on the reef had a correlation with oyster recruitment density. With a limited budget, clod cards and cubes made of hardened Plaster of Paris were used to calculate the relative flow intensity of the water based on the amount of dissolution of each cube placed in a standardized location. Clod cards were constructed using an ice tray, Plaster of Paris dry mix, and deionized water (Boizard and DeWreede 2006; Thompson & Glenn 1994). The standardized locations of the seven sites were constructed prior to the deployment of the cubes at 0.2 meters from the bottom. PVC pipe was used to construct the attachment site for the clod card and recruitment monitoring bag (Figure 3). Relative flow intensity of the seven sites were taken over the course of three months during summer 2018, with a deployment of three sets of clod cards. Photos were taken, and the mass was recorded for each clod card before deployment and after being submerged in the water for 72 hours (Figure 3). Clod cards lack the ability to calculate an accurate value for the velocity of the water on the oyster reef, but they offer a good estimate of relative flow intensity (Thompson and Glenn 1994). The relative flow intensity of each site can be calculated according to Thompson and Glenn (1994).

 V=4.31(Wi/Ai).25(Sf1.25/Sn)

Sf= [ 1 – (Wf/ Wi)1/3]/θ

Sn= [ 1 – (Wf/ Wi)1/3]/θ

 Within their equation Wiis the initial mass, Wfis the final mass, Aiis the initial area, θ is time in days, Sf is weight loss data from cards deployed at the site, Snis weight loss data from cards exposed to water from the site.  The relative flow intensity can be calculated for each site using both the values for the clod card calibration (Sn), which accounts for dissolution of the cubes due to being submerged in water alone, and the change in mass of the cubes once submerged at the site and exposed to moving water for three days (Sf) (Thompson and Glenn 1994). This method provided a cost-effective way to demonstrate the relative flow of water within channels and on the reef.

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Figure 3. Recruitment monitoring bag for Site 2 constructed with polyethylene cage material and filled with ten oysters shells that fit dimension requirement. Clod card is pictured in the upper left-hand corner of the bag

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Figure 4. Clod card from Site 1 before (A,B) and after (C,D) deployment in May 2018

Initially, I hoped to compare the relative flow intensity at the seven sites with recruitment data from settling oyster spat, but with the vast amount of sediment that accumulated on the shells, there was no recruitment recorded for the summer spawning season. Instead of looking at recruitment, I shifted my study to analyzing the problem of sedimentation and if there was a correlation between relative flow intensity of the water and sedimentation. The cause of sedimentation on the reef was in question, so we devised a sedimentation experiment where four bags from each site would be cleaned using a water pump while four bags would be left untouched at each site. After, four cleaned bags and four untouched bags were placed interchangeably in a row of eight at each site. Photos were taken of each bag, totaling 56, after the cleaning and a month later.

The increase in percent sediment cover on the cleaned bags (over the course of a month) was not significantly different at each site (Figure 5). Once the clod card calibration value is calculated using water collected from the site, the numerical value for the relative intensity of water flow at each of the sites can be calculated. If there is a difference of relative flow intensity of each site on the reef, then a linear regression can be used to compare the relative flow value with the increase in percent cover of sediment at each site. This study will provide more information on how relative flow relates to sediment buildup on the reef and choosing the most effective sites for an artificial reef.

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Figure 5. Average difference of percent sediment coverage in the experimental and control groups at each of the seven sites. Experimental bags were cleaned, and the control groups were untouched. The experimental bags did not show a significant difference in the percentage of reaccumulated sediments in a one-month period

I am very grateful for the opportunity to be involved with research as an undergraduate at the University of Florida. In addition to the opportunity to conduct my own study this summer, I gained valuable experience in field work, constructing my own materials for the study, and monitoring marsh grasses planted at Linda Pedersen Park as well as those at the nursery in Gulf Coast Academy in Hernando County, FL. I am eager to continue working on projects that both aim to enhance or restore the environment and involve the community.

References

Baggett, L.P., S.P. Powers, R.D. Brumbaugh, L.D. Coen, B.M. DeAngelis, J.K. Greene, B.T. Hancock, S.M. Morlock, B.L. Allen, D.L. Breitburg, and D. Bushek. 2015. Guidelines for evaluating performance of oyster habitat restoration. Restoration Ecology23:737-745.

Baggett, L.P., S.P. Powers, R. Brumbaugh, L.D. Coen, B.M. DeAngelis, J.K. Greene, B.T. Hancock, and S.M. Morlock. 2014. Oyster habitat restoration monitoring and assessment handbook. The Nature Conservancy, Arlington, VA, USA., 96pp.

Boizard, S.D., and R.E. DeWreede. 2006. Inexpensive water motion measurement devices and techniques and their utility in macroalgal ecology: a review. Science Asia 32: 43-49.

Thompson, T. L., and E. P. Glenn.1994. Plaster standards to measure water motion. Limnology and Oceanography39: 1768-1779.