The emerging infectious disease chytridiomycosis is driven by the spread of amphibian chytrid fungus (Batrachochytrium dendrobatidis, Bd), a highly virulent pathogen threatening global amphibian biodiversity. Although pandemic in distribution, previous intensive field surveys have failed to detect Bd in Madagascar, a biodiversity hotspot home to hundreds of endemic amphibian species. Due to the presence of Bd in nearby continental Africa and the ecological crisis that can be expected following establishment in Madagascar, enhanced surveillance is imperative. I sampled 565 amphibians commercially exported from Madagascar for the presence of Bd upon importation to the USA, both to assist early detection efforts and demonstrate the conservation potential of wildlife trade disease surveillance. Bd was detected in three animals via quantitative PCR: a single Heterixalus alboguttatus, Heterixalus betsileo, and Scaphiophryne spinosa. This is the first time Bd has been confirmed in amphibians from Madagascar and presents an urgent call to action. Our early identification of pathogen presence prior to widespread infection provides the necessary tools and encouragement to catalyze a swift, targeted response to isolate and eradicate Bd from Madagascar. If implemented before establishment occurs, an otherwise likely catastrophic decline in amphibian biodiversity may be prevented.
Citation: Kolby JE (2014) Presence of the Amphibian Chytrid Fungus Batrachochytrium dendrobatidis in Native Amphibians Exported from Madagascar. PLoS ONE 9(3): e89660. https://doi.org/10.1371/journal.pone.0089660
Editor: Matthew (Mat) Charles Fisher, Imperial College Faculty of Medicine, United Kingdom
Received: October 27, 2013; Accepted: January 17, 2014; Published: March 5, 2014
Copyright: © 2014 Jonathan E. Kolby. 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: Jonathan Kolby was supported by a James Cook University Australian Postgraduate Research Scholarship. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The author has declared that no competing interests exist.
Amphibian populations are experiencing global decline in response to a storm of assaults including habitat destruction, climate change, and the emerging infectious disease chytridiomycosis caused by amphibian chytrid fungus, Batrachochytrium dendrobatidis (Bd) –. Bd demonstrates low host species specificity and can potentially affect the entire class Amphibia, threatening the survival of thousands of amphibian species . This pathogen can be highly lethal and easily transmissible through direct physical contact with affected individuals or indirectly by exposure to water contaminated with aquatic Bd zoospores . Despite infection, certain species can act as reservoir hosts, allowing Bd to persist while driving others to extinction. This, together with prolonged environmental persistence provides an optimal situation for pathogen establishment and the collapse of amphibian diversity, especially in aquatic environments –.
Bd was first identified and described nearly 15 years ago , , by which time it had already spread to dozens of countries, potentially through the international trade in live amphibians –. Annually, millions of live amphibians are traded globally for the exotic pet trade, biomedical research, and human consumption and this movement of potentially infected animals may be a primary driving force of global Bd dispersal –. The transportation of Bd-contaminated environmental substrates and field equipment represent additional potential dispersal pathways , , suggesting that common activities such as freshwater aquaculture and mining may also contribute towards the spread of Bd even in the absence of amphibian movement.
Although the spread of Bd has continued seemingly unabated for many decades, there remain hotspots of amphibian biodiversity where this devastating pathogen is not yet established and has been presumed absent due to the lack of confirmed field detection, most notably in Madagascar. The first expansive survey for the presence of Bd in Madagascar failed to detect this pathogen in 527 amphibians from 79 species sampled from 2005–2006 . To complement this effort, a follow-up survey of 300 animals from 53 species at 12 additional locations were sampled in 2006 and 2007 , and a further 56 amphibians from 12 species were sampled in the country's central highlands ; all results similarly demonstrated the absence of Bd in amphibians sampled despite covering a range of host species and environments, and employing the most sensitive diagnostic tool, the Bd-specific quantitative PCR (qPCR) assay. It is remarkable that Bd is not already widespread in Madagascar because the country possesses high diversity of amphibians likely to be susceptible to chytridiomycosis, is in close proximity to regions of Bd presence in continental Africa (i.e. Tanzania, Malawi, South Africa), and provides high environmental suitability for Bd , .
Thousands of amphibians are exported annually from Madagascar and disseminated globally into the exotic pet trade. An analysis of records obtained from the United States Fish and Wildlife Service (USFWS) through a Freedom of Information Act (FOIA) request shows 39,020 amphibians were exported from Madagascar to the United States between 2006 and 2011, from at least 31 species. Although the international movement of amphibians is believed to help spread Bd and thus jeopardize global animal health, we considered access to traded animals a boon to our research goals: demonstration that trade can be approached as an efficient wildlife disease surveillance tool for the rapid detection of Bd in Madagascar. The commercial trade in amphibians generates large and diverse sample pools from which proactive surveillance can be performed with less human and financial resources than conventional field surveys. This investigation explored the presence of Bd in Madagascar by examining the contents of a shipment of wild-collected endemic amphibians exported directly to the USA.
In total, 565 amphibians of nine species exported from Madagascar were sampled for Bd detection (Table 1). Bd was detected in three of 565 animals and each displayed measureable amounts of Bd in at least two qPCR replicates, from at least two separate plates. The three species positive for Bd were Scaphiophryne spinosa (MGSS30), Heterixalus alboguttatus (MGHA54), and Heterixalus betsileo (MGHB42) (Table 1). Prior to its final qPCR with purified DNA, MGSS30 was tested in two separate qPCR plates and in each, one replicate came up positive; the zoospore loads were 0.332 and 0.040, respectively. When the purified DNA was run a final time, all three replicates of MGSS30 were negative for Bd. MGHA54 was tested in two plates prior to DNA purification and again, one replicate per plate came up positive for Bd, with zoospore loads of 0.189 and 0.089. After DNA purification, one replicate was again positive; its reported zoospore load was 0.400. MGHB42 was also tested in two plates prior to purification and unlike the other two samples, all six replicates were positive for Bd; the average zoospore load for the first plate was 0.395 and the average zoospore load in the second plate was 0.219. After DNA purification, all three MGHB42 replicates were again positive for Bd, reporting a mean value of 1.059 zoospores.
Examination of amphibians sampled for Bd collectively revealed ulcerations (1.8%), heavy skin sloughing (3.4%), and death on arrival (6.9%) in 68/565 animals (Table 1). Because not all deceased amphibians were sampled for Bd, the total number of DOA animals was greater with respect to the entire shipment (n = 99; 15.8%). No such conditions were observed in any of the three Bd-positive amphibians at the time of sampling.
The presence of Bd has been confirmed in Malagasy amphibians for the first time. These amphibians were collected from the wild for the pet trade, exported to the USA and sampled immediately upon arrival. One sample produced a strong signal for Bd presence (MGHB42), and two others displayed weak indications: MGSS30 and MGHA54. Despite the low intensities, these two samples certainly displayed positive signals and, most important from separate plates, suggesting the signals were real and not due to contamination from the positive controls. It is not uncommon for the standard controls to contaminate a single replicate, but to do so across multiple plates, has never been witnessed and is unlikely. After DNA purification, all three MGSS30 replicates were negative for Bd. This is perplexing and could suggest that the original DNA aliquot used in the first two plates (prior to purification) was contaminated. However, it is also possible that the particular aliquot of DNA used in the final run did not actually contain Bd DNA, although it existed in the sample; MGSS30's measured zoospore loads were incredibly low and give some credence to this possibility. MGHA54, like MGSS30, similarly never had all replicates within a single plate turn up positive. Its zoospore load was similarly low, again suggesting that Bd DNA similarly might not have been present in each replicate. Because a single replicate was positive from three different plates, including the final qPCR using purified DNA, these data do suggest MGHA54 was positive for Bd. Although it is difficult to discern the truth about MGSS30 and MGHA54, because separate plates yielded positive replicates and contamination was unlikely, I report these samples as Bd-positive. Regardless, all nine replicates of MGHB42 were positive for Bd, undeniably confirming its presence in material from Madagascar.
The status of Bd in wild amphibian populations in Madagascar remains uncertain and calls for urgent targeted field surveys in regions where these Bd-positive animals were likely collected. The human-assisted movement of traded animals introduces an opportunity for Bd cross-contamination between species and collection origins prior to exportation if animals are housed in shared enclosures where direct or indirect contact is allowed. Accordingly, transmission of Bd between Malagasy species from different collection localities may potentially exaggerate the number of affected species in wild populations and suggested distributional range of infection.. Furthermore, identifying the source of Bd detected in traded animals becomes especially challenging when animals from multiple countries are also present in the trade sector. Fortunately, this is not the case in Madagascar; commercial amphibian importation does not occur in the country  and only those of national origin are traded. The absence of foreign-sourced amphibian species suggests my detection of Bd is not simply an artifact of re-exportation through the amphibian trade, but instead a reflection of Bd presence in the wild in Madagascar. Still, other non-amphibian wildlife trade activities may unknowingly introduce foreign infectious material to Madagascar and expose wild-collected frogs prior to exportation if housed at a shared facility (i.e. exposure to Bd-contaminated water accompanying freshwater fish importations). Albeit unlikely the result of such cross contamination, I employed a conservative approach by interpreting these data as confirmation of Bd presence in Madagascar within the amphibian trade, but not yet irrefutable evidence for Bd presence in wild amphibian populations, despite the strong suggestion.
A second, more specific tier of surveillance via targeted field sampling applying this new information, is now imperative to determine the current extent of Bd in Madagascar outside the trade sector. A predictive model of Bd distribution  shows that the highest climatic suitability for Bd overlaps particularly closely with the distributional range of H. betsileo, from which MGHB42 was collected. Interestingly, the distributions of H. albuguttatus and S. spinosa fall on the periphery of this climatic range and may have been collected from areas with moderate to low Bd suitability, potentially explaining their exceptionally low Bd zoospore loads compared to that detected in the specimen of H. betsileo. Accordingly, surveys to trace back the source of the Bd detected herein should commence immediately within the distributional ranges of H. betsileo, H. alboguttatus, and S. spinosa, target the larvae and subadults expected to exhibit increased susceptibility to infection, and include bioregions suitable for Bd  to maximize the chances of rapidly detecting Bd if currently present in wild populations.
The lack of Bd detection in previous field surveys of wild amphibians – and these newly reported Bd-positive animals suggest the presence of Bd in Madagascar is a recent phenomenon and not yet widespread. Bd-related die-offs have not been documented and infection prevalence is expected to still be extremely low in wild populations, if currently affected. The conditions necessary to result in Bd establishment in amphibian populations following exposure are poorly understood. Infection with as little as one Bd zoospore can result in chytridiomycosis , and affected amphibians have been observed to release 68 infectious Bd particles per minute when in an aquatic environment . Therefore, the detection of Bd in H. betsileo and H. alboguttatus is especially concerning because these species breed in both permanent and temporary water bodies and an outbreak in wild populations may both promote extended environmental persistence and facilitate indirect transmission to nearby aquatic species, increasing the opportunity for pathogen establishment. The spread of Bd can occur rapidly following introduction to a naive region, estimated as much as 25–282 km/y , and the data presented herein provides impetus to quickly reevaluate the presence of Bd in Madagascar.
The confirmation of Bd in amphibians exported from Madagascar presents an opportunity to intervene prior to the first confirmed outbreak in wild populations – an outbreak with potentially irreparable ecological consequences. It is no longer questionable whether or not Bd will become introduced to Madagascar; it is now a tangible threat. Survival of the country's amphibians now requires an efficient network of proactive surveillance and rapid response to quickly identify additional introduction events and minimize exposure to wild populations , because pathogen eradication is considered implausible following establishment. The provenance of Bd detected in this investigation remains an enigma, especially considering the absence of commercial amphibian trade into Madagascar, suggesting a more insidious mechanism is responsible for the introduction. Accordingly, Bd may continue to arrive in Madagascar and creep closer towards establishment until the true introduction pathway is identified, targeted and controlled. Early detection now provides the opportunity to interrupt pathogen establishment, but if not acted upon with haste, disease-associated ecological decline in Madagascar may soon become inescapable.
Materials and Methods
Amphibians were imported under United States Fish and Wildlife Service (USFWS) License No. LE65317A-0 and accompanied by a cleared USFWS Declaration for Importation of Wildlife (Form 3-177). None of the species included in this investigation are currently protected or endangered and therefore, no additional special permits were necessary. Permission to export the amphibians was granted by the Government of Madagascar with permit #'s 017/12-MEF/SG/DREF.ATS/EXPORT and 018/11-MEF/SG/DREF.ATS.
In February 2012, a shipment containing 565 wild-collected amphibians from Madagascar was exported and sampled for the presence of Bd upon importation to the United States. Of 17 endemic amphibian species commercially available from this particular supplier, nine were systematically selected for sampling to represent a potentially wide coverage of biogeographical regions and altitudinal ranges where Bd was expected to thrive if present. These decisions were made by comparing species distribution maps provided by the IUCN Red List of Threatened Species  with work that identified a predicted region of optimal Bd survival based on climatic suitability . Sampling priority was accordingly directed towards species with distributions that overlap this high risk zone, in addition to those vulnerable to Bd exposure based on life history characteristics, most importantly association with aquatic habitats .
Following regulatory clearance for importation into the USA, the shipment was collected from the airport and immediately transported to a small greenhouse specifically constructed to provide a controlled area for receipt and sampling of these animals. This structure had no previous exposure to amphibians and all interior surfaces were first washed with a 10% bleach solution to minimize any potential risk of domestic Bd contamination. The shipment remained sealed for the duration of transport from Madagascar to the USA, and was not opened until first secured inside this sampling location to further prevent opportunities for contamination. All contents of the shipment were handled exclusively with fresh pairs of Nitrile gloves.
Amphibians arrived inside two wooden crates insulated with 1/4″ Styrofoam (Fig. 1). Within these crates, the amphibians were packed in plastic containers filled with damp sphagnum moss and leaves as bedding material. Some containers housed multiple amphibians whereas others were packed individually; this varied by species and size of amphibian, but containers housing multiple individuals did not combine species. All amphibians were adults, with the exception of S. spinosa, for which only subadult frogs were received. Upon opening the crates, containers were arranged by species and the contents of each examined and sampled for the presence of Bd. A sterile fine-tipped rayon swab (Medical Wire & Equipment Co., MW113) was drawn across each amphibian's hands, feet, and pelvic patch five times each. Samples were stored in 2 mL vials filled with 1 mL 70% ethanol as preservative. To prevent cross-contamination between samples, fresh pairs of Nitrile gloves were changed each time a new amphibian was handled.
Amphibians were shipped sealed in wooden crates, insulated with 1/4″ Styrofoam, and packed in plastic containers filled with damp sphagnum moss and leaves.
Each animal was examined immediately prior to swabbing and its condition recorded. Potential clinical symptoms of chytridiomycosis were noted, including the presence of skin ulcerations, skin sloughing, and death . Most specimens of the nine target species were sampled, except for dead animals that arrived in advanced stages of decomposition (n = 60), which were excluded from this investigation. All live amphibians were swabbed individually. When multiple dead animals arrived in the same container, a single swab was used to sample all carcasses; this maneuver increases cost-efficiency of analysis, resulting in fewer swabs (n = 551) than the total number of animals actually sampled (n = 565). Following the prompt completion of sample collection for this investigation, all live amphibians were transferred back into the course of the domestic pet trade.
Thirty swabs deemed as high priority, those most suspect of Bd infection based on physical examination, were first immediately shipped to the San Diego Zoo Amphibian Disease Laboratory for testing. Samples were processed via a sensitive quantitative PCR assay (qPCR) specific to Bd following standard methods , . Assays were run on an Applied Biosystems 7900HT thermocycler using 384 well plates with Applied Biosystems exogenous internal positive control labeled with Vic in separate wells to test for the presence of PCR inhibitors. For each sample, 5 µl of 1∶10 dilution of swab DNA was added to each well for a final total qPCR volume of 20 µl. Standard curves were generated with 10-fold serial dilutions (range 10,000 to 0.001 zoospores) of laboratory cultivated Bd zoospores.
The remaining 521 swabs were also processed via qPCR according to established protocols – at Yale University. Samples were extracted with 150 µl Prepman Ultra (Applied Biosystems, California, USA), with a final 30 µl of supernatant removed for downstream use. An aliquot of this supernatant was diluted 1∶10 in DNase-free water for qPCR. The qPCR protocol used SensiMix II Low Rox (Bioline, Massachusetts, USA) as the qPCR master mix . Samples and controls were run in triplicate with three positive, standard control samples (100, 10, and 1 zoospore/well, made from JAM81 pure culture; see Boyle et al. 2004 for standard control construction) and one non-template control (DNase free, molecular-grade water). When the qPCR assay failed to detect Bd in all replicate wells, the sample was deemed negative for Bd. When one of three replicates successfully detected Bd, the sample was rerun (in triplicate again) in a subsequent plate. For rerun samples that had at least a total of two of six replicates positive for Bd (from at least two separate plates) or samples that had Bd in all replicates, the original DNA supernatant stock was purified and tested, at full-strength, in a final qPCR. Full-strength DNA from PrepMan Ultra inhibits qPCR, so this DNA must be cleaned-up or diluted prior to use . The remaining full-strength DNA was purified using Performa DTR Gel Filtration Cartridges (Edge Biosystems, Maryland, USA). The cartridges were loaded and prepared by spinning at 750× g for two minutes. The remaining DNA was added to each column, loaded directly onto the gel matrix, and then spun for two minutes at 750× g. Five microliters of this eluted, purified, full-strength DNA was loaded into three replicate wells in a final qPCR (i.e., this DNA was not further diluted prior to qPCR). All zoospore loads described in this report have not been converted; here, reported zoospore loads come from 5 µl DNA (1∶10 or full-strength), placed into 20 µl reaction volumes.
I thank Cheryl Jacko for assisting with sample collection and both Kathryn Richards-Hrdlicka and Dana Lee for performing the sample extractions and qPCRs. Kathryn Richards-Hrdlicka also provided data interpretation. I am grateful for the helpful comments made on this manuscript by Lee Skerratt, Lee Berger, Laura Brannelly, and Kathryn Richards-Hrdlicka.
Conceived and designed the experiments: JEK. Performed the experiments: JEK. Analyzed the data: JEK. Wrote the paper: JEK.
- 1. Stuart SN, Chanson JS, Cox NA, Young BE, Rodrigues ASL, et al. (2004) Status and Trends of Amphibian Declines and Extinctions Worldwide. Science 306: 1783–1786.
- 2. Pounds AJ, Bustamante MR, Coloma LA, Consuegra JA, Fogden MPL, et al. (2006) Widespread amphibian extinctions from epidemic disease driven by global warming. Nature 439: 161–167.
- 3. Skerratt L, Berger L, Speare R, Cashins S, McDonald K, et al. (2007) Spread of Chytridiomycosis Has Caused the Rapid Global Decline and Extinction of Frogs. EcoHealth 4: 125–134.
- 4. Daszak P, Berger L, Cunningham AA, Hyatt AD, Green DE, et al. (1999) Emerging infectious diseases and amphibian population declines. Emerg Infect Dis 5: 735–748.
- 5. Carey C, Bruzgul J, Livo L, Walling M, Kuehl K, et al. (2006) Experimental Exposures of Boreal Toads (Bufo boreas) to a Pathogenic Chytrid Fungus (Batrachochytrium dendrobatidis). EcoHealth 3: 5–21.
- 6. Johnson ML, Speare R (2003) Survival of Batrachochytrium dendrobatidis in water: quarantine and disease control implications. Emerg Infect Dis 9: 922–925.
- 7. Daszak P, Strieby A, Cunningham AA, Longcore JE, Brown CC, et al. (2004) Experimental evidence that the bullfrog (Rana catesbeiana) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Herpetological Journal 14: 201–207.
- 8. Fisher MC, Garner TW, Walker SF (2009) Global emergence of Batrachochytrium dendrobatidis and amphibian chytridiomycosis in space, time, and host. Annu Rev Microbiol 63: 291–310.
- 9. Berger L, Speare R, Daszak P, Green DE, Cunningham AA, et al. (1998) Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proceedings of the National Academy of Sciences of the United States of America 95: 9031–9036.
- 10. Longcore JE, Pessier AP, Nichols DK (1999) Batrachochytrium dendrobatidis gen. et sp. nov., a Chytrid Pathogenic to Amphibians. Mycologia 91: 219–227.
- 11. Weldon C, du Preez LH, Hyatt AD, Muller R, Spears R (2004) Origin of the amphibian chytrid fungus. Emerg Infect Dis 10: 2100–2105.
- 12. Fisher MC, Garner TWJ (2007) The relationship between the emergence of Batrachochytrium dendrobatidis, the international trade in amphibians and introduced amphibian species. Fungal Biology Reviews 21: 2–9.
- 13. Schloegel LM, Toledo LF, Longcore JE, Greenspan SE, Vieira CA, et al. (2012) Novel, panzootic and hybrid genotypes of amphibian chytridiomycosis associated with the bullfrog trade. Mol Ecol 21: 5162–5177.
- 14. Schloegel LM, Picco AM, Kilpatrick AM, Davies AJ, Hyatt AD, et al. (2009) Magnitude of the US trade in amphibians and presence of Batrachochytrium dendrobatidis and ranavirus infection in imported North American bullfrogs (Rana catesbeiana). Biological Conservation 142: 1420–1426.
- 15. Farrer RA, Weinert LA, Bielby J, Garner TWJ, Balloux F, et al. (2011) Multiple emergences of genetically diverse amphibian-infecting chytrids include a globalized hypervirulent recombinant lineage. Proceedings of the National Academy of Sciences of the United States of America 108: 18732–6.
- 16. Webb R, Mendez D, Berger L, Speare R (2007) Additional disinfectants effective against the amphibian chytrid fungus Batrachochytrium dendrobatidis. Diseases of aquatic organisms 74: 13–16.
- 17. Weldon C, du Preez L, Vences M (2008) Lack of detection of the amphibian chytrid fungus (Batrachochytrium dendrobatidis) in Madagascar. In: Andreone F, editor A Conservation Strategy for the Amphibians of Madagascar Monografie XLV Torino, Museo Regionale di Scienze Naturali: 95–106.
- 18. Vredenburg VT, du Preez L, Raharivololoniaina L, Vieites DR, Vences M, et al. (2012) A molecular survey across Madagascar does not yield positive records of the amphibian chytrid fungus Batrachochytrium dendrobatidis. Herpetol Notes 5: 507–517.
- 19. Crottini AM, Barbuto M, Casiraghi M, Andreone F (2011) A rapid amphibian survey at Itremo-Ambatofinandrahana, central Madagascar, with confirmed absence of chytrid fungus and recommendations for future monitoring activities. North-Western Journal of Zoology 7: 346–351.
- 20. Lötters S, Rödder D, Kielgast J, Glaw F (2011) Hotspots, Conservation, and Diseases: Madagascar's Megadiverse Amphibians and the Potential Impact of Chytridiomycosis. In: Zachos FE, Habel JC, editors. Biodiversity Hotspots: Springer Berlin Heidelberg. pp. 255–274.
- 21. Olson DH, Aanensen DM, Ronnenberg KL, Powell CI, Walker SF, et al. (2013) Mapping the global emergence of Batrachochytrium dendrobatidis, the amphibian chytrid fungus. PloS ONE 8: e56802.
- 22. Skerratt LF, Berger L, Hines HB, McDonald KR, Mendez D, et al. (2008) Survey protocol for detecting chytridiomycosis in all Australian frog populations. Diseases of Aquatic Organisms 80: 85–94.
- 23. Reeder NM, Pessier AP, Vredenburg VT (2012) A reservoir species for the emerging amphibian pathogen Batrachochytrium dendrobatidis thrives in a landscape decimated by disease. PloS ONE 7: e33567.
- 24. Lips KR, Diffendorfer J, Mendelson JR, Sears MW (2008) Riding the wave: reconciling the roles of disease and climate change in amphibian declines. PLoS Biology 6: e72.
- 25. Andreone F, Carpenter AI, Cox N, du Preez L, Freeman K, et al. (2008) The challenge of conserving amphibian megadiversity in Madagascar. PLoS Biology 6: e118.
- 26. The IUCN Red List of Threatened Species. Version 2012.1. Available: http://www.iucnredlist.org. Accessed 2012 Jan 4.
- 27. Nichols DK, Lamirande EW, Pessier AP, Longcore JE (2001) Experimental transmission of cutaneous chytridiomycosis in dendrobatid frogs. J Wildl Dis 37: 1–11.
- 28. Boyle DG, Hyatt AD, Daszak P, Berger L, Longcore JE, et al. (2003) Cryo-archiving of Batrachochytrium dendrobatidis and other chytridiomycetes. Diseases of Aquatic Organisms 56: 59–64.
- 29. Hyatt AD, Boyle DG, Olsen V, Boyle DB, Berger L, et al. (2007) Diagnostic assays and sampling protocols for the detection of Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms 73: 175–192.
- 30. Boyle DG, Boyle DB, Olsen V, Morgan JA, Hyatt AD (2004) Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Diseases of Aquatic Organisms 60: 141–148.
- 31. Garland S, Baker A, Phillott AD, Skerratt LF (2010) BSA reduces inhibition in a TaqMan assay for the detection of Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms 92: 113–116.
- 32. Richards-Hrdlicka KL, Richardson JL, Mohabir L (2013) First survey for the amphibian chytrid fungus Batrachochytrium dendrobatidis in Connecticut (USA) finds widespread prevalence. Diseases of Aquatic Organisms 102: 169–180.