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-20^th 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.

Tampa Bay Habitat Restoration

By: Kailee Schultz, M.S University of Florida

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.

            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).

Fig 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.

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

Fig. 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 body condition 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.  

Fig 5. The ventral side of a juvenile Common Snook with its two otoliths exposed, the opaque, oval bones sitting within the brain cavity.

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.) 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.

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

Fig. 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 research 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 inestuaries. Estuarine, Coastal and Shelf Science 55:929–936

Fitzhugh TW, Richter BD (2004) Quenching urban thirst: growing cities and theirimpacts 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) Ecosys-tem responses to long-term nutrient management in an urban estu-ary: 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, BocaRaton, 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 fishhabitat loss. National Coalition for Marine Conservation Inc., Savannah, GA

Peters KM, Matheson RE Jr, Taylor RG (1998) Reproduction and early lifehistory 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 bycounty, 2015-2040, with estimates for 2014. University of Florida Bureauof Economic and Business Research Bulletin 171, Gainesville, FL

Russell M, Greening H (2015) Estimating benefits in a recovering estuary: TampaBay, 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 Commision Publication 167

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

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

A “How-To” Guide in Mapping: Memoirs of a First Time ArcGIS User.

By: Brent McKenna, FAU M.S

Step One: Find Data, Get Data

As a scientist, this sounds like it will be the easiest step.  Tag some fish, download the tag receivers, get an email, save the data.  Step one done.

In fact, finding and getting the data might be even easier.  Pretend that you inherited a large data set.  Each file was meticulously catalogued and organized.  Before you ever saw the files, fish movements had been divided into spawning and non-spawning seasons then by tag identification number.  Each ID was grouped and arranged chronologically.  This data is easy to consume and handle. 

Step Two: But Wait!  There’s More!

Turns out though, your data is organized great for the last person who used it.  Turns out, maybe not so much for you. 

With every change of hands, the interest in the data changes.  What you want to accomplish, isn’t the same as what the previous user had wanted to accomplish.    

Luckily, there are only some 30,000 lines of detections in your data set.  That should be easy to organize how you need it, right?  Right.

Step Three:  99 Excel Files open, 99 files of excel, take one down, open 4 more, 110 files of excel…on your desktop

A short period of reflection later, you decide that the best way to handle the data is to combine every data set into a single gigantic dataset or separate each data set by some identification parameter.  Turns out that 30,000 rows in Microsoft excel either crashes the computer program or forms one unwieldy file, so that isn’t an option.  Instead, you make each identification number into its own excel file. 

You diligently set about partitioning the data in ways that you need.  Check the tag identification number, copy, and paste.  Open the next file.  Check the tag identification number, copy, and paste.    After the 1000^th row, however, every line starts to blend together.  Your eyes cross.  You philosophize that there is no real difference between 2 and 5 or 3 and 8.  After all, two is but five upside down, and eight is only three next to a mirror.  Moreover, individual fish aren’t that important.  What’s important is that all the fish are the same species.

Alas, each individual fish is important.  Incredibly so.  Three is not eight, and 2 is not 5.  No matter how much you want them to be.  You must continue to organize the data.  Each tagged fish gets its own file. 

After copying over every instance of an ID’s detection, you open the next file, and the next and the next until your desktop is cluttered with files. 

At last, you are done, but don’t forget to save them.   

Step Four:  So Close, but not quite. 

You clicked “save as” for every file you made.  Named them after their respective tag numbers and saved them in a single folder.  Then you backed up that folder onto an external hard-drive.  With all of your files saved, you can load them into ArcGIS.  Your excitement peaks.  The first step is almost done.  Instead of a beautiful map of colored dots in ArcGIS, however, you get an error message.  You try another file.  Get the same error message.  You saved every one of your files in the wrong format. 

You go back and save each file again.  This time after using Google to make sure you know which file format is best for ArcGIS.  This takes a while.    

Step Five:  One drop of water in the bucket

You get that first file loaded into GIS.  Perfectly.  It is easy to manipulate and effective at showing the data that you want it to show.  Now, you just load another 109 files because you want to see your whole dataset. 

Step Six:  You learn something new every 2 hours or so

Next, you try to animate your map.  How do your fish move with time?  You somehow manage to create a time animation of fish you didn’t know you had from 250 years ago before figuring out that the times were saved in the wrong format in your files.  This will be easy to fix, or so you thought.  In the format that ArcGIS needs, you cannot format the whole date and time column at once.  You must reopen that original excel file, change the date and time format there, and resave the file for use on ArcGIS.  You must do this for every new file you made.

Step Seven: Your Computer Crashed.  Do Not Pass Go. 

Finally, everything is going smoothly.  Your data is organized in a way that you can load into your mapping program and do whatever you need to do.  The amount of adjustments made in ArcGIS are minimal.  It seems like you’ve accomplished a major goal in your project.  You’re close to being done.   Excitement abounded.  You make what should be your last click, that click to save.  That final moment before completion when you can show your lab that you can do something!  But…The program doesn’t reply.  Your click is not going through.  You try clicking again…maybe one more time.  Ok.  It froze.  Not a big deal.  Leave it alone.  Let the computer think.  Then, it’s like slow-motion.  Is that blue you see?  Is it blue?  Surely it’s just green or maybe some holdover from staring out the window too long.  Not a chance.

Blue screen of death. 

Your stomach slams into the ground.  Despair and disappoint permeate your body.  Your hands reach towards the sky, you fall to your knees, and scream, “WHY!!”  Cursing every deity of silicon and heavy metals.    

Step Eight: Time to start over

As it turns out, your files corrupted when your computer crashed.  Time to redo everything.  You text your friends.  You won’t be going to the bar with them tonight.  In fact, let’s cancel the weekend’s plans.  You have a lot of work to do again.  But this time, you learned your lesson.  Every 30 seconds, you click that save button.

Step Nine: What I Learned, a list of Pro-tips:

1. Consider getting a Mac. 
2. Save everything in the right file format the first time. 
3. Write your hypothesis and how you plan to test it on a piece of paper.  Then post that paper on the wall behind your computer.  Make sure you adhere it along the center line of your computer screen so that you can see it even when you go cross-eyed.   
4. Thank the people who worked with the data before you, a lot.  They worked hard to turn the data into what it is now.  They made your work that much faster and simpler. 

Do you know the history of your field site?

By: Cody Eggenberger, FIU M.S Candidate.

From nukes to murder to drug smuggling, the Everglades have had an interesting history to say the least. Since moving to south Florida, apart from the deep love I’ve developed for the recreational fish species I am lucky enough to research, I’ve developed a fascination with the history of the Everglades. Unfortunately, very few places still exist in the US that are able to give you the feeling that you’ve time traveled to a prehistoric, untouched past, where reptilian dinosaurs larger than boats and monstrous schools of fish larger than humans lurk beneath the water’s surface. Anyone who knows anything about the terrible ideas us Homo SAPIENS have had in the past regarding the Everglades knows that the ecosystem has been drastically altered and much of the beauty we see today is a mere shadow of what it once was. From experience and communication, the trials and tribulations that always seem to coincide with field research often causes graduate students conducting field research to form love/hate relationships with their study sites. I’ve developed a deep love for the Everglades, but more specifically, a deep love with one field site in particular. Aside from the beauty it holds, much of this love comes from learning its troubling history with man.

Figure 1. Look carefully and you’ll notice the dinosaur that competed for size with the 14’ jronboat we were using to do an acoustic receiver download in the Alligator Creek subestuary. The girth of its tail was as thick around as an average man’s torso.

The Alligator Creek subestuary stretches from north-central Florida bay past Garfield Bight, to about 7 miles north of Flamingo. The subestuary consists of four large mangrove-lined lakes that are connected to each other by long, meandering and overgrown creeks dense with spider webs, snags, and the whispers of the historic Everglades. This subestuary used to be a hot spot for waterfowl hunting. So much so that old timers have been quoted as saying that the usually remote and quiet lakes would “sound as if a war had broken out” with the amount of gunshots produced from waterfowl hunters gunning down the hundreds of thousands of waterfowl that would migrate to the system every year in winter months. This, of course, is no longer the case as hunting in Everglades National Park is illegal and the majority of the Alligator Creek subestuary is now protected by fairly strict regulations. Most of the subestuary is now a pole/paddle zone meaning that it’s illegal to propel a watercraft in the lakes with the use of an engine. While this makes getting into much of the subestuary very exhaustive and time consuming, one lake’s protective regulations go beyond this and is off limits to anyone without a permit to enter. This is Cuthbert Lake.  

Figure 2. West Lake hunting camp, late 1930s. Buddy Roberts, an Everglades pioneer, was interviewed in 1985 at age 96 and stated, “I seen West Lake and Coot Bay, that place be 200 cars bumper to bumper for 2-3 miles there. All of them hunting, and West Lake, and East Lake, and Cuthbert Lake, and all those lakes back there, The Lungs they call it on the map. It sounded just like war. “

Cuthbert Lake lies in the northeast corner of the Alligator Creek subestuary and nowadays, often has enormous crocodiles sunning on the mud flats and sub-adult tarpon rolling in the channel in front of the overgrown and hard-to-find creek entrance. Cuthbert Lake looks very similar to the other lakes in the Alligator Creek subestuary, but Cuthbert Lake is different in that it has a fairly dark and bloody history. In the late 1880’s and early 1900’s, many may know that it was fashionable in New York and London for women to don bird plumes on their hats. The more “airy” and “floaty” the feathers, the more desirable they were and many of the wading birds in the Everglades, such as Snowy Egrets, develop such feathery plumes during nesting season. At the time, plume feathers were worth about double that of gold by weight and 1oz of plume feathers would sell for about $30. George Cuthbert, apart from being a ship captain and fisherman, was a plume hunter.

Figure 3. Slowly motoring through a mangrove creek on a rainy afternoon of fieldwork in the Alligator Creek subestuary.

Seminole Indian rumors of enormous rookeries existing in the Everglades, composed of thousands of wading birds, drove Cuthbert to sail 80 miles south from his Marco Island home to the narrow creek opening of the Alligator Creek subestuary. Cuthbert waded across mud flats littered with crocodiles and bushwhacked upcurrent through dense mangrove creeks for days while living out of his canoe. The further north Cuthbert trudged, the more wading birds he would see flying to and from a remote region in the distance. Cuthbert followed the flight paths of the birds and was led to the hidden creek mouth of the mysterious lake that would later be named after him and ultimately cause the murder of the first game warden in the Everglades (but that is another story). Emerging from the northernmost creek of his journey, Cuthbert found an expansive lake with a 2-acre mangrove island in its northeast corner. From a distance, the island looked as though it was plagued with white flecks. As he paddled closer, thousands of great and snowy egrets, ibis, wood storks, tricolored herons, and roseate spoonbills came into focus. He later told his children that he had found his “flower, a beautiful white blossom.” The nesting birds were sitting “ducks”. Cuthbert quietly tied his canoe to a branch on the island, grabbed his rifle, and prepared, hidden amongst the mangroves. Plume hunters of Cuthbert’s day were not what one today would refer to as a conservation-minded “sportsmen”, not by any stretch of the imagination; they were ruthless harvesters. Cuthbert shot the adult birds, scalped the plume feathers from their heads, tossed the bodies aside, and left the orphaned chicks to starve. Cuthbert had found his legendary honey hole and like many others to come, littered it with carcasses.  

Cuthbert’s first two trips yielded him about $2,000 worth of plume feathers which equates to about $50,000 today. A few trips later and Cuthbert had made enough money to buy half of Marco Island and comfortably retire. Cuthbert and his family have since sold most of the land they acquired on Marco Island, but it’s estimated that their property would now be worth around $5 Billion. Of course, Cuthbert’s good fortune made a stir in the plume hunting community and others later found the legendary rookery and decimated the wading bird populations. One local plume hunter was quoted as saying that “you could have walked around the rookery on the bodies of the dead birds”.

Fortunately, the Migratory Bird Treaty Act was passed in 1918 and the plume trade died shortly after.  While I’ve never really been much for history, learning about the gruesome and exploitative history of one of my research sites has been very interesting and I think that the knowledge of its past has led me to appreciate the Alligator Creek subestuary’s beauty that much more. I encourage any graduate students who have read this to try to do the same and look into the history of their own field sites. You never know what you may find.

Tips on Researching Pre-collected Data

By: Lauren Kircher, Florida Atlantic University

When joining a new lab, you may sometimes run into a situation where you “inherit” data. Maybe there’s not time for the other person to run analysis, maybe more data needs to be collected, maybe they finished their study, but more can be done. This happened to me when I started my Ph.D. Inheriting data definitely has advantages. You can never have a bad field season and need to continue sampling. You don’t have to wake up super early to get the boat ready to take out. However, there are a fair amount of drawbacks as well. It can be difficult to work with someone else’s data, figure out how it is set up, etc.

• Organization is key.

The researchers who made the measurements and collected the data probably had a standardized format to record everything. Make sure that you learn how the data is structured. It may be more useful for you to format the data a different way just be  sure that you are organized and save the original data separately.

• Keep in touch with the people who collected the data.

One of your most important resources will be the initial researchers. You may think you have a handle on everything when you start, but questions always come up along the way. They can provide insight into how the initial study was conducted, any concessions they had to make designing the study, or if the procedure changed during the study.

• Check the temporal/spatial scale and units of collection.

Make sure that if you are combining data sets they have the same scale and units. Units are an easy thing to convert, but issues of scale may be harder to resolve.

• Don’t feel limited to just use that data.

You may have been given biological data, but that doesn’t have to be the only data you use in analysis. There are plenty of organizations (NOAA, South FL Water Management District, USGS, EPA, etc.) that host open-source environmental databases. Dataloggers measuring abiotic factors are verified, recorded, and stored for years. From the comfort of your own home, you can download data from decades of measurements.

• Brush up on your coding skills.

When organizing your data, especially if it contains years of measurements, it is handy to be able to code in R, Python, Excel, or other programs. You can set up codes or equations to perform functions that you would otherwise perform manually. This will save you time and frustration.

• Leave plenty of time for getting things done.

When communicating with scientists, you must leave plenty of time for them to respond or find other documentation and data for you. They are busy professionals with their own current research. You will also inevitably run into errors and obstacles while working with the data. You may need to reformat files, search for errors in the data, reference other data.

Remote Sensing of Oyster Reefs

By: Michael Espriella, M.S candidate, University of Florida

Oyster reefs filter pollutants, serve as habitat for hundreds of species, and control shoreline erosion among numerous other ecosystem services. Unfortunately, these resources are in decline due to various anthropogenic and environmental stressors including over-harvest, disease, and low freshwater flow events. Given the difficulty in accessing these habitats, there is very limited monitoring to assess declines and their causes.

That’s where remote sensing comes in to the conversation. Not only do remote sensing techniques have the benefit of collecting data on areas that are difficult to access, but they also allow information to be collected without potentially harming the reef as can sometimes be the case with more traditional transect sampling (Figure 1).

Figure 1: Transect sampling counting live oysters off the coast of Cedar Key, FL.

Unoccupied aircraft systems (UASs) are one of the most cost-effective techniques to accomplish this task. They can collect data virtually any day of the year, assuming appropriate conditions. Additionally, they can collect data at a much higher spatial resolution than commonly used remote sensing data sources, such as satellite imagery. This high spatial resolution will allow for more detailed analysis on the state of an individual reef.

Figure 2: Example of UAV imagery mosaic from Little Trout Creek, located north of Cedar Key, FL. Imagery courtesy of Dr. Peter Frederick’s lab at the University of Florida.

Our lab’s objective with this project is to use high-resolution imagery to generate mosaics and delineate inter-tidal habitats with a focus on oyster reefs. This will be done along Florida’s Big Bend coastline using a Geographic Object Based-Image Analysis (GEOBIA) technique. Borrowed from terrestrial remote sensing, GEOBIA is particularly useful when processing high-resolution imagery as it allows for the segmentation of pixels into meaningful objects. These objects are then classified using spectral, structural, and topographical characteristics. From there, we can assess the spatial dynamics that contribute to a successful or unsuccessful system.

Overcoming Nutritional Bottlenecks in Freshwater Ornamental Fish Larviculture

By: Taylor Lipscomb, Ph. D candidate, University of Florida

The focus of my doctoral research is early larval nutrition in select taxa of neotropical teleost fishes that are important to the freshwater ornamental aquaculture industry of Florida. Eleven species from six families (Cyprinidae, Characidae, Cichlidae, Callichthydae, Mochokidae, and Osphronemidae) are the subjects of this research. For initial feeding during early larval aquaculture production in these fishes, live Brine Shrimp (Artemia spp) nauplii predominate (Sales and Janssens 2003). These live organisms provide adequate nutrition for growth and survival, and their ease of culture relative to other live feed organisms have led to their near ubiquitous use in the industry (Lavens and Sorgeloos 2000). Recently, though, variation in the availability of Artemia due to environmental perturbation of the cues for cyst formation in wild stocks, as well as demand from other sectors of aquaculture, have inflated prices and led to inconsistent availability (Kolkovski 2001). In light of these developments, research into potential alternatives is increasingly warranted.

Initial dietetics research has evaluated survival, growth, and homogeneity of growth in the subject species reared using three commercially available microparticulate diets compared to the current standard Artemia from the initiation of exogenous feeding to 14 days beyond that time point. Comparable or superior survival, growth and homogeneity of growth for species of Cyprinids, Callichthyds, Mochokids, Osphronemids and Cichlids fed microparticulate diets relative to Artemia provides evidence for the successful detection, ingestion, digestion and assimilation of nutrients from the manufactured diets. However, optimization of feeding protocols by incorporating feed attractants, as well as the evaluation of potential impacts of feed items on digestive enzyme function, is still warranted for these taxa, and will be the focus of further research going forward. Feed attractants, namely some free amino acids and nucleotides, have been shown to elicit gustatory endocrine and subsequent behavioral responses in other species of farmed fish (Kubitza et al. 1997; Olsén and Lundh 2016), leading to increased feeding incidence and growth. Modulation of digestive enzyme activity has been used to indirectly evaluate digestibility of feeds, by inference of established positive feedback mechanisms for enzyme production when appropriate substrates are present in the gastrointestinal lumen (Zambonino Infante and Cahu 2007).

In species of the family Characidae that were evaluated in initial dietetics studies, significantly lower survival was achieved using microparticulate diets when compared to Artemia, indicating a physiological incapacity for the use of nutrients in the manufactured diets. Similar reliance on live feed organisms is present in many marine teleosts at the onset of exogenous feeding, which exhibit delayed development of the stomach and concurrent gastric secretion of pepsin and hydrochloric acid (Rønnestad et al. 2003). From an evolutionary perspective, this delay in development may have arisen because many of the invertebrate zooplankton present in the marine environment that are feed items for larval teleosts contain a relatively large reservoir of both free amino acids and small, soluble polypeptides when compared to their freshwater counterparts, and particularly compared to manufactured feeds (Rønnestad et al. 2003; Hamre et al. 2013). I hypothesize that the apparent reliance on live feed at the onset of exogenous feeding in ornamental Characids is due to a similar delay in gastric development, which has indeed been observed in other characiform fishes (Portella et al. 2014).

Black Tetra (Gymnocorymbus ternetzi)

Neon Tetra
(Paracheirodon innesi)

Research involving characterization of ontogenetic development of the gastrointestinal tract (Figure 1) and associated enzymes (Figure 2) in Characids has been conducted, which confirmed this protracted agastric phase in larval development. Future research will focus on optimization of weaning using the information elucidated from these observations. The appearance of the stomach, associated gastric glands, and the onset of pepsin activity will be used to inform the optimum time to introduce mircroparticulate diets and begin concurrent reduction of Artemia feeding in these species.

The goal of this research is to optimize early larval nutrition in ornamental freshwater fish, which will lead to advancement in the aquaculture practices of Florida tropical fish farmers, as well as contribute to the collective understanding of early teleost development.

Figure 1: Photomicrographs of G. ternetzi gastrointestinal tract at 2dph (a,100x), 17dph (b, 100x), 22dph (c, 100x), and 22dph (d, 400x). Esophagus (E), mucous cell (MC), stomach (S), gastric glands (GG), pyloric valve (PV), ileo-rectal valve (IRV), rectum (R), exocrine pancreas (EP),

Figure 2: Development of digestive enzyme activities (trypsin (a), lipase (b), pepsin (c)) of G. ternetzi fed exclusively newly hatched Artemia nauplii between 2 and 24dph. Error bars are absent as each data point represents enzyme activity of a pooled sample of ten larvae standardized to enzyme activity per fish.

References

Hamre, K., M. Yúfera, I. Rønnestad, C. Boglione, L. E. C. Conceição, and M. Izquierdo. 2013. Fish larval nutrition and feed formulation: Knowledge gaps and bottlenecks for advances in larval rearing. Reviews in Aquaculture 5(SUPPL.1).

Kolkovski, S. 2001. Digestive enzymes in fish larvae and juveniles—implications and applications to formulated diets. Aquaculture 200(1–2):181–201.

Kubitza, F., L. L. Lovshin, and R. T. Lovell. 1997. Identification of feed enhancers for juvenile largemouth bass Micropterus salmoides. Aquaculture 148(2–3):191–200.

Lavens, P., and P. Sorgeloos. 2000. The history, present status and prospects of the availability of Artemia cysts for aquaculture. Aquaculture 181(3–4):397–403.

Olsén, K. H., and T. Lundh. 2016. Feeding stimulants in an omnivorous species, crucian carp Carassius carassius (Linnaeus 1758). Aquaculture Reports 4:66–73. Elsevier B.V.

Portella, M. C., R. K. Jomori, N. J. Leitão, O. C. C. Menossi, T. M. Freitas, J. T. Kojima, T. S. Lopes, J. A. Clavijo-Ayala, and D. J. Carneiro. 2014. Larval development of indigenous South American freshwater fish species, with particular reference to pacu (Piaractus mesopotamicus): A review. Aquaculture 432:402–417. Elsevier B.V.

Rønnestad, I., S. K. Tonheim, H. J. Fyhn, C. R. Rojas-García, Y. Kamisaka, W. Koven, R. N. Finn, B. F. Terjesen, Y. Barr, and L. E. C. Conceição. 2003. The supply of amino acids during early feeding stages of marine fish larvae: A review of recent findings. Aquaculture 227(1–4):147–164.

Sales, J., and G. P. J. Janssens. 2003. Nutrient requirements of ornamental fish. Aquatic Living Resources 16(6):533–540. University of Florida.

Zambonino Infante, J. L., and C. L. Cahu. 2007. Dietary modulation of some digestive enzymes and Metabolic processes in developing marine fish: Applications to diet formulation. Aquaculture 268(1–4 SPEC. ISS.):98–105.

The History of Lake Alice

By: Marina Schwartz, M.S, University of Florida

Lake Alice, a revered cornerstone of the University of Florida campus in Gainesville, Florida holds many great memories for students, faculty, staff, and alumni. It seems like everyone at UF has a tale about “one time, at Lake Alice…”, but even the best of those don’t encompass the whole story. So here is a short history lesson on everyone’s favorite mascot home, Lake Alice.

Lake Alice is a 33-ha closed-basin lake/marsh system with a watershed of 461 ha that covers approximately 60% of UF’s main campus. The open-water portion (10 ha) is located at the west end of the basin, while the east end of the basin is a marsh (Figure 1). The lake-marsh system is now UF’s primary treatment system for nonpoint source runoff. Lake Alice is a sinkhole depression with a maximum depth of approximately 3 m. The mean depth of the open-water portion of the lake is < 1.3 m. (Korhnak 1996).

Figure 1. Satellite view of Lake Alice (Gainesville, Florida) and its adjoining marsh in 2018. Image from Google Maps

Lake Alice began as a small farm pond (1 ha) and was called Jonas’ Pond until it was renamed in the late 1800s. Many believe the lake was named after the farm owner’s daughter, Alice, but no such record of an Alice Witt exists. UF acquired the land in 1925 as part of an agricultural experiment station (Wells et al. 1996). Shortly after, UF built a wastewater treatment plant and discharged the unchlorinated effluent into Lake Alice (Guard 1932). In 1947, the effluent from the treatment plant was routed into a nearby sinkhole instead of the lake. The following year an earthen dam was constructed on the West side of the lake to help control flooding (Karraker 1953), this expanded the size of the lake to 36 ha. In the early 1950s, the lake area continued to flood and expand until 1959, when two drainage wells, the North and West wells were installed to allow the lake to reach a fixed stage and then overflow into the wells to drain lake water into the deep Floridan aquifer (Karraker 1953, University of Florida 2015).

During the 1960s, the sinkhole that received the unchlorinated effluent was sealed off and the effluent was once again routed into Lake Alice through the marsh at 3,800 to 7,600 m³/day, along with cooling and condenser water at a rate of 38,000 to 45,600 m³/day from UF heating plant #2 (Brezonik and Shannon 1971, Korhnak 1996). In the late 1960s, a large infestation of Water Hyacinth (Eichhorina crassipies) occupied most of the lake’s surface area (Figure 2, bottom left).

Figure 2. Historical aerial photographs of Lake Alice from University of Florida Digital Collections (University of Florida, 2018): (top row left to right) 1937, 1949, 1956, 1961,(bottom row left to right) 1969, 1974, 1990, 1994

In 1976, the heating plant water was diverted away from Lake Alice. UF’s Introduction to Fishery Science class began collecting water quality and fish data from the lake in 1988. In 1994, a new UF advanced wastewater treatment reclamation plant was built, and the treated effluent was routed directly into the northern drainage well and no longer into Lake Alice as of November 1994 (Korhnak 1996). In January 2001, a severe cold weather event resulted in a die-off of ~ 12,000 to 15,000 1.8 Kg Blue Tilapia (Oreochromis aureus). In 2002 and 2003, UF began managing aquatic submersed and emergent macrophytes in Lake Alice with herbicide applications and mechanical harvesting. In 2004, UF discontinued discharging 3,800 m³/day of cooling water from the Reitz Union to Lake Alice (University of Florida 2015). Additionally, severe weather events from three major hurricanes happened that same year. In 2005, UF stocked 500 Grass Carp (Ctenopharyngodon Idella) into the lake to control the growth of rooted submerged aquatic macrophytes. There was a low water event due to a severe drought in the Spring of 2011, and another, less severe low water event in the Spring of 2017. In January 2018 there was another Blue Tilapia die-off less severe than the event in 2001.

As for the management of Lake Alice… it’s complicated. The lake has multiple, conflicting classifications, such as a Class III water body, a stormwater management system, and a university-designated conservation area. Each of these designations has potentially conflicting goals in terms of policy and management. Although primarily managed for aesthetics, there is still conflict surrounding the nutrient input from the watershed. Lake Alice has a robust history of physical and hydrological changes due to management efforts, but regardless of nutrient input or physical alterations, Lake Alice will remain a cornerstone of the UF campus for many years to come.

Most of the information included here was compiled from a Master of Science thesis titled ‘Water, Phosphorus, Nitrogen, and Chloride Budgets for Lake Alice, Florida, and Documentation of the Effects of Wastewater Removal’ (Korhnak 1996) and extensive personal communications and the unpublished data of Dr. Daniel Canfield, Jr. and Dr. Charles Cichra, who have taught classes on Lake Alice since arriving at UF.

References

Brezonik, P. L., and E. E. Shannon. 1971. Trophic state of lakes in north central Florida. Florida Water Resources Research Center, Report B-004-FLA.

Guard, C. J. 1932. Some problems in the control of sewage treatment. Master’s Thesis. University of Florida, Florida

Karraker, D. O. 1953. The birds of lake Alice A comparative study between disturbed and undisturbed areas. Master’s Thesis. University of Florida, Florida.

Korhnak, V. L. 1996. Water, phosphorus, nitrogen, and chloride budgets for lake Alice, Florida, and documentation of the effects of wastewater removal. Master’s Thesis. University of Florida, Florida.

University of Florida. 2015. Campus master plan 2015-2025: General infrastructure data and analysis. :9-5-9-7.

Wells, O., T. T. Ankerson, M. Clark, M. Aldridge, G. West, and Q. Miralia. 2006. Waters of the university of Florida: A plan for achieving sustainable water management in the Lake Alice watershed. University of Florida Levin College of Law, Gainesville, Florida.

The work-life balance might be a seesaw

By: Gabrielle Love, M.S, University of Florida

It’s the middle of the semester. I’m taking two classes and developing my research project for my thesis. I spend plenty of hours reading about the many ways that a snail can eat an oyster. Some days I go to class in the morning, and in the evening, I sit down at my desk to do homework and read guides on how to actually get any writing done. But I don’t go home right after class – I go to work.

            In addition to being a graduate student, I am a full-time employee of the Florida Department of Agriculture and Consumer Services. There I work on research for the eradication of the invasive giant African land snail. I love my work, and I love that my work has allowed me to become a graduate student. I also love free time, but sometimes, that is in short supply.

All graduate students are well aware of the need for down time and the relief of getting a true break from the work and the studying. So how do you maintain any free time when it seems that all of your time is being demanded by other responsibilities? It certainly isn’t easy.

A typical day in my week involves morning classes, professional duties until the workday ends, then schoolwork until bed. For me, weekends are a crucial time to decompress. That’s not to say that I don’t do any work on these days (because I definitely do), but I try to limit myself. If I don’t, I know that I will get caught up in the next thing on my To-Do List and never stop working. It is important for me to get as much done during the week as possible, so the weekends can remain free and I can remain sane.

I have a list of tools and tips that help keep me on track with my schoolwork, while also accounting for the time I need to devote to my job and attempting to maintain the elusive work-life balance.

1. Prioritize Things – It’s easy to do all of the little things on my ever-expanding list of tasks, and they often are important, too. But if I focus too much of my time on those things, then my project proposal will never get done.
2. Keep a Calendar or Schedule – This is crucial for me. Knowing important dates and upcoming deadlines allows me to plan ahead and make sure that I can budget my time however I need to.
3. Set Personal Goals and Deadlines – In keeping with the above, this helps set a pace for my work and keeps me accountable, so I don’t end up trying to complete a term project two days before it is due.
4. Let Yourself Mess Up – No one is perfect. I know that I have procrastinated, turned in sub-par work, and neglected personal responsibilities because I just didn’t have the time. That’s ok. Making mistakes is normal, and it’s never as bad as the anxious voice in our heads might think.

It’s called it a work-life balance, and that is what I strive for, but I know that the scale will tip back and forth. It takes time and practice and mistakes to find that balance, and it’s always changing. My plan is to enjoy the times when it tips in my favor, and when it doesn’t, pray that the workload doesn’t get heavier – because it will.

How to catch a Guppy: A journey into the mountains of Trinidad

By: Allison Durland-Donahou, Ph.D Candidate, University of Florida

Figure 1. The Guppy Poecilia reticulata (Deacon et al. 2015).

The Northern Range mountains of Trinidad are home to a well-studied, model species: The Guppy Poecilia reticulata (Figure 1). This mountain range runs along the northern border of the island of Trinidad. Streams flow from the mountains into the plains below and either east into the Atlantic Ocean or west into the Gulf of Paria (Figure 2).

Figure 2. Map of northern Trinidad indicating the location of the Northern Range mountains and the connectivity of streams

Barriers to dispersal of predators caused divergent evolution in the Guppy within streams. Waterfalls act as natural barriers, causing unique populations between upstream and downstream locations. Predation level is determined by the predator composition of the site with high predation sites containing the Pike Cichlid Crenicichla frenata and low predation sites containing the Jumping Guabine Anablepsoides hartii (Figure 3). For the purposes of my research, I collected from four unique populations (four streams) that had similar human disturbance and predation level (Figure 4).

Figure 3. The Jumping Guabine Anablespoides hartii.

Figure 4. The sampling site at Lopinot. All four stream sampling sites were similar: high human disturbance and moderate predation risk.

The collection adventure begins by driving on (the left side of!) winding, narrow, pot-holey roads through small towns with gorgeous views of the Northern Range (Figure 5). Once at the sampling sites, which are just as gorgeous as the drive, scientists scan the clear stream water for schools of guppies. Guppies tend to school in the shallow pools of streams. Wading through the water, the scientist uses a modified seine to corral the schools towards the shore and then encircles them (Figure 6). The net is then set on the shore in order to pick out the guppies and place them in buckets (Figure 7). Sometimes the pools are too small to use a seine, so an aquarium net can be used to collect guppies (Figure 8). Other fish are sometimes caught with the guppies, such as the native Trinidad Pleco Hypostomus robinii (Figure 9).

Figure 5. View from the road to a sampling site

Figure 6. Kharran and Allison using a seine to catch guppies

Kharran and Amy picking fish out of a seine

Figure 8. Allison using an aquarium net to catch guppies

Once the number of fish needed have been collected, buckets of fish are strapped into the bed of the truck and the bumpy trek back to the lab ensues (Figure 10). It is essential to have a passenger keep an eye on the buckets to make sure they stay upright. Always remember to take a proud scientist picture at the end of a successful collection day (Figure 11)!

Figure 9. (most likely) A juvenile Trinidad Pleco Hypostomus robinii – a native fish that co-occurs with the Guppy

Figure 10. Buckets with guppies strapped in and ready for the bumpy ride back to the lab.

The guppies collected in Trinidad are being used in a series of experiments to compare life history and behavioral trait tradeoffs between domesticated and wild guppies.

For more information about this project, contact Allison at adurland@ufl.edu.

My sincere gratitude to Amy Deacon and Kharran Deonarinesingh for their knowledge and assistance in the field and lab and to Kharran for driving the Trinidadian roads so I didn’t have to!

Allison and Amy after a successful day of Guppy sampling

Oyster Reef Water Quality

By: Mel Moreno M.S, University of Florida

My name is Mel Moreno, and I am a graduate student at the University of Florida. Prior to becoming a graduate student, I was a full-time employee working on the Lone Cabbage Reef Restoration Project at the University of Florida. This project involves restoring an oyster reef to historic conditions. This oyster reef is located within the Big Bend, near Cedar Key, Florida. You can read more information about the project here https://wec.ifas.ufl.edu/oysterproject/. I was in charge of handling water quality data collected around the reef during my work on the project. At that time, with the help of others at the University of Florida Library, we were able to create a workflow to ensure quality control and securement of these data. Because of this effective workflow, we are able to report data analysis from the water quality sensors in 5 business days. You can locate the work I have managed here https://lcroysterproject.github.io/oysterproject/. My current research entails ensuring the workflow is as effective as it can be and documenting the final product.

Me in the field downloading water quality data.

Map of Oyster Reef and water quality sensors

Constructed oyster reef (Credit: Carlton Ward Jr., Florida Wild)