Ranaviruses have caused die-offs of amphibians across the globe. In North America, these pathogens cause more amphibian mortality events than any other pathogen. Field observations suggest that ranavirus epizootics in amphibian communities are common during metamorphosis, presumably due to changes in immune function. However, few controlled studies have compared the relative susceptibility of amphibians to ranaviruses across life stages. Our objectives were to measure differences in mortality and infection prevalence following exposure to ranavirus at four developmental stages and determine whether the differences were consistent among seven anuran species. Based on previous studies, we hypothesized that susceptibility to ranavirus would be greatest at metamorphosis. Our results did not support this hypothesis, as four of the species were most susceptible to ranavirus during the larval or hatchling stages. The embryo stage had the lowest susceptibility among species probably due to the protective membranous layers of the egg. Our results indicate that generalizations should be made cautiously about patterns of susceptibility to ranaviruses among amphibian developmental stages and species. Further, if early developmental stages of amphibians are susceptible to ranaviruses, the impact of ranavirus epizootic events may be greater than realized due to the greater difficulty of detecting morbid hatchlings and larvae compared to metamorphs.
Citation: Haislip NA, Gray MJ, Hoverman JT, Miller DL (2011) Development and Disease: How Susceptibility to an Emerging Pathogen Changes through Anuran Development. PLoS ONE 6(7): e22307. https://doi.org/10.1371/journal.pone.0022307
Editor: Raphaël Arlettaz, University of Bern, Switzerland
Received: March 19, 2011; Accepted: June 23, 2011; Published: July 22, 2011
Copyright: © 2011 Haislip et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: The University of Tennessee Institute of Agriculture and Tennessee Wildlife Resources Agency (TWRA) provided funding for this project. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Disease epidemics are driven by the complex interactions among the pathogen, host susceptibility, and the environment. Recent work in disease ecology seeks to understand mechanisms of pathogen infection during development that lead to developmental abnormalities and mortality events . There is increasing awareness that there are critical windows during development in which hosts are particularly sensitive to disease-causing agents leading to mortality, impairment, or malformation of the individual , . In humans, for example, differences in susceptibility to infection during development are demonstrated by the early childhood malformations and mortality associated with German measles (Rubella Virus; ). Such developmental perturbations can occur from exposure to toxins, parasites, and nutrient deficiencies , , . Thus, the connection between windows of developmental sensitivity and susceptibility to pathogens is an important mechanism in the emergence of wildlife diseases.
The role of pathogens in the recent declines of amphibians across the globe has received considerable attention . While amphibians are hosts for a diversity of pathogens , many die-off events have been associated with infection by ranaviruses , . Ranaviruses have been reported on five continents and are associated with nearly 50% of the reported amphibian mortality events in the United States , . Although ranaviruses have been well studied and characterized at the molecular level , , research has only recently begun to examine the mechanisms associated with ranavirus emergence in wild populations .
In 96% of reported ranavirus die-off events, recently metamorphosed individuals experienced the greatest mortality , . These field observations have led to the hypothesis that ranavirus epizootics in the wild occur most often as amphibians undergo metamorphosis, which is known to be a period of natural immune suppression . Previous studies suggest that there are varying degrees of immune system development across different amphibian life stages. Du Pasquier et al.  found that the production of thymic lymphocytes increases during larval development, drops substantially at metamorphosis, and peaks in adult Xenopus laevis. Decreases in immune function during metamorphosis are probably related to endogenous production of glucocorticoids associated with restructuring organ systems for postmetamorphic life . Thus, the immunological changes that occur during anuran development should affect host-pathogen interactions , . Unfortunately, experimental studies comparing the susceptibility of amphibians to pathogens at different developmental stages are rare , . Thus, the first objective of our research was to test for differences in susceptibility (as indexed by mortality and infection prevalence) to ranavirus among pre-terrestrial developmental stages in amphibians.
Traditionally, disease ecology has focused on pathogens that attack a single host, which has limited our ecological understanding of disease dynamics driven by pathogens that infect multiple host species –. There is growing evidence that amphibian species differ in their susceptibility to pathogens –. While not surprising, such variation in species susceptibility underscores the need for comprehensive studies that examine multiple host species to identify generalities that cannot be obtained from single-species studies. To date, very few studies have examined the relative susceptibility of amphibian larvae to ranaviruses , , , . Moreover, these studies tested only one developmental stage, thus their results may be limited. The second objective of our study was to identify trends in the relative susceptibility to ranavirus for seven North American anuran species.
All animal husbandry and euthanasia procedures followed an approved University of Tennessee IACUC protocol (#1755).
Animal collection and maintenance
We used seven anuran species for our study: Lithobates clamitans, L. pipiens, L. sylvaticus, Pseudacris feriarum, Hyla chrysoscelis, Scaphiopus holbrookii, and Anaxyrus americanus, which are widely distributed in eastern North America . Between February–July 2009, we collected 7–20 egg masses for each species from single populations (Table 1). Egg masses were collected within 48 hours of deposition, rinsed with sterile water, and transported in 19-L buckets filled with aged tap water to the University of Tennessee Joe Johnson Animal Research and Teaching Unit (JARTU). Egg masses were placed in covered (60% shade cloth) 300-L wading outdoor pools the day after collection to develop. After hatching, tadpoles were maintained in these pools and fed rabbit chow (Purina, St. Louis, Missouri) and ground TetraMin® (Tetra, Blacksburg, Virginia) ad libitum until used in the experiments. The experiments began as individuals reached the appropriate developmental stages (see below). Prior to each experimental trial, a random sample of 10 tadpoles was euthanized and frozen at −80°C for confirmation that they were negative for ranavirus using real-time quantitative polymerase chain reaction (qPCR, see Molecular Analyses section); all pre-experiment individuals tested negative.
A single isolate of Ranavirus was used for all experiments. The University of Georgia Veterinary Diagnostic and Investigational Laboratory (VDIL) extracted this isolate from morbid L. catesbeianus juveniles. Preliminary molecular analyses suggest that the isolate is similar to the ranavirus frog virus 3 (GenBank accession no. EF101698, ), and it has been shown to be virulent in anuran larvae . Titrated stock solutions of the isolate were sent overnight by the VDIL to the University of Tennessee for the experiments.
For each species, we conducted a 14-d experimental trial for each of four developmental stages: 1) embryo (stage 11), 2) hatchling (stage 21), 3) larval (stage 30), and 4) pro-metamorphosis (stage 41, ). For our experiments, embryos were contained in eggs. Experimental units for all trials were 1-L tubs filled with 0.5 L of aged tap water. The tubs were placed at a common shelf height in a completely randomized design at the JARTU laboratory facility. We randomly assigned a single individual to each tub. Treatments included a no-virus control and a virus exposure of 103 plaque-forming units (PFUs) mL−1 . Both treatments were replicated 20 times for a total of 40 experimental units per trial.
We inoculated the water (i.e., bath exposure) with 29.5 µL of Eagle's Minimal Essential Media (MEM) for the no-virus control tubs and 29.5 µL of MEM containing the virus for the virus tubs. The resulting virus concentration was 103 PFUs mL−1, which is within the range of doses used in other studies (102–106 PFUs mL−1; –) and ecologically relevant , . Given that some species in our study developed rapidly (e.g., S. holbrookii), we used a 3-day exposure in an attempt to target the intended developmental stage rather than a subsequent stage. After three days, individuals were removed from the containers, rinsed with sterile water, and placed into a new container with 500-mL of fresh aged tap water. For the remainder of the experiment, water was changed every three days to maintain water quality.
After each water change, individuals in the larval and metamorph experiments were fed ground TetraMin® at a daily rate of 8% body mass . Prior to the water change, we weighed a group of 10 non-experimental individuals housed under identical conditions to calculate food rations based on the average mass. Individuals in the embryo and hatchling experiments were fed if they reached stage 25 prior to the end of the experiment, which is when yolk reserves are exhausted and jaw development is complete in most species . After the initial exposure and water change, platforms were placed in the metamorph experimental units to allow individuals to crawl out of the water to complete metamorphosis. Once individuals in the metamorph stage experiments began tail resorption, feedings were terminated and water depth was slowly reduced until a minimal amount of water remained to provide moisture for the individual and TetraMin® was no longer added. Following tail resorption, individuals were fed 10 seed weevils (Callosobruchus sp.) every three days.
The experimental units were monitored three times daily for mortality. Dead larvae and metamorphs were necropsied using sterilized forceps and scissors. Because the kidneys and liver are known sites of ranavirus infection , we removed sections of these organs from each individual, placed the pooled sample in a 1.5-mL microcentrifuge tube, and froze at −80°C for molecular testing. Dead embryos and hatchlings were rinsed with sterile water and frozen at −80°C, because their small size prevented consistent necropsies. After 14 days, all live individuals were euthanized in benzocaine hydrochloride (1 g L−1) and the identical necropsy procedures followed. We set 14 days as the experiment duration because previous research has shown this is sufficient duration to observe disease from ranavirus infection with a 3-day water bath exposure .
For ranavirus testing, genomic DNA (gDNA) was extracted from a homogenate of the kidney and liver for tadpoles and metamorphs and from entire embryos (including vitelline membrane and mucoidal capsules) and hatchlings using a DNeasy Blood and Tissue Kit (Qiagen Inc., Valencia, CA). We used the Qubit™ fluorometer and the Quant-iT™ dsDNA BR Assay Kit to quantify the concentration of genomic DNA in each sample (Invitrogen Corp., Carlsbad, CA, USA) . The qPCR amplified a 70-bp region of the ranavirus major capsid protein. For each sample, we combined 12.5 µL of TaqMan Universal PCR Master Mix (Applied Biosystems, Foster City, California, USA), 1.5 µL of each primer (rtMCP-F [5′ – ACA CCA CCG CCC AAA AGT AC – 3′] and rtMCP-R [5′ – CCG TTC ATG ATG CGG ATA ATG – 3′]), and 1.5 µL of rtMCP-probe (5′-CCT CAT CGT TCT GGC CAT CAA CCA-3′). We added 0.25 µg of gDNA from each sample to standardize the total amount of gDNA added to the tubes. Because the volume containing this amount of gDNA varied depending on the gDNA concentration of the sample, we used the values from the fluorometer to calculate how much of the sample to add. We then added DNA grade water to the sample to bring the total volume to 30 µL. A SmartCycler® (Cepheid, Sunnyvale, California) thermal cycler was used for the qPCR. In each run of the qPCR, we included 4 controls, which were a ranavirus-negative tadpole sample, a negative DNA grade water sample, a ranavirus-positive tadpole sample, and a cultured virus sample. For each sample, we recorded the cycle number at which the sample crossed the fluorescent threshold level, which was set at 30 (i.e., CT value). Those samples that crossed the threshold level before CT = 30 were declared infected.
The response variables for each experiment included final mortality and infection prevalence calculated from binary data. Differences in final mortality and infection prevalence were tested among species and developmental stages using logistic regression analysis , . We did not include the control treatment in the analysis because control mortality was low resulting in low or zero counts for prevalence estimates of several developmental stages, which could have biased the logistic regression results , . Instead, median control mortality among developmental stages was provided for each species. If the Wald's chi-square test associated with the logistic regression analysis was significant, we used binomial tests that were Bonferroni corrected (α÷number of post-hoc comparisons) to test for pairwise differences between proportions . We estimated the likelihood of infection and mortality for each treatment in comparison with the treatment having the lowest rate by calculating odds-ratio statistics . If species and developmental stage effects interacted, we separated the analysis by species and performed a chi-square test for differences in mortality and infection prevalence among stages. All tests were performed at α = 0.05 using PROC LOGISTIC in the SAS® system . Test statistics and P-values were provided for evidence of differences in infection prevalence and mortality among effect levels. Test statistics with inequalities included results from more than one effect. Lastly, we regressed infection prevalence against mortality using linear regression in PROC GLM. Paired estimates for infection and mortality were the response variables and included in the analysis only if both proportions were not zero.
Across all species, final mortality and infection prevalence for the hatchling, larval and metamorph stages were significantly greater than the embryo stage (χ23>43.3, P<0.001). In the hatchling, larval, and metamorph stages, the odds of mortality were 3X, 4X, and 5X greater, respectively, when exposed to ranavirus compared to the embryo stage. Across all developmental stages, mortality and infection were greatest for L. sylvaticus and S. holbrookii, and were lowest for P. feriarum and A. americanus (χ26>40.67, P<0.001; Figure 1). Intermediate mortality and infection occurred for L. clamitans, L. pipiens, and H. chrysoscelis (Figure 1). Ranavirus exposed L. sylvaticus and S. holbrookii had 150X and 119X greater odds of mortality, respectively, than P. feriarum. Among species and stages, there was a strong positive relationship (R2 = 0.79) between mortality and infection prevalence (F1,20 = 74.52, P<0.001).
Similar shaded bars with unlike letters are different (P<0.006) by logistic regression analysis; n = 20 per developmental stage for each species.
Species and developmental stage effects interacted for final mortality and infection prevalence (χ218 = 128.9, P<0.001); thus, logistic regression analyses were performed separately for each species. For all species except L. sylvaticus, mortality and infection prevalence differed among developmental stages (χ23>12.6, P<0.006; Figure 1). For L. sylvaticus, infection prevalence was high (>82%) and did not differ among stages (χ23 = 6.3, P = 0.09). Mortality and infection prevalence were greatest during the metamorph stage for all Lithobates species. Mortality also was greatest during the metamorph stage for A. americanus, but these individuals were not infected with ranavirus. Mortality and infection prevalence tended to be greatest during the larval stage for the two hylid species: P. feriarum and H. chrysoscelis. The greatest infection and mortality for S. holbrookii occurred during the embryo, hatchling and larval stages, and were lowest during metamorphosis. Median control mortality was low for all species (≤10%), except for P. feriarum (22.5%), thus the results for this species should be interpreted cautiously. No control tadpoles tested positive for ranavirus infection.
Embryos that were contained within eggs were the least susceptible stage across species when exposed to ranavirus in a water bath. Previous research has shown that direct injection of ranavirus into embryos causes 97–100% mortality in L. pipiens . Thus, the vitelline membrane encasing the developing embryo or the mucopolysaccharide/mucoprotein capsules coating the surface of the egg likely affords protection against ranavirus infection. The mechanisms that contribute to this protection are unknown but may include structural barriers ,  or anti-viral properties of the egg capsules or membrane . Infection occurred in the embryo experiments for S. holbrookii and L. sylvaticus; however, embryos of these species hatched prior to the end of the 3-day virus challenge, hence exposing the hatchling to virions. No infection occurred during the embryo experiments in species that hatched after the virus challenge and first water change. Thus, it appears that eggs protect their developing embryos from ranavirus infection for the species we tested.
We documented high mortality during metamorphosis for all species of Lithobates tested, which is frequently the stage documented during anuran die-offs in the wild , . Cullen et al.  and Cullen and Owens  reported high susceptibility of several species of recently metamorphosed anurans compared to larvae or adults when exposed to ranavirus. Warne et al.  also reported higher mortality of ranavirus-exposed L. sylvaticus tadpoles during metamorphosis. High infection and mortality during metamorphosis may be associated with decreased immune function from endogenous production of corticosteroids and lymphocyte apoptosis , , , , which has been demonstrated in X. laevis , .
All other species that we tested had low mortality and infection prevalence during metamorphosis. The classic model of amphibian immune function during development, based on X. laevis, suggests that immune function increases through development then drops during metamorphosis . Down regulation of the immune system during metamorphosis may prevent destruction of new cell types that form for terrestrial life or may be a consequence of reduced physiological resources , . According to the X. laevis model of immune function, mortality associated with ranavirus infection should have been lowest during the larval (i.e., tadpole) stages. Lowest mortality during the larval stage did not occur for any of the anuran species that we tested, which may indicate that immune responses of North American anurans differ from those of X. laevis. The fully aquatic life cycle of X. laevis may result in unique immunological adaptations that are not shared with amphibian species that live terrestrially after metamorphosis. Pallister et al.  suggested that differences in larval development might contribute to differences in immune function. Indeed, comparative immunological studies between X. laevis and other anuran species are needed.
The greatest mortality and infection prevalence occurred during the hatchling stage for S. holbrookii, which was a different trend among the species that we tested. Infection and mortality decreased during the larval and metamorph stages, suggesting that immune function increased through development for this species. Compromised immunity during early development may be a consequence of physiological trade-offs associated with rapid development in this species. Spadefoots are among the fastest developing anuran species due to their association with ephemeral breeding sites , . Zettergren  reported cells synthesizing immunoglobulins (Ig) during embryogenesis and B lymphocytes circulating in pre-metamorphic L. pipiens at the onset of feeding. Leukocyte mobilization and anti-FV3 IgY antibody production have been reported as immune responses to ranavirus infection in X. laevis , . We hypothesize that development of these components of the amphibian immune system is delayed in S. holbrookii due to rapid growth during the embryo and hatchling stages.
Among species, L. sylvaticus was the most susceptible, with infection and mortality exceeding 80% in the hatchling, larval, and metamorph stages. These results support field observations for this species across its geographic range , , , . To date, no studies have explored the immunological mechanisms underlying the high susceptibility of L. sylvaticus to ranavirus compared to other species, although see Warne et al. . Cotter et al.  reported that poor lymphocyte production in the spleen was a mechanism driving high susceptibility of larval Ambystoma mexicanum to ranavirus. Significant increases in total leukocytes and natural killer cells are detected after 1 and 3 days post-infection with ranavirus, respectively, in X. laevis . Pre-metamorphic L. catesbeianus and X. laevis produce antibodies , , and therefore may resist ranavirus infection . Thus, minimal innate and adaptive immune response to ranavirus infection may be mechanisms contributing to high infection and mortality rates in ranavirus-exposed L. sylvaticus.
Our study is the first to report mortality of anuran hatchlings by ranavirus. The possibility for hatchling mortality from ranaviruses raises a significant conservation concern considering that detecting die-offs of hatchlings is extremely difficult in the wild. Differential susceptibility among developmental stages also indicates that studies that focus on one stage ,  may provide narrow insight into species susceptibility. If testing only one stage is feasible, we recommend using the larval stage because mortality and infection prevalence were either greater or similar to hatchling and metamorph stages for most species.
More research is needed investigating the role of immune function in regulating differences in susceptibility to ranavirus among anuran species. To date, few studies have quantified immune responses to ranavirus in pre-metamorphic amphibians , . Identifying commonalities among immunogenetic, evolutionary and life history traits of susceptible species will improve our understanding of host-pathogen interactions , and help facilitate identification of amphibian communities at greatest risk of ranavirus epizootics. To this end, we recommend that additional amphibian species and ranavirus strains be tested for relative susceptibility. Various multivariate techniques exist (e.g., canonical correspondence analysis, ) that can elucidate patterns between host characteristics and indices of susceptibility. We also encourage studies that challenge amphibian species with ranavirus at each stage of development and follow individual survival through metamorphosis. This knowledge is fundamental to developing stage-structured disease models that predict epizootic outcomes .
We thank the staff at the University of Georgia VDIL and UTIA JARTU Facility for logistical support. We especially thank C. Baldwin and D. Ingram for providing the virus isolate, L. Whittington for conducting the real-time PCR, and J. Hodges, B. Simpson, M. Campbell, and R. Long for various resources at JARTU. We also thank H. Edwards, E. Haislip, N. Hilzinger, L. Rucker, and W. Thompson for providing assistance with the experiments, and 2 anonymous referees for helpful comments on our manuscript. Collection of egg masses was approved by the TWRA (Scientific Collection Permit #1990).
Conceived and designed the experiments: JTH MJG. Performed the experiments: JTH NAH. Analyzed the data: JTH NAH MJG. Contributed reagents/materials/analysis tools: DLM. Wrote the paper: JTH NAH MJG DLM.
- 1. Johnson PJ, Kellermanns E, Bowerman J (2011) Critical windows of disease risk: amphibian pathology driven by developmental changes in host resistance and tolerance. Functional Ecology. PJ JohnsonE. KellermannsJ. Bowerman2011Critical windows of disease risk: amphibian pathology driven by developmental changes in host resistance and tolerance.Functional EcologyIn press. In press.
- 2. Pryor JL, Hughes C, Foster W, Hales B, Robaire B (2000) Critical windows of exposure for children's health: the reproductive sustem in animals and humans. Environmental Health Perspectives 108: 491–503.JL PryorC. HughesW. FosterB. HalesB. Robaire2000Critical windows of exposure for children's health: the reproductive sustem in animals and humans.Environmental Health Perspectives108491503
- 3. Webster WS (1998) Teratogen update: congenital rubella. Teratology 58: 12–23.WS Webster1998Teratogen update: congenital rubella.Teratology581223
- 4. Tabin CJ (1998) A developmental model for thalidomide defects. Nature 396: 322–323.CJ Tabin1998A developmental model for thalidomide defects.Nature396322323
- 5. Wake DB, Vredenburg VT (2008) Are we in the midst of the sixth mass extinction? A view from the world of amphibians. Proceedings of the National Academy of Sciences 105: 11466–11473.DB WakeVT Vredenburg2008Are we in the midst of the sixth mass extinction? A view from the world of amphibians.Proceedings of the National Academy of Sciences1051146611473
- 6. Wright KM, Whitaker BR (2001) Amphibian medicine and captive husbandry. Malabar, Florida: Krieger Publishing Company. 499 p.KM WrightBR Whitaker2001Amphibian medicine and captive husbandryMalabar, FloridaKrieger Publishing Company499
- 7. Green DE, Converse KA, Schrader AK (2002) Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996–2001. Annals of the New York Academy of Sciences 969: 323–339.DE GreenKA ConverseAK Schrader2002Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996–2001.Annals of the New York Academy of Sciences969323339
- 8. Muths E, Gallant AL, Campbell EHC, Battaglin WA, Green DE, et al. (2006) E. MuthsAL GallantEHC CampbellWA BattaglinDE Green2006The Amphibian Research and Monitoring Initiative (ARMI): 5-year report: U.S. Geological Survey Scientific Investigations Report 2006-5224. The Amphibian Research and Monitoring Initiative (ARMI): 5-year report: U.S. Geological Survey Scientific Investigations Report 2006-5224.
- 9. Converse KA, Green DE (2005) Diseases of tadpoles. In: Majumdar SK, Huffman JE, Brenner FJ, Panah AI, editors. Wildlife diseases: Landscape epidemiology, spatial distribution and utilization of remote sensing technology. Easton, Pennsylvania: The Pennsylvania Academy of Science. pp. 72–88.KA ConverseDE Green2005Diseases of tadpoles.SK MajumdarJE HuffmanFJ BrennerAI PanahWildlife diseases: Landscape epidemiology, spatial distribution and utilization of remote sensing technologyEaston, PennsylvaniaThe Pennsylvania Academy of Science7288
- 10. Chinchar VG (2002) Ranaviruses (family Iridoviridae): emerging cold-blooded killers - Brief review. Archives of Virology 147: 447–470.VG Chinchar2002Ranaviruses (family Iridoviridae): emerging cold-blooded killers - Brief review.Archives of Virology147447470
- 11. Chinchar VG, Essbauer S, He JG, Hyatt A, Miyazaki T, et al. (2005) Iridoviridae. In: Fauguet CM, Mayo MA, Maniloff J, Desselberger U, Ball LA, editors. Virus taxonomy: 8th report of the international committee on the taxonomy of viruses. London: Elsevier. pp. 163–175.VG ChincharS. EssbauerJG HeA. HyattT. Miyazaki2005Iridoviridae.CM FauguetMA MayoJ. ManiloffU. DesselbergerLA BallVirus taxonomy: 8th report of the international committee on the taxonomy of virusesLondonElsevier163175
- 12. Gray MJ, Miller DL, Hoverman JT (2009) Ecology and pathology of amphibian ranaviruses. Diseases of Aquatic Organisms 87: 243–266.MJ GrayDL MillerJT Hoverman2009Ecology and pathology of amphibian ranaviruses.Diseases of Aquatic Organisms87243266
- 13. Du Pasquier L, Schwager J, Flajnik MF (1989) The immune system of Xenopus. Annual Review of Immunology 7: 251–275.L. Du PasquierJ. SchwagerMF Flajnik1989The immune system of Xenopus.Annual Review of Immunology7251275
- 14. Rollins-Smith LA (1998) Metamorphosis and the amphibian immune system. Immunological Reviews 166: 221–230.LA Rollins-Smith1998Metamorphosis and the amphibian immune system.Immunological Reviews166221230
- 15. Gantress J, Maniero GD, Cohen N, Robert J (2003) Development and characterization of a model system to study amphibian immune responses to iridoviruses. Virology 311: 254–262.J. GantressGD ManieroN. CohenJ. Robert2003Development and characterization of a model system to study amphibian immune responses to iridoviruses.Virology311254262
- 16. Scotthoefer AM, Cole RA, Beasley VR (2003) Relationship of tadpole stage to location of echinostome cercariae encystment and the consequences for tadpole survival. Journal of Parasitology 89: 475–482.AM ScotthoeferRA ColeVR Beasley2003Relationship of tadpole stage to location of echinostome cercariae encystment and the consequences for tadpole survival.Journal of Parasitology89475482
- 17. Daszak P, Cunningham AA, Hyatt AD (2000) Wildlife ecology - Emerging infectious diseases of wildlife: threats to biodiversity and human health. Science 287: 443–449.P. DaszakAA CunninghamAD Hyatt2000Wildlife ecology - Emerging infectious diseases of wildlife: threats to biodiversity and human health.Science287443449
- 18. Dobson A, Foufopoulos J (2001) Emerging infectious pathogens of wildlife. Philosophical Transactions of the Royal Society of London Series B-Biological Sciences 356: 1001–1012.A. DobsonJ. Foufopoulos2001Emerging infectious pathogens of wildlife.Philosophical Transactions of the Royal Society of London Series B-Biological Sciences35610011012
- 19. Woolhouse MEJ, Haydon DT, Antia R (2005) Emerging pathogens: the epidemiology and evolution of species jumps. Trends in Ecology & Evolution 20: 238–244.MEJ WoolhouseDT HaydonR. Antia2005Emerging pathogens: the epidemiology and evolution of species jumps.Trends in Ecology & Evolution20238244
- 20. Johnson PTJ, Hartson RB (2009) All hosts are not equal: explaining differential patterns of malformations in an amphibian community. Journal of Animal Ecology 78: 191–201.PTJ JohnsonRB Hartson2009All hosts are not equal: explaining differential patterns of malformations in an amphibian community.Journal of Animal Ecology78191201
- 21. Schock DM, Bollinger TK, Chinchar VG, Jancovich JK, Collins JP (2008) Experimental evidence that amphibian ranaviruses are multi-host pathogens. Copeia 133–143.DM SchockTK BollingerVG ChincharJK JancovichJP Collins2008Experimental evidence that amphibian ranaviruses are multi-host pathogens.Copeia133143
- 22. Hoverman JT, Gray MJ, Miller DL (2010) Anuran susceptibilities to ranaviruses: role of species identity, exposure route, and a novel virus isolate. Diseases of Aquatic Organisms 89: 97–107.JT HovermanMJ GrayDL Miller2010Anuran susceptibilities to ranaviruses: role of species identity, exposure route, and a novel virus isolate.Diseases of Aquatic Organisms8997107
- 23. Kilpatrick AM, Briggs CJ, Daszak P (2010) The ecology and impact of chytridiomycosis: an emerging disease of amphibians. Trends in Ecology & Evolution 25: 109–118.AM KilpatrickCJ BriggsP. Daszak2010The ecology and impact of chytridiomycosis: an emerging disease of amphibians.Trends in Ecology & Evolution25109118
- 24. Schock DM, Ruthig GR, Collins JP, Kutz SJ, Carriere S, et al. (2009) Amphibian chytrid fungus and ranaviruses in the Northwest territories, Canada. Diseases of Aquatic Organisms 92: 231–240.DM SchockGR RuthigJP CollinsSJ KutzS. Carriere2009Amphibian chytrid fungus and ranaviruses in the Northwest territories, Canada.Diseases of Aquatic Organisms92231240
- 25. Cullen BR, Owens L (2002) Experimental challenge and clinical cases of Bohle iridovirus (BIV) in native Australian anurans. Diseases of Aquatic Organisms 49: 83–92.BR CullenL. Owens2002Experimental challenge and clinical cases of Bohle iridovirus (BIV) in native Australian anurans.Diseases of Aquatic Organisms498392
- 26. Cullen BR, Owens L, Whittington RJ (1995) Experimental infection of Australian anurans (Limnodynastes terraereginae and Litoria latopalmata) with Bohle iridovirus. Diseases of Aquatic Organisms 23: 83–92.BR CullenL. OwensRJ Whittington1995Experimental infection of Australian anurans (Limnodynastes terraereginae and Litoria latopalmata) with Bohle iridovirus.Diseases of Aquatic Organisms238392
- 27. Lang E, Gerhardt C, Davidson C (2009) The frogs and toads of North America. New York, New York, USA: Houghton Mifflin Company. E. LangC. GerhardtC. Davidson2009The frogs and toads of North AmericaNew York, New York, USAHoughton Mifflin Company
- 28. Miller DL, Rajeev S, Gray MJ, Baldwin CA (2007) Frog virus 3 infection, cultured American bullfrogs. Emerging Infectious Diseases 13: 342–343.DL MillerS. RajeevMJ GrayCA Baldwin2007Frog virus 3 infection, cultured American bullfrogs.Emerging Infectious Diseases13342343
- 29. Gosner KL (1960) A simplified table for staging anuran embryos and larvae with notes and identification. Herpetologica 16: 183–190.KL Gosner1960A simplified table for staging anuran embryos and larvae with notes and identification.Herpetologica16183190
- 30. Bollinger TK, Mao JH, Schock D, Brigham RM, Chinchar VG (1999) Pathology, isolation, and preliminary molecular characterization of a novel iridovirus from tiger salamanders in Saskatchewan. Journal of Wildlife Diseases 35: 413–429.TK BollingerJH MaoD. SchockRM BrighamVG Chinchar1999Pathology, isolation, and preliminary molecular characterization of a novel iridovirus from tiger salamanders in Saskatchewan.Journal of Wildlife Diseases35413429
- 31. Brunner JL, Richards K, Collins JP (2005) Dose and host characteristics influence virulence of ranavirus infections. Oecologia 144: 399–406.JL BrunnerK. RichardsJP Collins2005Dose and host characteristics influence virulence of ranavirus infections.Oecologia144399406
- 32. Pearman PB, Garner TWJ (2005) Susceptibility of Italian agile frog populations to an emerging strain of ranavirus parallels population genetic diversity. Ecology Letters 8: 401–408.PB PearmanTWJ Garner2005Susceptibility of Italian agile frog populations to an emerging strain of ranavirus parallels population genetic diversity.Ecology Letters8401408
- 33. Rojas S, Richards K, Jancovich JK, Davidson EW (2005) Influence of temperature on Ranavirus infection in larval salamanders Ambystoma tigrinum. Diseases of Aquatic Organisms 63: 95–100.S. RojasK. RichardsJK JancovichEW Davidson2005Influence of temperature on Ranavirus infection in larval salamanders Ambystoma tigrinum.Diseases of Aquatic Organisms6395100
- 34. Relyea RA (2002) Competitor-induced plasticity in tadpoles: Consequences, cues, and connections to predator-induced plasticity. Ecological Monographs 72: 523–540.RA Relyea2002Competitor-induced plasticity in tadpoles: Consequences, cues, and connections to predator-induced plasticity.Ecological Monographs72523540
- 35. Thibaudeau G, Altig R (1999) Endotrophic anurans. In: McDiarmid RW, Altig R, editors. Tadpoles: the biology of anuran larvae. Chicago, Illinois, USA: University of Chicago Press. pp. 170–188.G. ThibaudeauR. Altig1999Endotrophic anurans.RW McDiarmidR. AltigTadpoles: the biology of anuran larvaeChicago, Illinois, USAUniversity of Chicago Press170188
- 36. Picco AM, Brunner JL, Collins JP (2007) Susceptibility of the endangered California tiger salamander, Ambystoma californiense, to ranavirus infection. Journal of Wildlife Diseases 43: 286–290.AM PiccoJL BrunnerJP Collins2007Susceptibility of the endangered California tiger salamander, Ambystoma californiense, to ranavirus infection.Journal of Wildlife Diseases43286290
- 37. Stokes ME, Davis CS, Kock GG (2000) Categorical data analysis using the SAS® system. Cary North Carolina, USA: SAS Institute. ME StokesCS DavisGG Kock2000Categorical data analysis using the SAS® systemCary North Carolina, USASAS Institute
- 38. Zar JH (1999) Biostatistical analysis. Upper Saddle River, New Jersey, USA: Prentice Hall. JH Zar1999Biostatistical analysisUpper Saddle River, New Jersey, USAPrentice Hall
- 39. Tweedell K, Granoff A (1968) Viruses and renal carcinoma of Rana pipiens. V. Effect of Frog Virus 3 on developing frog embryos and larvae. Journal of the National Cancer Institute 40: 407–410.K. TweedellA. Granoff1968Viruses and renal carcinoma of Rana pipiens. V. Effect of Frog Virus 3 on developing frog embryos and larvae.Journal of the National Cancer Institute40407410
- 40. Berrill M, Coulson DR, McGillivray L, Pauli BD (1998) Toxicity of endosulfan to aquatic stages of anuran amphibians. Environmental Toxicology and Chemistry 17: 1738–1744.M. BerrillDR CoulsonL. McGillivrayBD Pauli1998Toxicity of endosulfan to aquatic stages of anuran amphibians.Environmental Toxicology and Chemistry1717381744
- 41. Pauli BD, Coulson DR, Berrill M (1999) Sensitivity of amphibian embryos and tadpoles to MIMIC 240LV insecticide following single or double exposures. Environmental Toxicology and Chemistry 18: 2538–2544.BD PauliDR CoulsonM. Berrill1999Sensitivity of amphibian embryos and tadpoles to MIMIC 240LV insecticide following single or double exposures.Environmental Toxicology and Chemistry1825382544
- 42. Han Y, Yu H, Yang X, Rees HH, Liu J, et al. (2008) A serine proteinase inhibitor from frog eggs with bacteriostatic activity. Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology 149: 58–62.Y. HanH. YuX. YangHH ReesJ. Liu2008A serine proteinase inhibitor from frog eggs with bacteriostatic activity.Comparative Biochemistry and Physiology B-Biochemistry & Molecular Biology1495862
- 43. Green DE, Converse KA (2005) Diseases of frogs and toads. In: Majumdar SK, Huffman JE, Brenner FJ, Panah AI, editors. Wildlife diseases: Landscape epidemiology, spatial distribution and utilization of remote sensing technology. Easton, Pennsylvania: The Pennsylvania Academy of Science. pp. 89–117.DE GreenKA Converse2005Diseases of frogs and toads.SK MajumdarJE HuffmanFJ BrennerAI PanahWildlife diseases: Landscape epidemiology, spatial distribution and utilization of remote sensing technologyEaston, PennsylvaniaThe Pennsylvania Academy of Science89117
- 44. Greer AL, Berrill M, Wilson PJ (2005) Five amphibian mortality events associated with ranavirus infection in south central Ontario, Canada. Diseases of Aquatic Organisms 67: 9–14.AL GreerM. BerrillPJ Wilson2005Five amphibian mortality events associated with ranavirus infection in south central Ontario, Canada.Diseases of Aquatic Organisms67914
- 45. Warne RW, Crespi EJ, Brunner JL (2011) Escape from the pond: stress and developmental responses to ranavirus infection in wood frog tadpoles. Functional Ecology 25: 139–146.RW WarneEJ CrespiJL Brunner2011Escape from the pond: stress and developmental responses to ranavirus infection in wood frog tadpoles.Functional Ecology25139146
- 46. Flajnik MF, Hsu E, Kaufman JF, Du Pasquier L (1987) Changes in the immune system during metamorphosis of Xenopus. Immunology Today 8: 58–64.MF FlajnikE. HsuJF KaufmanL. Du Pasquier1987Changes in the immune system during metamorphosis of Xenopus.Immunology Today85864
- 47. Rollins-Smith LA (2001) Neuroendocrine-immune system interactions in amphibians - Implications for understanding global amphibian declines. Immunologic Research 23: 273–280.LA Rollins-Smith2001Neuroendocrine-immune system interactions in amphibians - Implications for understanding global amphibian declines.Immunologic Research23273280
- 48. Rollins-Smith LA, Blair PJ (1993) The effects of corticosteroid hormones and thyroid hormones on lymphocyte viability and proliferation during development and metamorphosis of Xenopus laevis. Differentiation 54: 155–160.LA Rollins-SmithPJ Blair1993The effects of corticosteroid hormones and thyroid hormones on lymphocyte viability and proliferation during development and metamorphosis of Xenopus laevis.Differentiation54155160
- 49. Grant PR, Clothier H, Johnson RO, Ruben LN (1998) In situ lymphocyte apoptosis in larval Xenopus laevis, the South African clawed toad. Developmental and Comparative Immunology 22: 449–455.PR GrantH. ClothierRO JohnsonLN Ruben1998In situ lymphocyte apoptosis in larval Xenopus laevis, the South African clawed toad.Developmental and Comparative Immunology22449455
- 50. Pallister JA, Halliday DC, Robinson AJ, Venables D, Voysey RD, et al. (2011) Assessment of Virally Vectored Autoimmunity as a Biocontrol Strategy for Cane Toads. PLoS ONE 6(1): e14576.JA PallisterDC HallidayAJ RobinsonD. VenablesRD Voysey2011Assessment of Virally Vectored Autoimmunity as a Biocontrol Strategy for Cane Toads.PLoS ONE61e14576
- 51. Newman RA (1992) Adaptive plasticity in amphibian metamorphosis. Bioscience 42: 671–678.RA Newman1992Adaptive plasticity in amphibian metamorphosis.Bioscience42671678
- 52. Denver RJ (1997) Environmental stress as a developmental cue: Corticotropin-releasing hormone is a proximate mediator of adaptive phenotypic plasticity in amphibian metamorphosis. Hormones and Behavior 31: 169–179.RJ Denver1997Environmental stress as a developmental cue: Corticotropin-releasing hormone is a proximate mediator of adaptive phenotypic plasticity in amphibian metamorphosis.Hormones and Behavior31169179
- 53. Zettergren LD (2000) Ontogeny of B cells expressing IgM in embryonic and larval tissues of the American grass frog, Rana pipiens. Journal of Experimental Zoology 286: 737–744.LD Zettergren2000Ontogeny of B cells expressing IgM in embryonic and larval tissues of the American grass frog, Rana pipiens.Journal of Experimental Zoology286737744
- 54. Maniero GD, Morales H, Gantress J, Robert J (2006) Generation of a long-lasting, protective, and neutralizing antibody response to the ranavirus FV3 by the frog Xenopus. Developmental and Comparative Immunology 30: 649–657.GD ManieroH. MoralesJ. GantressJ. Robert2006Generation of a long-lasting, protective, and neutralizing antibody response to the ranavirus FV3 by the frog Xenopus.Developmental and Comparative Immunology30649657
- 55. Morales HD, Abramowitz L, Gertz J, Sowa J, Vogel A, et al. (2010) Innate immune responses and permissiveness to ranavirus infection of peritoneal leukocytes in the frog Xenopus laevis. American Society of Microbiology 84: 4912–4922.HD MoralesL. AbramowitzJ. GertzJ. SowaA. Vogel2010Innate immune responses and permissiveness to ranavirus infection of peritoneal leukocytes in the frog Xenopus laevis.American Society of Microbiology8449124922
- 56. Harp EM, Petranka JW (2006) Ranavirus in wood frogs (Rana sylvatica): Potential sources of transmission within and between ponds. Journal of Wildlife Diseases 42: 307–318.EM HarpJW Petranka2006Ranavirus in wood frogs (Rana sylvatica): Potential sources of transmission within and between ponds.Journal of Wildlife Diseases42307318
- 57. Gahl MK, Calhoun AJK (2010) The role of multiple stressors in ranavirus-caused amphibian mortalities in Acadia National Park wetlands. Canadian Journal of Zoology 88: 108–121.MK GahlAJK Calhoun2010The role of multiple stressors in ranavirus-caused amphibian mortalities in Acadia National Park wetlands.Canadian Journal of Zoology88108121
- 58. Cotter JD, Storfer A, Page RB, Beachy CK, Voss SR (2008) Transcriptional response of Mexican axolotls to Ambystoma tigrinum virus (ATV) infection. BMC Genomics 9: 493.JD CotterA. StorferRB PageCK BeachySR Voss2008Transcriptional response of Mexican axolotls to Ambystoma tigrinum virus (ATV) infection.BMC Genomics9493
- 59. Haimovich J, Du Pasquier L (1973) Specificity of antibodies in amphibian larvae possessing a small number of lymphocytes. Proceeding of the National Academy of Science (USA) 70: 1898–1902.J. HaimovichL. Du Pasquier1973Specificity of antibodies in amphibian larvae possessing a small number of lymphocytes.Proceeding of the National Academy of Science (USA)7018981902
- 60. Hsu E, Du Pasquier L (1984) Ontogenyof the immune system in Xenopus. II. Antibody repertoire differences between larvae and adults. Differentiation 28: 116–122.E. HsuL. Du Pasquier1984Ontogenyof the immune system in Xenopus. II. Antibody repertoire differences between larvae and adults.Differentiation28116122
- 61. Robert J, Abramowitz L, Gantress J, Morales HD (2007) Xenopus laevis: A possible vector of ranavirus infection? Journal of Wildlife Diseases 43: 645–652.J. RobertL. AbramowitzJ. GantressHD Morales2007Xenopus laevis: A possible vector of ranavirus infection?Journal of Wildlife Diseases43645652
- 62. Richmond JQ, Savage AE, Zamudio KR, Rosenblum EB (2009) Toward immunogenetic studies of amphibian chytridiomycosis: Linking innate and acquired immunity. Bioscience 59: 311–320.JQ RichmondAE SavageKR ZamudioEB Rosenblum2009Toward immunogenetic studies of amphibian chytridiomycosis: Linking innate and acquired immunity.Bioscience59311320
- 63. ter Braak CJF (1986) Canonical correspondence analysis: a new eigenvector technique for multivariate direct gradient analysis. Ecology 67: 1167–1179.CJF ter Braak1986Canonical correspondence analysis: a new eigenvector technique for multivariate direct gradient analysis.Ecology6711671179
- 64. Allen LJS (2006) An Introduction to Mathematical Biology. Upper Saddle River, New Jersey: Prentice Hall. LJS Allen2006An Introduction to Mathematical BiologyUpper Saddle River, New JerseyPrentice Hall