Sheepshead Shuffle 2020

Hello Fellow AFS Members,

The FL AFS Student Subunit invites you to join us for the annual Sheepshead Shuffle Virtual 5k to celebrate World Oceans Day and give back to your community of fisheries scientists!

The Sheepshead Shuffle is a fun, virtual event where you participate by walking, running, or shuffling (whatever mode of exercise you want!) 5k at the location of your choosing. Participants are encouraged to track their times to compete with other shufflers. Prizes for the fastest shuffler, slowest shuffler, and most interesting location (within limits since we’re all supposed to be staying close to home) will be awarded.

Participating in the Sheepshead Shuffle is not only the perfect opportunity to get outside, exercise, and de-stress during these uncertain times but it also allows you to donate to one of the AFS Student Subunits whose chapter meeting got cancelled this spring. All proceeds from registration will fund student travel grants to AFS meetings for members of those Student Subunits.

How to participate? Register at sheepsheadshuffle5k2020.eventbrite.com and be sure to indicate which Chapter you are representing!

You can complete your 5k in as many legs as you want between April 6th and June 14th. Post your best #SheepsheadSelfie to the FL AFS Student Subunit Facebook or Instagram with your time (or just email them) and you’ll be eligible to win prizes.

Make sure to check out our Facebook and Instagram for other’s #Sheepshead Selfies (we promise there will be pets and kids in selfies)!

Any questions? Please contact the FL AFS Student Subunit at flafsstudent@gmail.com.

Happy Shuffling!

FL AFS Student Subunit 

Facebook page: https://www.facebook.com/AmericanFisheriesSocietyFlStudentChapter

FisheriesBlog: https://flafsstudentsubunit.wordpress.com/

Email: flafsstudent@gmail.com

Amazon Smile: http://smile.amazon.com/ch/52-1208319

Instagram: @flafsstudentAsk us to add you to our listserv!

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.

President

Lauren Kircher

CroppedLaurensnook

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

Vice President

Brent McKenna0818201228c_HDR

Brent is a Master’s of Science student at Florida Atlantic University.  His interests lie primarily with fishing and the best ways to ensure the survival and success of fishes.  He believes that fishing is one of the best ways to interest people in conserving fishes. Thank you to everyone in Dr. Baldwin’s lab for their experience and friendship.

University Liaison

Casey Murray

casey

Casey Murray is a PhD student at the University of Florida’s Tropical Aquaculture Lab working with Dr. Matt DiMaggio. Her research interests include larval fish nutrition, gut enzyme ontogeny, and improving larval feeding protocols. Casey is studying both freshwater and marine ornamental fishes to develop species-specific feeding protocols and weaning schedules. Casey received her B.A. in Biology from St. Mary’s College of Maryland in 2015 where she discovered her passion for ornamental aquaculture during her senior thesis research on the determination of juvenile Banggai cardinalfish habitat preference.  Casey graduated from the University of Miami with a Master of Professional Science degree in 2017 where she studied the factors affecting loggerhead sea turtle hatch success in Everglades National Park.  Prior to starting at the Tropical Aquaculture Lab in 2019, Casey worked at Roger Williams University where she helped culture Atlantic lookdowns, glassy sweepers, and smallmouth grunts along with researching alternative protein sources in salmonid feeds.

In her spare time, Casey enjoys traveling, baking and spending time with her pet duck, Tiny.

Secretary/Treasurer

Matt Woodstock

Woodstock_headshot

Matt Woodstock is from Footville, Wisconsin where he received a B.S. in Ecology, Evolution, and Behavioral Biology from Beloit College. During the summer of his junior year, he participated in shark and stingray tagging research near Clearwater, Florida and decided to follow the Marine Biology career path. He received an MS in Marine Biology from Nova Southeastern University and is currently a PhD student at Florida International University. While at NSU, he worked for the Broward County Sea Turtle Conservation Program, taught at a local museum, and found a love for deep-sea fishes. For his PhD, Matt is developing ecosystem models for the oceanic (seaward of the 1000m isobath) Gulf of Mexico to highlight the ecological importance of deep-sea organisms as both predators to zooplankton assemblages and prey to tunas and billfishes. Matt is also investigating the role vertically migrating fishes (e.g., lanternfish) have in carbon export/sequestration in the open ocean and the vertical movement of nutrients by deep-diving cetaceans.

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 1000th 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.