This article discusses the species of introduced herpetofauna (amphibians and reptiles) of Florida’s terrestrial and aquatic habitats along with a general discussion of the possible effects of biological invasions on native wildlife and habitats.  The first part of this three-part series was on introduced fishes in the state.  The final part of the series will be on introduced mollusks (bivalves and gastropods, or clams and snails & slugs) of Florida.  

As of this writing, at least 110 species of nonindigenous herpetofauna (colloquially called ‘herptiles’ for short) representing 34 families have been introduced to Florida (Exhibits 1 and 4).  Of the species introduced to Florida, about 43% are now considered to have established breeding populations in one or more counties (Exhibit 2).  This amounts to 47 established herptile species in Florida as of this writing.  Both urban and natural areas of Florida are affected by these biological invaders.  For example, the first reticulated python (Python reticulatus) observed in Florida was during the 1980s, where it was seen living under a house in Miami.  This species has since been observed and (or) collected in several other areas of Florida, although it is not known whether the species has established self-sustaining breeding populations (Exhibit 3). Lizards are the most successful group and account for the majority (72%) of established herptiles in Florida today.  The list in Exhibit 4 below contains the species known to have been introduced, although it is important to note that new species are introduced on a regular basis in Florida, so the list is constantly expanding.  Most introduced herptiles are native to the tropics (Wilson and Porras 1983).  The fact that Florida’s climate is subtropical is a major reason why many introduced species have successfully established themselves in the state.  Nonindigenous herptiles have been introduced via a variety of mechanisms:

• Stowaways in shipments of ornamental plants or produce
• Intentional or accidental release by pet dealers or owners
• Intentional or accidental release from zoological parks
• Intentional release by government agencies to combat nuisance organisms

 Wilson and Porras predicted in the early 1980s that southern Florida would eventually be overrun with introduced exotic wildlife.  The current trends in established and spreading introduced species suggest that these authors may have been right. 

Reducing the effects of invasive nonindigenous species is an important part of restoration and management efforts in natural areas of Florida, United States, and worldwide, as these species cause significant stress to native ecosystems (Adams and Steigerwalt 2010) and biological invasion is widely viewed as a major cause of the reduction in native plant and animal diversity (Elton 1958, Wilcove et al. 1998).  Invasive species are known to affect most natural areas of the United States (Villazon 2009) and worldwide (Sala et al. 2000).

It should go without saying that the intentional introduction of any nonindigenous species, whether it be a plant or animal and regardless of size or assumed innocuousness, should never be attempted.  The reasons are many and the costs can be severe, both in terms of biological effects and economic impacts.  Nonindigenous species introduced to new areas have the capacity to explode in numbers and outcompete native species for limited resources such as food, water, and shelter.  Native species are at a competitive disadvantage because they have not had time to evolve defense mechanisms that would otherwise allow them to successfully compete against the introduced species.  The competition between native and nonindigenous species can result in the extinction of native species, the spread of diseases and parasites, displacement of whole communities, and may even cause physical changes to the environment.  

Exhibit 4.  Nonindigenous amphibians and reptiles recorded in Florida.

Sources: Florida Museum of Natural History (http://www.flmnh.ufl.edu/herpetology/florida-amphibians-reptiles/checklist-atlas/), USGS Nonindigenous Aquatic Species online database (http://nas.er.usgs.gov/queries/SpeciesList.aspx?group=Amphibians&state=FL&Sortby=1), Krysko et al. (2011), J.C.Seitz unpublished data.

 Sources:

Adams, C.R. and N.M. Steigerwalt.  2010.  Research Needs and Logistic Impediments in Restoration, Enhancement, and Management Projects: A Survey of Land Managers. Publication ENH1161 [online resource].  Environmental Horticulture Department, Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, FL.  Accessed 11/21/10 at: http://edis.ifas.ufl.edu/ep423.

Elton, C.S.  1958.  The Ecology of Invasions by Animals and Plants.  Methuen and Co., Ltd., Strand, London.

Florida Museum of Natural History.  2014.  Checklist & Atlas of Amphibians and Reptiles in Florida [online resource].  Accessed 02/24/15 at http://www.flmnh.ufl.edu/herpetology/florida-amphibians-reptiles/checklist-atlas/.

Krysko, K.L., K.M. Enge, P.E. Moler.  2011.  Atlas of Amphibians and Reptiles in Florida.  Project Agreement 08013, report submitted to Florida Fish and Wildlife Conservation Commission, Tallahassee, FL.

Sala, O.E. F.S. Chapin, J.J. Armesto, E. Berlow, J. Bloomfield, R. Dirzo, E. Huber-Sanwald, L.F. Huenneke, R.B. Jackson, A. Kinzig, R. Leemans, D.M. Lodge, H.A. Mooney, M. Oesterheld, N.L. Poff, M.T. Sykes, B.H. Walker, M. Walker, and D.H. Wall.  2000.  Global biodiversity scenarios for the year 2100.  Science 287:1770–1774.

U.S. Geological Survey.  2015.  NAS – Nonindigenous Aquatic Species [online resource].  Accessed 03/03/15 at http://nas.er.usgs.gov/queries/CollectionInfo.aspx?SpeciesID=963&State=FL.

Villazon, K.A.  2009.  Methods to Restore Native Plant Communities after Invasive Species Removal: Marl Prairie Ponds and an Abandoned Phosphate Mine in Florida.  MS thesis, University of Florida, Gainesville, FL.

Wilcove, D.S., D. Rothstein, J. Dubow, A. Phillips, and E. Losos.  1998.  Quantifying threats to imperiled species in the United States. Bioscience 48:607–615.

Wilson, L.D. and L. Porras.  1983.  The Ecological Impact of Man on the South Florida Herpetofauna.  The University of Kansas Museum of Natural History Special Publication No. 9, University of Kansas, Lawrence, KS.

This article discusses the species of introduced fishes in Florida’s freshwater and marine habitats, along with a general discussion of biological invasions as a potential driver of loss-of-habitat functions.  Future articles in the series will discuss introduced mollusks (bivalves and gastropods) and herptiles (amphibians and reptiles) of Florida.

Waterbodies such as streams, lakes, ponds, and oceans are well known for their habitat functions, especially their ability to support aquatic wildlife by providing sustenance and shelter.  A myriad of animals, from tiny arthropods to 12-meter-long whale sharks, rely on native organisms as food.   Many waterbodies support some of the most productive habitats in the world, providing food and shelter for mollusks, crustaceans, fishes, amphibians, reptiles, birds, and mammals, and often serve as vital nursery grounds for these species.  Others are nutrient-poor and relatively unproductive.  Nevertheless, hundreds of imperiled species require aquatic habitats for survival.  Along with their threatened or endangered wildlife, waterbodies themselves are threatened in many ways.  Anthropogenic disturbances include groundwater depletion, reallocation of surface water, nutrient inputs, habitat fragmentation, fire suppression, pollution, land use changes, overharvesting, climate change, dredging, and the introduction of nonindigenous plants and animals (Exhibit 1).

Reducing the effects of invasive nonindigenous species is an important part of restoration and management efforts in natural areas of Florida, the United States, and worldwide.  These species cause significant stress to native ecosystems (Adams and Steigerwalt 2010), and biological invasion is widely viewed as a major cause of the reduction in native plant and animal diversity (Elton 1958, Wilcove et al. 1998).  Invasive species are known to affect most natural areas of the United States (Villazon 2009) and worldwide (Sala et al. 2000), and aquatic habitats are particularly susceptible to nonindigenous species due in part to the fact that aquatic habitats act as biological sinks, receiving plant and animal genetic material from upstream sources.

As of this writing, at least 192 species of fishes representing 42 families have been introduced to Florida (Exhibit 2).  Nearly all waterbodies are affected by fish introductions, from small wetlands to the Atlantic and Gulf coasts of Florida.  The list below contains the species known to have been introduced, although it is important to note that new species are introduced on a regular basis in Florida, so the list is constantly expanding.  Many species ultimately fail to gain a foothold in Florida, while a smaller number of species successfully establish themselves.  Some have spread like a cancer across the state.  The Brown Hoplo (Hoplosternum littorale) is an example of an introduction that is now established throughout the peninsula of Florida, much to the detriment of native aquatic species that have not had time to adapt to this new competitor for limited resources.  Marine habitats are not immune to biological invasions.  The detrimental effects of the (likely intentional) introduction of two species of invasive lionfishes (Red Lionfish and Devil Firefish [Pterois volitans and P. miles]) are still being determined but likely include direct predation on native fishes, crabs, and shrimps and competition with native reef species for limited resources.  Red Lionfish and Devil Firefish are now firmly established throughout the Atlantic coast of Florida and are actively invading much of the Gulf of Mexico.  The spread of lionfishes throughout the western North Atlantic Ocean is occurring at an unprecedented rate (see Exhibit 3) (Schofield 2010).  Many of the introduced fishes in Florida are from tropical or subtropical areas of Asia and South America, and to a lesser extent, Africa (Idelberger et al. 2010).  The fact that Florida’s climate is also subtropical is a major reason why many introduced species have successfully established themselves in the state. 

It should go without saying that the intentional introduction of any nonindigenous species, whether it be a plant or animal and regardless of size or assumed innocuousness, should never be attempted.  The reasons are many and the costs can be severe, both in terms of biological effects and economic impacts.  Nonindigenous species introduced to new areas have the capacity to explode in numbers and outcompete native species for limited resources such as food, water, and shelter.  Native species are at a competitive disadvantage because they have not had time to evolve defense mechanisms that would otherwise allow them to successfully compete against the introduced species.  The competition between native and nonindigenous species can result in the extinction of native species, the spread of diseases and parasites, and the displacement of whole communities, and may even cause physical changes to the environment. 

Exhibit 2.  Freshwater and marine nonindigenous fishes recorded from Florida.

Sources: Schofield (2010), USGS Nonindigenous Aquatic Species online database (http://nas.er.usgs.gov/queries/SpeciesList.aspx?group=Fishes&state=FL&Sortby=1)

Below is a link to an interactive map showing the spread of the Red Lionfish and the Devil Firefish in the western North Atlantic from the 1980s to 2013:

http://nas.er.usgs.gov/queries/FactSheets/LionfishAnimation.aspx

Sources:

Adams, C.R. and N.M. Steigerwalt.  2010.  Research Needs and Logistic Impediments in Restoration, Enhancement, and Management Projects: A Survey of Land Managers. Publication ENH1161 [online resource]. Environmental Horticulture Department, Florida Cooperative Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, FL. Accessed 11/21/10 at: http://edis.ifas.ufl.edu/ep423.

Didham, R.H., J.M. Tylianakis, M.A. Hutchison, R.M. Ewers, and N.J. Gemmell. 2005. Are invasive species the drivers of ecological change? Trends in Ecology and Evolution 20(9):470–474.

Elton, C.S. 1958. The Ecology of Invasions by Animals and Plants. Methuen and Co., Ltd., Strand, London.

Idelberger, C.F., C.J. Stafford, and S.E. Erickson.  2011.  Distribution and abundance of introduced fishes in Florida’s Charlotte Harbor estuary.  Gulf and Caribbean Research 23:13–22.

Sala, O.E. F.S. Chapin, J.J. Armesto, E. Berlow, J. Bloomfield, R. Dirzo, E. Huber-Sanwald, L.F. Huenneke, R.B. Jackson, A. Kinzig, R. Leemans, D.M. Lodge, H.A. Mooney, M. Oesterheld, N.L. Poff, M.T. Sykes, B.H. Walker, M. Walker, and D.H. Wall. 2000. Global biodiversity scenarios for the year 2100. Science 287:1770–1774.

Schofield, P.J.  2010. Update on geographic spread of invasive lionfishes (Pterois volitans [Linnaeus, 1758] and P. miles [Bennett, 1928]) in the western North Atlantic Ocean, Caribbean Sea and Gulf of Mexico. Aquatic Invasions 5, Supplement 1:S117–S122.  http://www.aquaticinvasions.net/2010/Supplement/AI_2010_5_S1_Schofield

U.S. Geological Survey.  2015. NAS – Nonindigenous Aquatic Species [online resource].  Accessed 01/23/15 at http://nas.er.usgs.gov/queries/CollectionInfo.aspx?SpeciesID=963&State=FL.

Villazon, K.A. 2009. Methods to Restore Native Plant Communities after Invasive Species Removal: Marl Prairie Ponds and an Abandoned Phosphate Mine in Florida. MS thesis, University of Florida, Gainesville, FL.

Vitousek, P.M., C.M. D’Antonio, L.L. Loope, and R. Westbrooks. 1996. Biological invasions as global environmental change. American Scientist 84:468–478.

Vitousek, P.M., H.A. Mooney, J. Lubchenco, and J.M. Melillo. 1997. Human domination of Earth’s ecosystem. Science 277:494–499.

Wilcove, D.S., D. Rothstein, J. Dubow, A. Phillips, and E. Losos.  1998.  Quantifying threats to imperiled species in the United States. Bioscience 48:607–615.

Recently, there has been considerable interest and research regarding the laurel wilt disease, which affects members of the Lauraceae family, most notably red bay (Persea borbonia) and swamp bay (Persea palustris).  This article attempts to summarize the aspects of this disease that are of particular interest to land owners and land managers of Florida and elsewhere in the southeastern United States.

The Story of the Ambrosia Beetle, a Symbiotic Fungus, and the Disease Called Laurel Wilt

The disease Laurel wilt is spread by a nonindigenous beetle called the Asian red bay ambrosia beetle, Xyleborus glabratus.  This beetle measures only about 2 mm in length and is cigar-shaped and amber-brown to black in color.  This species has significantly less hair on its dorsal surface and is shinier than other species of ambrosia beetles.  The female ambrosia beetle spreads a nonindigenous fungus, Raffaelea lauricola, into the sapwood of a tree by boring pinhole-sized holes into the branches or trunk and either actively or passively depositing spores of the fungus in the tunnels.  The fungal spores are carried by the beetle in specialized pouch-like structures called ‘mycangia’ that are located at the base of each mandible.  Both adult and larval ambrosia beetles feed on the fungus growing in the tunnels.  Larvae are white with an amber-colored head.  Unlike most other species of ambrosia beetle, which attack dead or dying trees, the Asian red bay ambrosia beetle attacks healthy trees.  The native range of the fungus includes India, Japan, Taiwan, Burma, Bangladesh, and Myanmar.  As you probably guessed, the Asian red bay ambrosia beetle is native to Asia, including the same countries that the fungus is native to. 

The exact mechanism that causes death to a tree infected with the fungus and symbiont ambrosia beetle to die is unknown.  In simplified terms, the death of the tree is the result of it over-reacting to the presence of the pathogen.

The ambrosia beetle and associated fungus are thought to have arrived in the United States from Asia in untreated wood (such as wooden pallets) or in logs.  They were first detected in the United States in Port Wentworth near Savannah, Georgia, in May 2002.  The disease has since spread throughout the Southeast, from North Carolina south to Florida and west to Mississippi.

Relative Infestations of Laurel Wilt Disease among Infected States

What Tree Species Does Laurel Wilt Infect?

Although the beetle is named Asian red bay ambrosia beetle, it actually infects several other species, including both native trees and introduced trees of importance to the agricultural and ornamental plant industries.  Below is a list of species known to be susceptible to Laurel wilt.

Trees and Shrubs Known or Suspected to be Susceptible to Laurel Wilt Disease

Summary of the Biology and Symptoms of Laurel Wilt Disease

The female ambrosia beetle, attracted to the smell of a red bay tree, bores into the branches or trunk of the tree and deposits spores of the fungus in the tunnels.  Initial symptoms are wilting of the leaves.  Often, wilting is seen in all the leaves associated with the distal portion of an infected branch.  More and more leaves begin to wilt over time as the disease progresses.  Mild discoloration may be seen in the sapwood and can escalate to extensive black/brown streaking over time.  Frass tubes, looking like white bent straws sticking out from the bark, begin to appear months later as beetle activity increases.  It can take as little as a week for a tree to die from laurel wilt during the warm summer months.

Both adult ambrosia beetles and their larvae feed on the fungus growing in the tunnels.  It takes some 30 days from the time the eggs hatch to the development of adult ambrosia beetles.  Males are smaller than females, lack wings, and are haploid.  Females are winged and are diploid.  The fungus can remain alive inside a standing dead tree for at least 1 year according to recent research.  The biology of laurel wilt disease remains poorly understood, and there is significant research to be done to understand the mechanisms involved in susceptibility and resistance.

Management and Prevention of Laurel Wilt Disease

It is no longer logistically feasible to eradicate or stop the progression of the disease considering how widely distributed it is in the southeastern United States.  However, one way to slow the spread on a given site is to cut down and chip dead trees killed by the disease and place the wood chips into piles.  The fungus was found to die about 2 days following chipping, and the ambrosia beetle population of the tree was found to be reduced by 99% following chipping.  The chipping will also reduce the wood available for female beetles to reproduce.

Use of the fungicide propiconazole (Alamo^®), injected into the tree, was found to be only mildly effective (approximately 60% survivorship) at protecting red bay trees from the disease.  This treatment is expensive and testing for use against Laurel wilt has been limited so far.  Best results are achieved by systemic injection before any symptoms of the disease are observed on the tree and the pruning of any diseased areas following treatment.  Another method of injecting fungicide, developed by Arborjet^®, involves delivering smaller amounts of fungicide using microinjectors.  The results of the effectiveness of the Arborjet^® method have not been published as of this writing.  Similarly, the results of the effectiveness of applying fungicide to the soil around a tree have yet to be published.  Fungicides should be administered only by a knowledgeable professional or by the homeowner and in accordance with the instructions and mixing rates on the label.

Insecticides are unlikely to be useful at protecting a tree again the ambrosia beetle.  Broadcast spraying would be harmful to the environment and to beneficial insects, is not likely to be effective against the ambrosia beetle, and is therefore strongly discouraged. 

An attempt was made to protect some trees in Volusia County, Florida, by spraying Pinesol^® as a way of “hiding” the trees from detection by the ambrosia beetle.  Pinesol^® spraying took place at about 6‑ to 10‑week intervals.  However, all treated trees eventually contracted the disease and subsequently died.  It is possible that baits may be developed in the future that may be more attractive to the beetles than are the trees, but at this point in time no compounds have been identified for use as baits.

Anyone can help reduce the spread of the ambrosia beetle and the associated Laurel Wilt disease.  Refrain from moving untreated firewood far distances.  The State of Florida prohibits movement of untreated firewood farther than 50 miles within the state.  When camping, buy only local firewood or use certified firewood rather than bringing your own.  When traveling abroad, do not bring back untreated wood products or raw plant parts (including seeds or fruits).

Laurel wilt disease is one of at least a dozen tree diseases and insect pests within Florida or neighboring states.  Minimizing the movement of untreated wood and firewood can help reduce the spread of insect pests and diseases such as the emerald ash borer (kills ash trees), Asian longhorned beetle (kills maples), oak wilt and bot canker of oaks (kills oaks), spiraling whitefly (kills several native and ornamental trees), walnut twig beetle and thousand-cankers disease (kills walnuts), sudden oak death (kills oaks), and others.  The reader is encouraged to visit the website www.dontmovefirewood.org for more information.  

Sources and Further Reading:

Global Invasive Species Database.  2010.  Global Invasive Species Database, Raffaelea lauricola (fungus) [online database].  Accessed 10/17/2014 at http://www.issg.org/database/species/ecology.asp?si=1549&lang=EN.

Global Invasive Species Database.  2010.  Global Invasive Species Database, Xyleborus glabratus (insect) [online database].  Accessed 10/17/2014 at http://www.issg.org/database/species/ecology.asp?si=1536.

Spence, D. and J. Smith.  2013.  The status of Laurel Wilt.  Palmetto 30(3):4–5, 8–10. 

U.S. Department of Agriculture, Forest Service.  2013.  Laurel Wilt Distribution Map [online resource].  Accessed 10/17/2014 at http://www.fs.fed.us/r8/foresthealth/laurelwilt/dist_map.shtml.

The jumping of Gulf sturgeon (Acipenser oxyrinchus desotoi) on the Suwannee River is well publicized and is widely known among Floridians. Some jumping sturgeon have even caused injuries to unfortunate boaters who get in the way of these armored aerial acrobats. Sturgeon jump most often in the early morning, and each fish averages one jump per day. Jumping is not confined to large individuals either, as fish as small as 1 foot long will jump. Most jumping activity occurs from June to early July, when some reaches of the river can experience as many as 6 to 10 jumps per minute! The most intense jumping occurs when the water level is low, before the summer convective storm season sets in. Yes, the fact that these massive timeless fish jump is not the question here. They jump. We know. What’s not well publicized is why exactly these sturgeon feel the need to jump in the first place.

Gulf sturgeon are not trying to shed ectoparasites as many believe, nor are they trying to capture prey. The jumping is also unrelated to spawning. They are not reacting to the presence of boats.

They loiter in the deep dark pools of the Suwannee River and in other Gulf Slope rivers from April through September and slowly digest the 20 percent of body fat they put on during the previous 6 months of easy living, and easy feeding, in the Gulf of Mexico. When in the Suwannee they seldom feed, so put that fishing pole away.

Resting in the sluggish deep recesses of the Suwannee, Gulf sturgeon are conserving energy. But just hovering above the river bottom requires neutral buoyancy. Similar to how a SCUBA diver adjusts his BC, these fish must add air to the swim bladder. Air slowly leaks out of the swim bladder and is absorbed in the surrounding tissues. To keep from scraping along the bottom and to swim more efficiently in the sluggish current, the swim bladder must be refilled an average of once daily. Since the swim bladder in Gulf sturgeon is connected to the digestive tract by a duct, the bladder can be recharged by gulping air. But how do fishes, being totally aquatic as they are, find any atmospheric air to gulp? Well, we have nearly reached our answer as to why they jump.

Any SCUBA diver knows that you can’t descend easily through the water column with a BC full of air, even with dive weights. By the same principle, a sturgeon can’t just rise gently to the surface, take a big gulp of air, and glide back down into the depths. Did I mention that while sturgeon are lollygagging in the river, they are also chatting up a storm? Call them gabby or chatty, Gulf sturgeon really know how to bend an ear (or otolith, in this case). These fish communicate with one another via a series of snapping sounds. Jumping high up into the air and then falling powerfully back into the water, with a loud report, and swimming directly to the bottom, serves two purposes. The main purpose is to allow the fish to gulp air and yet have enough momentum to reach the river bottom immediately afterward (despite the increased buoyancy from the added air). The secondary purpose may be to communicate. Immediately prior to jumping, these fish emit a series of snapping sounds, and following the belly-flop-style smack when entering the water, the sturgeon emit more snapping sounds. The role jumping plays in sturgeon communication, if any, is far from clear and would be very difficult to prove scientifically. However, it seems probable that these fish are communicating with one another using not just snapping sounds (which have been recorded by U.S. Geological Survey scientists and heard by SCUBA divers), but perhaps also by the splash following a jump. So, the next time you encounter a jumping sturgeon while enjoying a lazy day on the Suwannee, ask it why it jumps. Listen carefully to its answer and don’t be surprised if its response sounds something like “snap, snap-snap, snaaap, snaaap, snap”.

Source and Suggested Reading:

Sulak, K. 2013. Catching air – those magnificent jumping Suwannee sturgeons. American Currents 38(2):23–25.

During a benthic survey off the Coquille River, Oregon, in September 2013, ANAMAR was collecting samples of epifauna using a 12-foot otter trawl when suddenly the gear encountered unidentified rock.  The trawl net snagged and the cable instantly snapped, losing the gear on the seafloor in about 45 feet of water.  Although many attempts were made to recover the trawl using a grapple hook off the deck of the survey vessel (R/V Pacific Storm), the gear was too entangled on the seafloor to be brought up with that method.  Directly following completion of the benthic survey, an ANAMAR subcontractor returned to the site and recovered the trawl gear using SCUBA divers.  The trawl was still in good shape and the remaining trawl tows were completed for the survey.  In addition to finding the trawl gear, the divers also observed several fossil scallop shells embedded in the rock on the seafloor.  The fossil scallops were in excellent condition (see images below).  The divers were able to pry a few of the fossil shells loose for closer inspection and photography.

Because the area where the survey took place is an ocean dredged material disposal site (ODMDS), information on the naturally occurring rocks found there is of interest to agencies tasked with managing the site (U.S. Army Corps of Engineers and the U.S. Environmental Protection Agency).  For this reason, and also out of personal interest, I began collaborating with paleontologists to determine the identity of the fossil scallops in the hopes of learning more about the rock they were found in.  I soon found my answer after contacting specialists at the Burke Museum of Natural History in Seattle, Washington.  Dr. Elizabeth Nesbitt, Curator of Paleontology, graciously identified the fossil scallops as either Patinopecten coosensis or P. oregonensis based on photos I sent her.  The flared portions of the shell adjacent to the hinge (called auricles) serve as key characteristics differentiating these two species.  These fossils lacked auricles so they could not be identified beyond these two species.  However, based on the fossils and the associated matrix, Dr. Nesbitt was able to identify the rock formation the fossils were found in!

The rocks and fossils are part of the Empire Formation which is better known from exposures about 20 miles south of the Coquille River, at Cape Blanco, Oregon.  The Empire Formation, composed mostly of sandstone, along with the fossils it contains, are as old as 12 million years (Miocene) but it is theorized to be closer to 8 to 5 million years (Miocene-Pliocene epoch boundary).  Since we know the identity of the rock as being part of the Empire Formation, we therefore know something about its composition.  In this case, the rocks that snagged the trawl gear must have been composed of sandstone and some siltstone.  This formation represents sands deposited in what was then a small marine basin, which now is only represented by Coos Bay.  It is probable that other rocks within the ODMDS are also fossiliferous sandstone/siltstone from the Empire Formation.

The above is an example of how fossils can help us infer the identity of the surrounding substrate.  In this case, the identity of the fossil scallops, along with the matrix attached to the fossils, were used to pinpoint the exact formation they represent.  Knowing the formation, we then were able to learn more about the composition and approximate geological age of surrounding rocks that represent the same formation.  All this information came from observing and collecting a handful of fossils incidental to recovering of some equipment from the seafloor!

Interestingly, the French word for scallop is Coquille.  Thus, the Coquille River, where the fossils were collected, was actually named after a scallop!

Sources:

Ehlen, J.  1967.  Geology of state parks near Cape Arago, Coos County, Oregon.  The Ore Bin 29(4):61–82. 

Nesbitt, E.  Department of Paleontology, Burke Museum of Natural History, University of Washington, Seattle, WA.  Pers. comm. 12/06/13.

Portell, R.W.  Department of Invertebrate Paleontology, Florida Museum of Natural History, University of Florida, Gainesville, FL.  Pers. comm. 11/18/13.

All sawfishes are highly modified and elongate rays that swim like a shark and have a long snout with laterally-placed spines. The snout (called a ‘rostrum’) is actually an extension of the skull (known as the ‘chondrocranium’) and the lateral spines are called ‘rostral teeth’ by scientists. Like the rest of the skeleton of the sawfish, the rostrum is composed of cartilage, albeit reinforced with extra calcium. The rostrum and rostral teeth are used in food gathering. The sawfish uses the rostrum to stun prey, such as fishes and invertebrates, which it then sucks into its mouth positioned under the head. There is no cutting or tearing and sawfish can only consume fish and invertebrates that fit into the mouth whole.

The modern sawfish group (family Pristidae) first showed up in the fossil record between the beginning of the Cenozoic (about 66 million years ago) and the beginning of the Eocene (about 56 million years ago). The only exception is the nominal genus Peyeria, which first appeared during the upper Cretaceous (about 100 million years ago). However, the fossil material attributed to the genus Peyeria may actually represent another type of ray—a member of the sharkfin guitarfish family (Rhinidae) such as the bowmouth guitarfish (Rhina ancylostoma).

Living (extant) sawfishes include the following species:

The Knifetooth complex (one species):

• Anoxypristis cuspidata (knifetooth sawfish [western Pacific and Indian oceans])

The Smalltooth complex (three species):

• Pristis clavata (dwarf sawfish [western Pacific Ocean])
• Pristis pectinata (smalltooth sawfish [eastern and western Atlantic Ocean])
• Pristis zijsron (green sawfish [western Pacific and Indian oceans])

The Largetooth complex of sawfish has recently been combined into one species:

• Pristis pristis (largetooth sawfish [eastern and western Atlantic, eastern and western Pacific, and Indian oceans])

There were also several additional members of the modern sawfish group that are only represented as fossils:

• Anoxypristis mucrodens (fossils of Europe, North America, West and East Africa)
• Peyeria libyca (nominal species; fossils of northeast Africa including Egypt)
• Propristis schweinfurthi (fossils of North and West Africa)
• Pristis spp. (at least eight extinct species described; fossils of West and East Africa, Europe, and North America)

Prior to the modern sawfishes, a diverse group of sawfishes (family Sclerorhynchidae) lived during the Cretaceous epoch. Members of the Cretaceous sawfishes had a diverse array of rostral tooth morphologies, ranging from closely-spaced thin spines to widely-spaced massive barbed teeth on sturdy, widened bases. Although most species reached a modest size of not more than 1 meter in total length, fossil rostra measuring well over 1 meter in length have been unearthed in Cretaceous sediments of Morocco.

There are at least 20 genera of Cretaceous sawfishes, including the following:

• Ankistrorhynchus (fossils of Europe and North America)
• Atlanticopristis (fossils of South America)
• Baharipristis (fossils of East Africa)
• Biropristis (fossils of South America)
• Borodinopristis (a nominal genus of about three species [fossils of North America])
• Ctenopristis (fossils of West and East Africa)
• Dalpiaza (fossils of West Africa)
• Ganopristis (fossils of Europe)
• Ischyrhiza (fossils of North and South America)
• Kiestus (fossils of North America)
• Libanopristis (fossils of Middle East)
• Marckgrafia (fossils of West Africa)
• Micropristis (fossils of Europe and Middle East)
• Onchopristis (fossils of North America, Europe, West and East Africa, Asia, New Zealand)
• Onchosaurus (fossils of North America and West Africa)
• Plicatopristis (fossils of the Middle East)
• Pucapristis (fossils of South America)
• Renpetia (fossils of the Middle East)
• Schizorhiza (fossils of North America and Middle East)
• Sclerorhynchus (fossils of North America and Middle East)

Modern sawfishes and Cretaceous sawfishes evolved independently from the guitarfishes (Rhinobatidae). Thus, while both groups are commonly referred to as sawfish, and both share similar morphological characteristics, they are not closely related. Sawfishes should not be confused with the saw sharks (family Pristiophoridae), which are true sharks and therefore only distantly related to the sawfishes (which are rays, not sharks).

The rostral teeth of Cretaceous sawfishes are attached to the dermis of the rostrum via connective tissue. Their rostral teeth are thought to be continually replaced throughout the life of the animal in the same conveyer-belt fashion as are the oral teeth of all sharks and rays. In contrast, modern sawfishes have rostral teeth firmly embedded in sockets (called ‘alveoli’) and their teeth are not replaced if lost. Cretaceous sawfish rostral teeth are covered with an enamel-like coating along the cusp; this coating is lacking in modern sawfishes. Cretaceous sawfishes also differ from modern sawfishes in that they possess a long, whip-like caudal fin. Although the average fossil remains consist of isolated rostral teeth or oral teeth, some beautiful, fully articulated fossil skeletons have been unearthed in Lebanon quarries.

Sources:

Alroy, J. and M. McClennen. 2013. Paleobiology Database [online resource]. Accessed 06/10/13 online at http://www.paleodb.org/?a=displaySearchColls&type=view.

Faria, V.V., M.T. McDavitt, P. Charvet, T.R. Wiley, C.A. Simpfendorfer, and G.J.P. Naylor. 2013. Species delineation and global population structure of critically endangered sawfishes (Pristidae). Zoological Journal of the Linnean Society 167(1):136–164.

Seitz, J.C. 2013. Fossil Sawfish [online resource]. Accessed 06/10/13 online at http://www.fossilsawfish.com/.

Wueringer, B.E., L. Squire Jr., and S.P. Collin. 2009. The biology of extinct and extant sawfish (Batoidea: Sclerorhynchidae and Pristidae). Reviews in Fish Biology and Fisheries 19:445–464.

Authored by Jason C. Seitz of ANAMAR

Sawfishes are large highly modified and elongate rays that swim like a shark and have a long snout with laterally-placed spines.  The snout (called a ‘rostrum’) is actually an extension of the skull (known as a ‘chondrocranium’) and the lateral spines are called ‘rostral teeth’ by scientists.  Like the rest of the skeleton of the sawfish, the rostrum is composed of cartilage, albeit reinforced with extra calcium.  The rostrum and rostral teeth are used in food gathering.  The sawfish uses the rostrum to stun fishes and to a lesser extent, invertebrates, which it then sucks into its mouth positioned under the head.  There is no cutting or tearing and sawfish can only consume fish and invertebrates that fit into the mouth whole.  The longest species of sawfish, the green sawfish (Pristis zijsron), is reported to reach a length of 23.9 feet according to Last and Stevens (1994).

Sawfishes can be distinguished from saw sharks (Pristiophurus spp. and Pliotrema warreni) by the lack of barbels, ventrally located gills (versus laterally located), dorso-laterally compressed body, and uniformly sized rostral teeth.  Sawfishes grow much larger than do saw sharks.  Further, sawfishes prefer warm coastal waters while saw sharks inhabit deeper cooler offshore waters.

Living sawfish species are globally distributed in tropical and sub-tropical coastal marine and estuarine waters, and sometimes inhabit rivers and associated freshwater bodies such as Lake Nicaragua.  The center of distribution is the western Pacific including northern Australia and Papua New Guinea. 

All modern sawfish species are considered imperiled and regarded by the International Union for the Conservation of Nature (IUCN) as ‘critically endangered’ with declining populations (www.iucnredlist.org).  Unfortunately, the IUCN is not a regulatory agency and thus no protection is afforded by this group.  Only two species currently have protection under the endangered species act of 1972, and only in waters of the United States.  The Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES) has protected all species of sawfish from international trade of sawfish parts since 2007 (www.cites.org).  However, the confusing taxonomy of modern sawfishes has hampered conservation efforts and such efforts are further disadvantaged by the poorly known geographical population structure.

Until very recently, there were six valid living species worldwide. The Knifetooth Complex of sawfish currently consists of one species:

Anoxypristis cuspidata (knifetooth sawfish [western Pacific and Indian oceans])

 The Smalltooth Complex of sawfish consists of three species:

Pristis clavata (dwarf sawfish [western Pacific Ocean])

Pristis pectinata (smalltooth sawfish [eastern and western Atlantic Ocean])

Pristis zijsron (green sawfish [western Pacific and Indian oceans])

 The Largetooth Complex of sawfish consisted of two species:

Pristis microdon (freshwater sawfish [western Pacific and Indian oceans])

Pristis perotteti (largetooth sawfish [eastern and western Atlantic Ocean, eastern Pacific Ocean])

Some researchers also considered the species Pristis pristis to be a valid member of the Largetooth Complex, although most considered it as an invalid synonym due to problems relating to its original description and lack of voucher specimens in museums (Faria 2007).  Also, the eastern Pacific population of the largetooth sawfish was considered to be a separate species (Pristis zephyreus) by some researchers following molecular phylogenetic work by Faria (2007), although no reliable morphological differences have been found between the eastern Pacific population and largetooth sawfish from the western Atlantic.

The freshwater sawfish and the largetooth sawfish have been problematic for researchers because the two species cannot be reliably differentiated by morphology and thus, these species were differentiated solely by region.  For specimens lacking collection data, this presents a challenge as the species may not be reliably determined at all.  Molecular work has shown that these species group closely together in terms of genetic background (Naylor et al. 2012).

Because of the fact that the largetooth sawfish and the freshwater sawfish are indistinguishable by morphology and share similar genetic backgrounds, it has been very recently proposed by researchers that these two species should be combined.  In a paper published in early 2013 and authored by Faria et al. (2013), all the Largetooth Complex species have been combined into a composite species under the resurrected name of Pristis pristis.  The resurrection of the formerly invalid name of Pristis pristis was based on Rule 23.1 (principle of priority) of the International Code of Zoological Nomenclature, which states that ‘the valid name of a taxon is the oldest available name applied to it’ (International Commission on Zoological Nomenclature 1999).  Since the species name Pristis pristis was published in 1758 (as Squalus pristis by Carl Linnaeus), it precedes the naming of all other Largetooth Complex species.  For this reason and because this name was used as a valid species after 1899, Pristis pristis is proposed by Faria et al. (2013) for use in place of Pristis microdon, Pristis perotteti, and the species of questionable validity, Pristis zephyreus.

Based on Faria et al. (2013), there are now a total of five valid sawfish species:

The Knifetooth complex (one species):

Anoxypristis cuspidata (knifetooth sawfish [western Pacific and Indian oceans])

The Smalltooth complex (three species):

Pristis clavata (dwarf sawfish [western Pacific Ocean])

Pristis pectinata (smalltooth sawfish [eastern and western Atlantic Ocean])

Pristis zijsron (green sawfish [western Pacific and Indian oceans])

The Largetooth complex of sawfish now consists of only one species:

Pristis pristis (largetooth sawfish [eastern and western Atlantic, eastern and western Pacific, and Indian oceans])

It is possible that in the future, changes may occur within the taxonomy of the knifetooth sawfish.  Gene flow between Indian Ocean and western Pacific specimens was found to be very low in a study by Faria et al. (2013).  Specimens from the Indian Ocean were found to have a higher average number of rostral teeth per side (average of 25.6) versus western Pacific specimens (average of 21.2) (faria et al. 2013).  Results of the Faria et al. (2013) study suggest that the knifetooth sawfish may actually represent multiple species.  In fact, a DNA-sequencing based analysis by Naylor et al. (2012) showed that knifetooth sawfish had considerable genetic differences from all other living members of the sawfish family.  This suggests that in the future the knifetooth sawfish may even be placed into a separate family distinct from the other sawfishes.

It is clear that there is still a lot to learn about sawfishes.  Although it is uncertain what changes will occur in sawfish taxonomy, or what new information will come to light from future scientific research, it remains clear that this interesting group of animals will continue to captivate scientists and layman alike for many years to come.

Sources Cited: Faria, V.V.  2007.  Taxonomic Review, Phylogeny, and Geographical Population Structure of the Sawfishes (Chondrichthyes, Pristiformes).  PhD dissertation, Iowa State University, Ames, IA.

Faria, V.V., M.T. McDavitt, P. Charvet, T.R. Wiley, C.A. Simpfendorfer, and G.J.P. Naylor.  2013. Species delineation and global population structure of critically endangered sawfishes (Pristidae).  Zoological Journal of the Linnean Society 167(1):136–164.

International Commission on Zoological Nomenclature.  1999.  International Code of Zoological Nomenclature. The International Trust for Zoological Nomenclature, The Natural History Museum, London, UK.

Last, P.R. and J.D. Stevens.  1994. Sharks and Rays of Australia.  CSIRO Division of Fisheries, Victoria, Australia.

Naylor, G.J.P., J.N. Caira, K. Jensen, K.A.M., Rosana, W.T. White, and P.R. Last.  2012. A DNA sequence-based approach to the identification of shark and ray species and its implications for global elasmobranch diversity and parasitology. Bulletin of the American Museum of Natural History 2012(367):1–262.

Harbor sediment is often subjected to physical, chemical, toxicological, and bioaccumulation analyses as part of dredging projects that keep our harbors navigable; however, the fossils in the sediment often go unnoticed and unappreciated during such testing.  What can the fossil remains—the critters that once inhabited the area—tell us about the age and history of the sediment?  An ANAMAR biologist set out to learn just that, by sieving about 5 gallons of extra material from a recent sediment sampling event at Charleston Harbor, South Carolina.

The sample was collected with a vibracore sampler at the area of Mount Pleasant Range and Benice Reach.  The matrix was a mixture of shells, sand, silt, and clay.  The particles smaller than the 2-mm mesh size screen were washed away during the sieving process.  The remaining shells and other hard objects were left to dry over a few week’s time.  The remains were then identified and photographed. 

So what was found?

The sample contained:

• Crustacean remains
  • Crab and shrimp claws
  • Barnacle shells
• Echinoid (sea urchin) spines
• Mollusk shells
  • Ark shells of the order Arcoida
  • Limpets of the genus Diodora sp.
  • Many other gastropod and bivalve shells
• Shark teeth
  • Hemipristis serra (extinct snaggletooth shark)
  • Physogaleus (Galeocerdo) contortus (extinct tiger shark)
  • Galeocerdo cuvier (modern [extant] tiger shark)
  • Hundreds of small teeth from other carcharhinids (requiem sharks)
• Shark vertebrae
• Ray teeth
  • Dasyatis sp. or Himantura sp. (stingrays)
  • Rhinopteridae and/or Myliobatidae (cownose rays, eagle rays)
  • cf. Paramobula fragilis and cf. Plinthicus stenodon (extinct devil rays)
• Bony fish remains:
  • Otoliths (ear bones)
  • Teeth from Sphyraena sp. (barracuda)
• Undetermined bone fragments
• Apatite (a phosphate-containing mineral)

Estimating the age of the sediment based on fossil remains

The snaggletooth shark Hemipristis serra occurred during the Pleistocene (12,000 to 2.6 million years ago) to the Eocene (34 million to 56 million years ago) and its fossil remains indicate warm, well-oxygenated marine conditions back then (Cappetta 1987).  The extinct tiger shark Physogaleus (Galeocerdo) contortus occurred during the Miocene (5 million to 23 million years ago) (possibly also during the Eocene) and is a characteristic fossil of the Southeastern Coastal Plain (Cappetta 1987).  The devil ray teeth were damaged but appeared to resemble the extinct species Paramobula fragilis of the Oligocene epoch (23 to 34 million years ago) and Plinthicus stenodon of the Miocene (Cappetta 1987, www.elasmo.com).

The tiger shark Galeocerdo cuvier and many other requiem shark species are still around today.  The stingray genera Dasyatis and Himantura, the eagle rays, and the cownose rays are also still around today, although the stingray genus Himantura is now absent from the western North Atlantic.  Similarly, the barracuda genus Sphyraena is still around today including four modern species inhabiting the western North Atlantic (Robins and Ray 1986).

Identification of the invertebrate remains was not yet attempted to the lowest practical taxonomic level, but they appeared to range in age from the Holocene (recent) to the Pleistocene (12,000 to 2.6 million years ago) based on initial observations. 

Based on the shark and ray fauna represented by fossils, the sediment taken from Charleston Harbor probably dates back to the Miocene.  Given that some fish and invertebrate remains may be modern, the sediment probably also includes relatively recently deposited material.  The fact that multiple geological epochs appear to be represented in the sediment is typical for fluvial (riverine) deposits such as in Charleston Harbor.  In addition to the fossil remains, the mineral apatite, containing phosphate, was found as smooth spherical black stones.  Mineral identification was made by W.G. Harris (pers. comm.).

Sources Cited:

Cappetta, H.  1987.  Handbook of Paleoichthyology, Chondricththyes II. Mesozoic and Cenozoic Elasmobranchii.  Gustav Fischer Verlag, Stuttgart, Germany.

Harris, W.G.  2013.  Professor of Soil Mineralogy, Soil and Water Science Department, University of Florida, Gainesville, FL. Pers. Comm.

Robins, C.R. and G.C. Ray.  1986.  A Field Guide to Atlantic Coast Fishes. North America.  Houghton Mifflin Co., New York, NY.

The movie Jaws, released June 20, 1975, was a box office blockbuster with a plot that revolved around a great white shark wreaking havoc around a fictional island called Amity off Long Island, New York.  Based on the 1974 novel of the same name by Peter Benchley, the film included impressive underwater video shot by diving and spear-fishing aficionados Ron and Valery Taylor.  The shark was filmed in waters off Australia and the video included a scale model of a dive cage, complete with a small-framed actor, which made the great white seem bigger by comparison. 

However, Jaws was not the first to feature great white sharks in their natural habitat.  The film Blue Water, White Death is a documentary of sharks in their natural habitat which included the first underwater video of great white sharks ever presented.  Released on June 1, 1971, Blue Water, White Death was released four years before the movie Jaws.  Similar to Jaws, Blue Water, White Death includes video shot by Ron and Valerie Taylor along with footage by underwater photographer Stan Waterman.  The film took a full 9 months to complete during which time the film crew, lead by filmmaker and photojournalist Peter Gimbel, traveled between the waters off South Africa and Australia in search of great white sharks.  While the video was made using technology of limited quality compared to today’s standards, the scenes in the film are spectacular for their time.  The adventures of the crew were also documented in the book Blue Meridian: the Search for the Great White Shark (1971) by award-winning writer Peter Matthiessen. 

Ladyfish (rarely called ‘tenpounders’) are economically valuable and are landed throughout the southeastern United States in both commercial and recreational fisheries (Levesque 2011).  Ladyfish are common in coastal areas throughout most of the western central Atlantic Ocean, including the Caribbean and Gulf of Mexico (Smith and Crabtree 2002).  The genus Elops in the western central Atlantic was traditionally treated as a single species, E. saurus.  This has changed recently with the description of a new species, E. smithi, by McBride and co-workers (2010).  Here’s how to tell the two species apart:

Counts of myomeres (in leptocephala larvae) or vertebrae (in post-larval to adult specimens) are the only known distinguishing morphological characters (McBride and Horodysky 2004, McBride et al. 2010).  E. saurus has 79–87 (usually 81–85) myomeres or vertebrae versus 73–80 (usually 75–78) in E. smithi.  (Number of myomeres = number of vertebrae).The number of myomeres or vertebrae appears to be a good diagnostic character, despite the overlap in range, as this overlap occurred in only 2.9% of the 3,255 specimens examined by McBride et al. (2010).  In counts of leptocephalus pre-anal myomeres or vertebrae, there is no overlap (see following table).

Myomeres should be counted using a compound microscope at 40x magnification, beginning with the first myomere behind the head and ending with the group of three myomeres near the caudal peduncle (McBride and Horodysky 2004).  Vertebrae (in post-larval to adult specimens) can be counted using radiographs and a microscope or by filleting, steaming, scraping, and directly counting the vertebrae.  Vertebral counts include all centra between the proatlas to the urostyle (McBride and Horodysky 2004). 

There are seasonal and geographic recruitment differences between the two species that can help you determine the species by date of capture: E. saurus larvae are most often found during winter through spring versus summer through fall in E. smithi (McBride and Horodysky 2004).  Generally speaking, Elops along the northern U.S. Atlantic seaboard are most often E. saurus while Elops from the Caribbean basin are most often E. smithi (McBride and Horodysky 2004).

The following table presents morphological, seasonal, and geographic comparisons between E. saurus and E. smithi that can help in species-level determinations.  Although it is admittedly difficult to discern the two species from one another, particularly when field-identification is needed, this write-up and associated table should make it more straightforward.

Sources Cited:

Levesque, J.C.  2011.  Is today’s fisheries research driven by the economic value of a species?  A case study using an updated review of ladyfish (Elops saurus) biology and ecology.  Reviews in Fisheries Science 19(2):137–149.

McBride, R.S. and A.Z. Horodysky.  2004.  Mechanisms maintaining sympatric distributions of two ladyfish (Elopidae: Elops) morphs in the Gulf of Mexico and the western North Atlantic Ocean.  Limnology and Oceanography  49(4):1173–1181.

McBride, R.S., C.R. Rocha, R. Ruiz-Carus, and B.W. Bowen.  2010.  A new species of ladyfish, of the genus Elops (Elopiformes: Elopidae), from the western Atlantic Ocean.  Zootaxa 2346:29–41.

Smith, D.G. and R. Crabtree.  2002.  Tenpounders (ladyfishes).  Pp. 679–680.  In:  Carpenter, K.E. (ed.), FAO Species Identification Guide for Fishery Purposes:  The Living Marine Resources of the Western Central Atlantic.  Vol. 2:  Bony Fishes Part 1 (Acipenseridae to Grammatidae).  FAO, Rome, Italy.

Papa Hemingway, as he preferred being called over Ernest, was an avid fisherman throughout his life.  He purchased his famous sport fishing boat, the 38-foot Pilar, in 1934 from a company in Brooklyn, New York, for $7,495.  Hemingway named the vessel after a nickname given to his then-current wife Pauline.  The Pilar came fitted with a flying bridge, a live well, and a special modification of the transom to allow large fish to be hauled into the vessel.  Hemingway fished in the Florida Keys and off Cuba and the Bahamas.  He was particularly fond of fishing the Gulf Stream, where he often targeted tuna and marlin. 

His well-honed techniques at successfully landing big tuna and marlin, coupled with his propensity to fish the Gulf Stream using heavy tackle and the many photos of him with landed heavyweights, suggest that Hemingway was most interested in large tuna and marlin.  Anyone having read The Old Man and the Sea probably would guess that Hemingway held a special spot in his heart for marlin.  While speaking of fishing, he often would mention wanting to land a “grander”, a term that refers to 1000-pound-class marlin.  Although the largest of the marlins—Indo-Pacific blue marlin—are legendary heavyweights often referred to as granders, Hemingway’s fishing exploits are outside of the range of that species.  Similarly, the massive black marlin is absent from Hemingway’s area of exploit.  Instead, Hemingway coveted the huge Atlantic blue marlin of the Gulf Current, and he targeted them in his Key West-Havana-Bimini fishing triangle.  Although the Atlantic blue marlin averages only between 300 and 400 pounds, the species’ maximum size of over 12 feet and robust body puts it well into the grander category.  The International Game Fish Association (IGFA) all-tackle angling record, a 1,282 pound monster, was landed off St. Thomas in the U.S. Virgin Islands.  Unfortunately, the weights of some of Hemingway’s biggest marlin remain unknown to this day due to the depredations of sharks while landing the marlin.

Of the tunas, Hemingway probably most eagerly sought the yellowfin, blackfin, bigeye, and Atlantic bluefin tunas, all of which can be caught in Hemingway’s Key West-Havana-Bimini fishing triangle.  Although all of these tunas reach impressive sizes worthy of the most skillful and resourceful of anglers, one species is perhaps the most coveted of all the tunas.  The Atlantic bluefin is often referred to simply as a “giant”.  With a maximum size of over 10 feet, coupled with a torpedo-shaped body made of pure muscle, the Atlantic bluefin may well have been Hemingway’s most prized fish.  The IGFA all-tackle record tipped the scales at about 1,497 pounds for a giant bluefin caught off Nova Scotia in 1979.

In celebration of Hemingway’s passion and aptitude for big game fishing, the Hemingway International Billfish Tournament is held each year near Havana, Cuba, where anglers target marlin, tuna, wahoo, dolphinfish, and other heavyweights over a several-day period.  A full-size replica of Hemingway’s Pilar, complete with several fighting chairs, can be seen on display at the Worldwide Sportsman store in Islamorada, Florida.  The original Pilar is on display in the Museo Ernest Hemingway in Cuba, near Havana.

Searobins are small to medium-sized benthic fishes (to 45 cm total length) that are common along Florida’s coastlines.  Although they are not targeted in commercial or recreational fisheries, searobins are captured in bottom trawls intended for shrimp or captured during epifaunal research surveys.  Searobins are important members of the benthic community and can amount to considerable biomass in some areas of Florida.  There are a total of 15 species within the family Triglidae in Florida waters.  First, one needs to determine which of the two genera his or her specimen belongs:

Bellator versus Prionotus

The genus Bellator can be distinguished from Prionotus by the absence of scales on the opercular membrane above the opercular spine.  There are usually 11 dorsal spines (rarely 10 or 12).  The first or second dorsal spines of males of Bellator species often end in long filaments (except in B. brachychir) (Richards and Miller 2002).  Also, species of the genus Bellator are relatively small (less than 17 cm standard length) (Richards and Miller 2002).  Finally, species of the genus Bellator are not normally found in inshore waters below about 20 m depth such as shallow bays, inlets, or estuaries.

In contrast to Bellator, the genus Prionotus never has a long, filamentous second dorsal spine.  Also, the dorsal spines usually number 10 (rarely 9 or 11) in Prionotus, and the opercular membrane is partially scaled above the opercular spine (Richards and Miller 2002).  Species of the genus Prionotus are often found in inshore waters, including bays and inlets.

See below for determining species within each genus.

Bellator Species

If you determined that your specimen falls within the genus Bellator, use the following table to help narrow down the possible species by geographic range or depth in Florida waters.

Bellator Morphological Characters by Species

The following characters are based on Richards and Miller (2002).

Bellator brachychir

• No filamentous dorsal spine present
• First free pectoral fin ray considerably longer than length of pectoral fins
• Usually less than 11 cm standard length (maximum size is 16 cm)

Bellator egretta

• Males with elongate filament extending from the first dorsal spine
• Head very spiny, including sharp spine in front of eye, and long opercular and preopercular spines
• First free pectoral fin rays shorter than length of pectoral fins
• Usually less than 10 cm standard length (maximum size is 15 cm)
• Alternating light and dark pigment appearing as brown patches or bands on upper one or two pectoral fin rays

Bellator militaris

• Males with elongate filaments extending from the first and second dorsal spines
• Dorsal-most pectoral fin rays marked with black and white bands
• No dark ventral margin on pectoral fin
• Colors in life are rosy with yellow lines along mid-body extending to caudal fin
• Usually less than 11 cm standard length (maximum size is 16 cm)

Prionotus Species

If you determined that your specimen falls within the genus Prionotus, use the following table to help narrow down the possible species by geographic range or depth in Florida waters.

Prionotus Morphological Characters by Species

The following characters are based primarily on Richards and Miller (2002), McEachran and Fechhelm (2005).

Prionotus alatus

• Lower pectoral fin rays very long, reaching past the posterior margin of anal fin
• Pectoral fins with black bands
• Small nasal spines present (can be detected by running finger along snout region towards snout tip)
• Maximum size is 20 cm standard length
• Reported to hybridize with P. paralatus between Gulfport and Panama City

Prionotus carolinus

• Branchiostegal rays dark (dusky or black)
• Pectoral fins attractively patterned throughout with spots in fresh specimens (see following figure)
• Dorsal fin with a single dark non-ocellated spot between 4^th and 5^th spines
• Can be distinguished from P. scitulus by the dark dorsal spine mentioned above
• Can be distinguished from P. martis by the dark dorsal spine and by range
• Caudal peduncle with white blotch on dorsal side
• Maximum size is 38 cm standard length

Prionotus evolans

• Nasal spines are absent
• Pectoral fins dark (patterned with very narrow, closely spaced, dark lines) (see following figure)
• Pectoral fins long, reaching posterior portion of anal fin
• Two distinct thin dark stripes along sides to caudal peduncle, contrasting with light background color
• Maximum size is 45 cm standard length (1.55 kg all-tackle record)

Prionotus longispinosus

• Branchiostegal rays very light-colored (whitish)
• Pectoral fins with small light-colored spots
• Caudal peduncle without a light-colored blotch on dorsal side
• Maximum size is 35 cm standard length

Prionotus martis

• Throat (gular area) is completely scaled
• Branchiostegal rays are dark (dusky or black)
• Gill rakers on lower limb of first arch usually 9 (range = 8–11)
• Dorsal fin with two distinct dark spots between 1^st and 2^nd, and 4^th and 5^th spines (see following figure)
• Body heavily spotted
• Small sized (maximum size is 18 cm standard length)

Prionotus ophryas

• Nasal cirri are present
• Preopercular spine long, reaching beyond posterior edge of operculum
• Pectoral fins very long, reaching well beyond posterior edge of anal fin
• Pectoral fins rounded
• Body color is variable, but not silvery
• Usually in 18–64 m depth
• Small sized (maximum size is 20 cm standard length)

Prionotus paralatus

• Nasal spines are absent
• Preopercular spine is long, reaching just beyond operculum
• Pectoral fins with dark spots and some scattered pinkish coloration throughout
• Reported to hybridize with P. alatus between Gulfport and Panama City
• Most abundant in 60–120 m depths
• Small sized (maximum size is 18 cm standard length)

Prionotus roseus

• Pectoral fins long, reaching or approaching the posterior edge of anal fin
• Dorsal free ray of pectoral fins short, not reaching posterior edge of pelvic fins
• Pectoral fins with bright blue or dark ocellated spots throughout (sometimes not ocellated)
• Pectoral fins with dark ventral edge (edge not blue)
• Small sized (maximum size is 20 cm standard length)

Prionotus rubio

• Nasal spines are absent
• Pectoral fins long, lowermost rays reaching beyond the posterior edge of anal fin
• Pectoral fins uniformly very dark (blackish) with distinct blue margin on ventral edge (see following figure)
• Small sized (maximum size is 23 cm standard length)

Prionotus scitulus

• Throat (gular area) is completely without scales
• Branchiostegal rays light-colored (whitish), never dusky or black
• Gill rakers on lower limb of first arch usually 11 (range = 10–13)
• Dorsal fin with two distinct dark spots between 1^st and 2^nd, and 4^th and 5^th spines (see following figure)
• Body spotted throughout
• Maximum size is 25 cm standard length
• Often found in shallow bays

Prionotus stearnsi

• Mouth with small bony knob on ventral side at symphysis
• Throat (gular area) completely scaled
• Pectoral fins relatively short (not reaching beyond origin of anal fin)
• Pectoral fins very dark (blackish)
• Trunk light colored (silvery)
• Small sized (maximum size is 18 cm standard length)

Prionotus tribulus

• Head is large and broad
• Total gill rakers on first arch usually 8–16
• Pectoral fin with broad, dark vertical bands (may be narrow in Gulf of Mexico specimens)
• Maximum size is 35 cm standard length

Sources Cited:

Hastings, R.W.  1979.  The origin and seasonality of the fish fauna on a new jetty in the northeastern Gulf of Mexico.  Bulletin of the Florida State Museum, Biological Sciences 24(1):1–124.

McEachran, J.D. and J.D. Fechhelm.  2005.  Fishes of the Gulf of Mexico, Volume 2: Scorpaeniformes to Tetraodontiformes.  University of Texas Press, Austin, TX.

Richards, W.J. and G.C. Miller.  2002.  Searobins.  Pp. 1266–1277.  In:  Carpenter, K.E. (ed.), FAO Species Identification Guide for Fishery Purposes:  The Living Marine Resources of the Western Central Atlantic.  Vol. 2:  Bony Fishes Part 1 (Acipenseridae to Grammatidae).  FAO, Rome, Italy.

Robins, C.R. and G.C. Ray.  1986.  A Field Guide to Atlantic Coast Fishes of North America.  Houghton Mifflin Co., New York, NY.

What types of invertebrates can be identified from the remains contributing to the sediment in continental slope waters?  An ANAMAR biologist set out to learn just what groups of invertebrates could be identified from sediment collected in the deep waters off Port Everglades Harbor, Florida.  The sample was collected using a custom-made bucket sampler that was towed a short distance across the sediment surface in 664 feet of water off Port Everglades Harbor (Fort Lauderdale), Florida. 

About 2 gallons of the sediment was sieved through a 2-mm screen.  An additional 15 ounces of the sediment was wet-sieved with a 0.8-mm fine-mesh screen to uncover the remains of smaller invertebrates.  The sample matrix was mostly greenish-gray silty fine sand, most of which was washed away during the sieving process.  The remaining shells and other hard objects were left to dry over a few week’s time.  The remains were then identified and photographed. 

So what was found?

The sample contained:

• Foraminiferidas (commonly called foraminifera or forams)
• Mollusks
  • Bivalves
  • Sea butterflies of the order Thecosomata (previously of Pteropoda)
  • Other gastropods
• Crustacean (crab and shrimp) remains
• Echinoid (sea urchin) spines and tests

Are any of these remains fossilized? 

All of the shells and other remains appeared to be of Holocene age (modern).  Thus, none of the remains represent fossils.  Based on the age of the animal remains, it appears that the sediment surface is mostly recently deposited material.  However, the silt component of the sediment sample may have been older than the shell component of the sample.

Although harbor sediment is often subjected to physical, chemical, toxicological, and bioaccumulation analyses, the biological properties often go unnoticed and unappreciated during such testing.  What can the biological properties—the critters inhabiting the sediment—tell us about the age and history of the sediment?  An ANAMAR biologist set out to learn just that, by sieving extra material from a recent sediment sampling event at Canaveral Harbor, Florida.

About 12 gallons of extra sediment sampled from Canaveral Harbor, Florida, was wet sieved with a 2-mm screen to uncover modern and fossil animal remains.  The sample matrix was mostly gray-green colored clay which was washed away during the sieving process.  The remaining shell hash component left in the sieve showed that it contained at least the remains of mollusks.  The shells and other hard objects were left to dry over a few week’s time.  The remains were then identified and photographed.

So what was found?

The sample contained:

• Crustacean (crab and shrimp) claws including the purse crab genus Persephona
• Barnacle shells
• Echinoid (sea urchin) spines and other remains
• Mollusk shells
  • Ark shells of the order Arcoida
  • Tusk shells including the genus Dentalium
  • Wentletraps including Epitonium cf. rupicola
  • The limpet Diodora cf. floridana
  • The land snail Polygyra septemvolva (Florida flatcoil)
• Fish remains
  • A tooth from Carcharhinus isodon (finetooth shark)
  • Vertebrae from teleost (bony) fishes
  • A pectoral spine from the sea catfish family Ariidae

Are any of these remains fossilized? 

It is not always easy to differentiate a fossil from the remains of modern animals, especially when considering the remains of mollusks having calcium carbonate shells.  However, we can say with certainty that the limpet shell is a fossil because the species Diodora floridana lived only during the Pleistocene (it has been extinct for thousands of years [Peterson and Peterson 2008]).  The Florida flatcoil shells may also represent fossils as they appeared to contain consolidated mineralized material filling the internal voids, and were a much darker than modern shells of this species.  The finetooth shark tooth also represents a fossil although this species still occurs around Florida today.  Most shark teeth found in sediment or on the ground are fossils because, although sharks are abundant in today’s oceans and they continually lose and replace teeth throughout their lives, it takes a build-up of teeth over thousands or millions of years for them to be numerous enough to be easily found. 

Most of the remaining shells are Holocene in age (modern).  Overall, the shells within the sediment range in age from Holocene (recent) to the Pleistocene (12,000 to 2.6 million years ago), based on the taxonomy of the animal remains.  The presence of the land snail Polygyra septemvolva among the remains of the marine animals suggests that the sediment had been mixed with other deposits originating from terrestrial sources.

Source Cited:

Peterson, C. and B. Peterson.  2008.  Southern Florida’s Fossil Seashells.  Blue Note Publications, Inc., Cocoa Beach, FL.

FULL WRITE-UP:

Action levels provided by the U.S. Food and Drug Administration (FDA) are used as screening benchmarks during Tier III testing evaluations under Section 103 of the Marine Protection, Research, and Sanctuaries Act of 1972 (MPRSA).  These action levels are based on human health and economic considerations and represent limits above which the FDA can take legal action to remove products from the marketplace (EPA and USACE 1991, known as the ‘Green Book’).  The Southeast Regional Implementation Manual (EPA and USACE 2008, known as the ‘SERIM’) provides guidance for MPRSA Section 103 testing evaluations in the southeastern United States.  Some discrepancies exist between the action levels provided by the FDA and those given in the SERIM.  The table below compares action levels between the SERIM, the source document to the SERIM (FDA 2001), and the most current FDA action levels (FDA 2011).

The action level presented in the SERIM for cadmium in bivalve tissue is switched with that of crustacea (see table above).  The SERIM applies the crustacea action levels to include polychaete worms as the FDA lacks any action levels intended for polychates.  The action levels given in the SERIM for mercury are intended for use specifically for methylmercury (FDA 2001 and 2011) rather than for total mercury.

The SERIM (Section 3.3.2.2 [page 24]) and the Green Book (Section 6.3 [not paginated]) suggest the reader use an updated version of the FDA action levels when available.  FDA action levels have been updated since the SERIM was published in 2008 and, therefore, it is useful to refer to FDA (2011) for some or all action levels rather than Appendix H of the SERIM.  However, since FDA (2011) omits arsenic, cadmium, chromium, lead, and nickel, the values given in FDA (2001) may, with the approval of USACE, continue to be used as screening benchmarks for these metals in Tier III evaluations.

Sources Cited Above:

U.S. Environmental Protection Agency and U.S. Army Corps of Engineers.  1991.  Evaluation of Dredged Material Proposed for Ocean Disposal, Testing Manual [Green Book].  EPA 503-8-91-001.  EPA, Office of Marine and Estuarine Protection and Department of the Army, USACE, Washington, DC.

U.S. Environmental Protection Agency and U.S. Army Corps of Engineers.  2008.  Southeast Regional Implementation Manual (SERIM), Requirements and Procedures for Evaluation of the Ocean Disposal of Dredged Material in Southeast U.S. Atlantic and Gulf Coast Waters.  EPA Region 4, Atlanta, GA, and USACE South Atlantic Division, Atlanta, GA.

U.S. Food and Drug Administration.  2001.  Fish and Fishery Products Hazards and Controls Guidance, Third Edition — June 2001 [online document].  Accessed 02/25/11 online at:  http://www.fda.gov/Food/‌GuidanceComplianceRegulatoryInformation/GuidanceDocuments/Seafood/FishandFisheriesProductsHazardsandControlsGuide/default.htm.

U.S. Food and Drug Administration.  2011.  Fish and Fishery Products Hazards and Controls Guidance, Fourth Edition — April 2011 [online document].  Accessed 01/09/12 online at:  http://www.fda.gov/Food/‌GuidanceComplianceRegulatoryInformation/GuidanceDocuments/Seafood/FishandFisheriesProductsHazardsandControlsGuide/default.htm.