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.

A 2018 study published in the peer-reviewed journal Gulf and Caribbean Research represents a collaborative effort between ANAMAR biologist Jason Seitz and colleagues at the University of New England and the Florida Fish and Wildlife Research Institute.  Southern stingrays (Hypanus americanus, previously Dasyatis americana) were sampled and measured at fishing tournaments from 2004 and 2012 in Charlotte Harbor, Florida.  Vertebral centra obtained from the stingrays were sectioned and mounted on slides, and their growth bands were counted by two independent readers.  A total of 18 female southern stingrays, measuring from 412 to 1127 mm disc width (DW) were aged.  Ages ranged from 0 to 17 years.  The results suggest that southern stingrays obtain relatively old ages (17 years) and large sizes (to at least 1127 mm DW).  The results are comparable to other large stingray species such as the brown stingray (Bathytoshia lata; to at least 24 years and 1790 mm DW) and the common stingray (Dasyatis pastinaca; to at least 16 years and 1140 mm DW).  The results are also comparable to size-at-width data from a captive population of southern stingrays (13 years for a 1000-mm DW captive female, compared to an estimated 12 years for a 1005-mm DW wild female in this study).  The age-at-width estimates given in the 2018 study provide a preliminary foundation for future studies on age and growth of southern stingrays for the generation of mortality rates, production rates, and growth models, such as a Von Bertalanffy growth function or a Gompertz function, once ages of each life stage are obtained.  This study is the first investigation of the age-at-widths of wild southern stingrays. 

Here is a link to the open-access paper: Southern-stingray-age-and-growth-FINAL-2018

Source

Hayne, A.H.P., G.R. Poulakis, J.C. Seitz, and J.A. Sulikowski.  2018.  Preliminary age estimates for female Southern Stingrays (Hypanus americanus) from southwestern Florida, USA.  Gulf and Caribbean Research 29:SC1–4.  DOI:10.18785/gcr.2901.03  

An anonymous letter to The Pennsylvania Journal in December 1775 describes how the writer observed an image of a rattlesnake depicted on a drum belonging to a Marine with the words ‘Don’t Tread on Me’ spelled underneath the snake’s image.  The writer went on to theorize as to why this symbol was chosen by the Marine and its intended meaning.  The writer began by outlining the characteristics that distinguish rattlesnakes from other animals and supposed why these same properties may be appropriately used to symbolize the United States of America:

“I recollected that her eye excelled in brightness, that of any other animal, and that she has no eye-lids.  She may therefore be esteemed an emblem of vigilance.  She never begins an attack, nor, when once engaged, ever surrenders: she is therefore an emblem of magnanimity and true courage.  As if anxious to prevent all pretensions of quarreling with her, the weapons with which nature has burnished her, she conceals, in the roof of her mouth, so that, to those who are unacquainted with her, she appears to be a most defenseless animal; and even when those weapons are shown and extended for her defense, they appear weak and contemptible; but their wounds however small, are decisive and fatal.  Conscious of this, she never wounds ‘till she has generously given notice, even to her enemy, and cautioned him against the danger of treading on her.

       -An American Guesser”

Although the above paragraph was written anonymously, history scholars believe the writer to have been Benjamin Franklin.  The entire letter can be viewed at http://greatseal.com/symbols/rattlesnake.html.

This short blog will consider the venom attributes of rattlesnakes.

What is the Purpose of Rattlesnake Venom?

There are two main categories of venoms: those that are intended to subdue prey and those that are intended to dissuade predators.  Rattlesnakes evolved their venom to subdue prey and to begin the process of digestion.  Figure 1 shows a timber rattlesnake (Crotalus horridus) in the act of consuming an eastern gray squirrel (Sciurus carolinensis) after subduing it with venom.  They have a special suite of proteins and enzymes to help accomplish this task.  Unfortunately, although rattlesnakes evolved their venom for use in obtaining food rather than as a defensive measure, a defensive bite nonetheless has the effect of these digestive compounds tearing through bodily tissues and causing pain, swelling, and necrosis. 

Figure 1.  This timber rattlesnake (Crotalus horridus) was photographed by the author while it consumed an eastern gray squirrel (Sciurus carolinensis) in Alachua County, Florida, after subduing it with venom.  This snake is among the population thought to have a higher concentration of neurotoxic venom than populations north of Interstate 10.

Photographed and copyrighted by the author, Jason Seitz

What Is Rattlesnake Venom Composed of?

The venom of rattlesnakes is a mixture of hemotoxins and neurotoxins, but are mostly hemotoxins.  Hemotoxins target tissues and blood, causing hemorrhaging and necrosis.  Their venom is really a cocktail of chemical elements.  Neurotoxins target the nervous system, some of which can cause paralysis.  While each species of venomous snake has its own particular cocktail of proteins and enzymes compared to other species, there is some evidence to suggest that the relative concentration of neurotoxins to hemotoxins may vary regionally even within a given species of snake.  For instance, a significant percentage of the timber rattlesnakes south of Interstate 10 (I-10) in Florida are believed by some researchers to have a higher concentration of neurotoxic venom than do timber rattlers north of this corridor.  The various proteins and enzymes in rattlesnake venom have a synergistic effect that has evolved to trigger total cardiovascular collapse of the snake’s intended prey.  When a rattler bites in defense, the effects are watered down due to the large size of a person compared to their prey (typically a rodent).

Some recent works by scientists have found that some forms of hemotoxic venoms are not immunogenic, meaning that they do not trigger an immune response by the victim.  They slip by the immune system of the envenomated animal so no antibodies are produced to fight the toxins.  This is troubling since antivenoms are produced by injecting venom into a large animal, typically a horse, and later harvesting the antibodies created by the horse that can be later used to treat bite victims. 

Hemotoxin venoms such as those of rattlesnakes begin to disassemble the structural components of blood vessels and tissues as soon as they are injected.  This is done by metalloproteases, which are proteases enzymes that use a metal as a catalyst in the hydrolysis of peptide bonds.  Because these enzymes break down even the proteins responsible for keeping the cell walls of blood vessels intact, localized hemorrhaging results, sending blood into surrounding tissues.  The same metalloproteases also act to break down skeletal muscles.  Another component of rattler toxin, phospholipases cause the death of muscle tissue by attacking their cellular membranes.  Some of these phospholipases have enzymes that create holes in the muscle cell walls by breaking apart the phospholipids that hold the membranes together.  Other phospholipases use as‑yet‑unidentified means of destroying muscle cells. 

There are still other enzymes contained within rattler venom that cause destruction.  These include hyaluronidases and serine proteases, each having its own type of destructive mechanism.  Some chemical compounds from the venom travel far from the site of the bite and wreak havoc on blood vessels and skeletal muscles elsewhere in the body. 

In addition to the destructive actions of the venom components themselves, some proteins trick our own immune system to fight against our own cells.  Specifically, the actions of metalloproteases and phospholipases trigger an immune response at the site of the wound.  Immune cells such as leukocytes signal an increased immune response by releasing messengers such as interleukin-6.  Since the venom components are not a cohesive force, and with no bacteria to attack, the immune system instead launches an attack that adds to the destruction of our own tissues.  The damage done by our own immune system is doubly troubling considering that antivenom does not help to mitigate its effects.  Studies have found that, when the immune system is shut down, necrotic effects of snake venom are greatly reduced.  The use of Benadryl, for instance, can lessen the swelling and edema associated with envenomation.

A Note on Conservation

Although snakes are often feared and persecuted throughout much of their ranges in the United States and elsewhere, they nonetheless occupy a valuable place in the ecology of many ecosystems.  There are hundreds of species of snakes in the U.S., but only a small portion of them are venomous.  For example, in Florida there are 50 species of snakes but only 6 (12%) of those species are venomous.  Venomous snakes can be safely and effectively avoided by using common sense.  If a snake is thought to be venomous, or if it is not known whether it is venomous, the safe thing to do is to leave it alone.  Remember that most bite victims are bitten because they are attempting to handle or kill the snake. 

Snakes perform valuable services such as controlling rodents and other pests.  A recent study has shown that timber rattlesnakes may reduce the incidence of Lyme disease in the northeastern U.S. by preying on the host (rodents) that carries the tick that spreads the disease.  An estimated 2,500 to 4,500 ticks were removed from the northeastern U.S. study areas each year by timber rattlesnakes consuming their mammal hosts.  

Recent research suggests that rattlesnakes may help disperse the seeds of grasses and other plants.  Rodents eat grass seeds and those of other plants, but the seeds do not normally survive through the digestive process of rodents.  However, rodents often carry the seeds in their cheek pouches, and if the rodent is then killed and eaten by a rattlesnake, these seeds are capable of passing through the snake’s digestive tract and remaining viable.  Three species of desert-dwelling rattlesnakes were found to have consumed rodents with seeds in their cheek pouches and that the seeds are capable of germinating in the snake’s colon and may be passed with the snake’s feces, allowing dispersal of the plant propagules.

Rattlesnakes and many other species of snakes are experiencing declines in populations in the United States due to loss of habitat, continued persecution, and emerging diseases such as the snake fungus Ophidiomyces ophiodiicola.  The eastern diamondback rattlesnake (Crotalus adamanteus) (Figure 2) has declined to the point where the species has been petitioned for protection as a threatened species under the Endangered Species Act.

Figure 2.  The eastern diamondback rattlesnake (Crotalus adamanteus) is one of the more well-known (and often persecuted) rattlesnake species.
Photographed and copyrighted by the author, Jason Seitz

A Fun Way to Report your Amphibian and Reptile Sightings Using Citizen Science

Use the HerpMapper mobile phone app to record and submit your amphibian and reptile sightings!  It’s fun, simple, and easy to do.  See the website https://www.herpmapper.org/ for more information and to download the app.

Sources

Adkins, C.L., D.N. Greenwald, D.B. Means, B. Matturro, and J. Ries.  2011.  Petition to List the Eastern Diamondback Rattlesnake (Crotalus adamanteus) as Threatened under the Endangered Species Act. Petition submitted 08/11/11 to U.S. Fish and Wildlife Service (USFWS), Washington, D.C., and USFWS Region 4, Atlanta, GA.

Brown, W.S.  1993.  Biology, Status, and Management of the Timber Rattlesnake (Crotalus horridus): A guide for Conservation.  Society for the Study of Amphibians and Reptiles, Herpetological Circular No. 22, The University of Kansas, Lawrence, KS.

Kabay, E., N.M. Caruso, and K.R. Lips.  2013.  Timber Rattlesnakes May Reduce Incidence of Lyme Disease in the Northeastern United States.  98^th Annual Meeting of the Ecological Society of America, 08/06/13, Minneapolis, MN.  Accessed online 02/05/18 at https://eco.confex.com/eco/2013/webprogram/Paper44305.html.

Lorch, J.M, S. Knowles, J.S. Lankton, K. Michell, J.L. Edwards, J.M. Kapfer, R.A. Staffen, E.R. Wild, K.Z. Schmidt, A.E. Ballmann, D. Blodgett, T.M. Farrel, B.M. Glorioso, L.A. Last, S.J. Price, K.L. Schuler, C.E. Smith, J.F.X. Wellehan, Jr., and D.S. Blehert.  2016.  Snake fungal disease: an emerging threat to wild snakes.  Philosophical Transactions of the Royal Society B 371: 20150457.  Accessed online 02/13/18 at http://rstb.royalsocietypublishing.org/content/royptb/371/1709/20150457.full.pdf.

Reiserer, R.S., G.W. Schuett, and H.W. Greene.  2018.  Seed ingestion and germination in rattlesnakes: overlooked agents of rescue and secondary dispersal.  Proceedings of the Royal Society B: 285: 20172755. http://dx.doi.org/10.1098/rspb.2017.2755.

Robertson, M.  2017.  pers. comm. regarding regional differences in neurotoxins in the timber rattlesnake.

Wilcox, C.  2016.  Venomous, How Earth’s Deadliest Creatures Mastered Biochemistry.  Scientific American / Farrar, Straus, and Giroux, New York, NY.

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.

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.