Research Activities: The Potential for Fish Stocking to Spread Disease to Aquatic Amphibians

Background

       Amphibian populations and species are declining or disappearing from many regions and habitats world wide (Stuart et al., 2004). No single cause has been demonstrated, although acid precipitation, environmental contaminants, introduction of nonindigenous species, disease agents, climate change, parasites, and the effects of UV-B radiation have been suggested as factors causing amphibian declines. Indeed, several factors may interact in such a manner as to threaten species and populations locally or regionally (Carey and Bryant, 1995).  However, the impacts of disease can be especially devastating, and declines due to disease, particularly from chytrid fungus and ranaviruses (Berger et al., 1998; Chinchar, 2002), are well documented from many disparate regions, including North America (Daszak et al., 2003; Kiesecker et al., 2004).  In addition to the more well-known fungi and viruses, a disease of undetermined affinity (Anuraperkinsus) is now known to have had serious effects on populations of ranid frogs in the southeast (Rana sevosa in Mississippi; R. Seigel, C.K. Dodd, Jr., unpublished data). Of additional relevance, the spread of pathogens has been linked specifically with the introduction of nonindigenous amphibians (Mazzoni et al., 2003; Hanselmann et al., 2004; Beard and O'Neill, 2005; Jancovich et al., 2005).

       The primary objective of this research is to determine whether diseases which could have a detrimental effect on southeastern amphibians (primarily chytrids, ranaviruses, Anuraperkinsus, saprolegniasis) are present in amphibian larvae living in warm-water National Fish Hatcheries (NFHs) in the southeastern U. S. Second, we want to make a preliminary assessment of the potential for amphibian larvae in stocking locations to have a higher incidence of disease than non-stocked locations. This research will provide a foundation for an ongoing assessment of the extent of disease in southeastern amphibians, and the role fish hatcheries may play in disease spread.  An examination of hatchery records also will allow a preliminary assessment of the extent to which hatchery shipments could contribute to the mixture of amphibian genotypes from very different habitats on the southeastern coastal plain.  Although the results may not demonstrate that hatchery shipments have spread disease in the past, they will help in the prevention of future disease outbreaks by allowing researchers to make recommendations to minimize such threats. In order to carry out this research, we partnered with the National Wildlife Health Center (NWHC) and various southeastern USFWS entities (NWRs and NFHs). Larvae were (and are being) screened at the NWHC for pathogens.

       In the southeastern U. S., warm water fish hatcheries supply National Wildlife Refuges and other land management agencies with stock for sport fishing, ecological restoration, and as food vital in endangered species management. Several million fish may be transported from one region to another across state lines in a single restocking event. For example, three million bluegills were stocked at HNNWR in 2004 in order to provide food for a nesting colony of endangered Wood Storks. With the fish come a host of other aquatic invertebrates and vertebrates, including tadpoles and potentially salamander larvae. Shipments are not screened for amphibian larvae or amphibian diseases. Moving large numbers of non disease-screened amphibian larvae throughout a region has the potential to transfer disease pathogens quickly and with serious consequences to resident amphibian populations. For example, chytrid fungi can remain virulent for seven days in contaminated water, thus offering the potential for disease transmission even without direct contact with infected amphibians (Johnson and Speare, 2003). In South Carolina, chytrids were first reported in bullfrogs in 1978 from the Savannah River Site, not far from the USFWS Orangeburg NFH, a source of fish stock for the southeastern Atlantic Coastal Plain. Chytrid infections in amphibians also have been reported from the coastal plain of North Carolina.

Methods

       We sampled tadpoles at four warm water NFHs (Welaka, FL., Warm Springs, GA., Orangeburg, SC, Edenton, NC) and at HNNWR (a fish stocked site) from May to July, 2005. The species of most interest a priori were ranid frogs (R. catesbeiana, R. clamitans, R. grylio, R. sphenocephala) which are easily captured by dip-netting or in crayfish traps, and which have been shown to be susceptible to chytrid, ranavirus and other pathogens in the southeast. Other species were collected as they were encountered (see Results and Appendix III). We sampled a variety of ponds in order to secure a representation of species, securing usually five tadpoles per species (as per NWHC instructions).  Tadpole samples were housed separately, and packed and shipped within 4-hr to Madison WI using the packing equipment and standard operating procedures developed by the NWHC (http:www.nwhc.usgs.gov/research/amph_dc/ sop_mailing. html).

       Necropsy: Amphibians that were dead on arrival at NWHC were  necropsied the same day as they are received. Live larvae were euthanized in 1:500 solution of MS222 (methanesulfonate salt, Sigma Chemical Co., St. Louis, Missouri. External and internal examinations were performed using a dissecting microscope equipped with a 35 mm camera.

       Virus isolation: Samples of the liver, mesonephros ("kidney") and spleen were pooled for virus cultures and isolations will be attempted on fathead minnow cell lines (Docherty et al., 2003, Journal of Wildlife Diseases 39:556-566).

       Bacterial and fungal cultures: Samples of liver, urine, mesonephros, bile, spleen or lung were  submitted for aerobic bacterial cultures.  A 2 mm  3 mm segment of cloaca and a 2-4 mm segment of distal toe (when available) were submitted for fungal cultures. Tissues and body fluids for routine aerobic bacterial cultures (approximately 1 mm³) were placed directly into vials of 2 ml tryptic soy broth with glycerine (TSB) and incubated at room temperature (25-27ºC). Cultures for Salmonella spp. were done in Rappaport-Vassiliadis R10 broth (Becton, Dickinson & Co., Cockeysville, Maryland). Subcultures were performed on 5 % sheep blood agar plates and eosin methylene blue plates. Biochemical identifications of bacterial isolates were performed using the Biolog MicroStation Microbial Identification System (Hayward, California). Fungal cultures were performed on Sabouraud dextrose agar plates with chloramphenicol and tetracycline (Hardy Diagnostics, Santa Maria, California). Fungal isolates were identified morphologically by features of their hyphae and spores.

       Parasitology: Parasites were identified to phylum during necropsies by a pathologist. Some helminths and insects were archived in hot buffered formalin or 70 % ethanol. Identifications to genus were based on external morphology of the live helminths at a dissecting microscope, tissue location in the host and histological features. Representative insects and helminths were identified by collaborating parasitologists and aquatic ecologists.

       Histology: Portions of ventral skin, digits, heart, liver, lung, spleen, mesonephros, stomach, intestine, pancreas, urinary bladder and gonads were fixed in 10 % buffered neutral formalin, processed routinely, sectioned at 5 microns, and stained with hematoxylin and eosin. Portions of liver, ventral skin, muscle, lung and mesonephros were placed in 1.8 ml cryovials and archived at –70ºC at NWHC (Madison, Wisconsin USA).

Results

       Tadpoles were found in large number at all the NFHs, and included a variety of taxa in various stages of development. Most species were what one might expect in large open-air ponds, such as bullfrogs and leopard frogs. However, a number of species associated with temporary ponds (Hyla femoralis, H. gratiosa, Gastrophryne carolinensis) also were collected. We sent the following specimens to the NWHC for disease screening:

Edenton NFH                20 tadpoles of 3 species; 15 Jun
Bufo terrestris, Hyla cinerea, Rana catesbeiana [large and small tadpoles]
 
Warm Springs NFH       20 tadpoles of 4 species; 13 Jun
Bufo terrestris, Hyla cinerea, Rana catesbeiana, Rana clamitans
 
Orangeburg NFH           25 tadpoles of 5 species; 14 Jun
Bufo terrestris, Hyla cinerea, Hyla femoralis, Gastrophryne carolinensis, Rana sphenocephala
 
Welaka NFH                   85 tadpoles of 4 species; 26 May
Hyla cinerea (2), Hyla gratiosa (5), Hyla squirella (30), Rana sphenocephala (48)
 
Harris Neck NWR          10 tadpoles of 1 species; 26 Jul
Rana catesbeiana (two different phenotypes, however).  This is a location where  fish have been stocked.

       We have received a preliminary report of the initial disease-screening results, covering virus cultures, bacterial cultures, some parasitological exams and gross examinations. This report (see Appendix III) indicated that no positive indications of disease had been detected as yet from any of the tadpoles collected.

       Histological examinations still are in progress. Preliminary results, however, confirm the presence of a chytrid infection of the oral cavity in four of five R. catesbeiana examined from Warm Springs NFH. As soon as the final report is available, we will provide the results to ARMI and the USFWS.

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