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Amphibian chytrid in Fact sheet

Introductory statement

Batrachochytrium dendrobatidis (B. dendrobatidis or Bd) (Longcore et al. 1999), the chytrid fungus, is a pathogen capable of driving to extinction. In Australia, the fungus has been directly implicated in the extinction of at least four species and the dramatic decline of at least 10 others.

There are many lessons to be learnt from chytrid and its global spread including: the importance of foresighting and horizon scanning in Australia’s wildlife biosecurity framework; the need for targeted, risk- based surveillance of selected Australian wildlife taxa, and; prior to an incursion the production and agreement on coordinated response plans that clearly articulate areas of responsibility and action.

Aetiology

Batrachochytrium dendrobatidis, the amphibian chytrid fungus. Fungi, phylum , Order Chytridiales. Until recently it was thought to be the only member in the . However, a second species, Batrachochytrium salamandrivorans, was discovered in the Netherlands in 2013 (Martel et al 2013).

Natural hosts

Amphibians (Berger et al. 1998). Currently found in all three extant amphibian orders: Anura ( and toads), Caudata ( and newts) and Gymnophiona (caecilians). The fungus has been found in 52 of 82 countries sampled and 516 of 1240 amphibian species examined. It is currently known from at least 449 anuran, 59 and six caecilian species worldwide (Doherty-Bone et al 2013, Gower et al 2013, Olsonet al 2013) but this number will rise as search effort and reporting continues.

No sex-linked predisposition. No age-linked predisposition to , since are commonly infected, but there is age-linked mortality. Adults and juveniles die from chytridiomycosis while tadpoles have not been reported to die from chytridiomycosis. Mortality in susceptible species is in general higher in metamorphs than adults. Recent work has also shown that the fungus can live within the gastrointestinal tracts of several species in the US: the blue crayfish ( alleni), red swamp crayfish (Procambarus clarkii) and northern crayfish (Orconectes virilis) (McMahon et al 2013).

World distribution

Known from all continents where occur. - Botswana, Ghana, Kenya, Lesotho, Morocco, South Africa, , Zambia; Australia; Pacific - , Hawaii; - Costa Rica, Cuba, , Honduras, Mexico, , Puerto Rico; - Canada, USA; - Argentina, Brazil, Ecuador, Peru, Uruguay, ; - Denmark, France, Germany, Hungary, Italy, Portugal, Spain, Switzerland, United Kingdom; Asia – Indonesia, Japan, Philippines.

Occurrences in Australia

History

In Australia, the oldest record of B. dendrobatidis is from a museum specimen collected in south-east Queensland near Brisbane in 1978 (Department of the Environment and Heritage 2006), which coincides with sudden frog declines in a number of species and two species extinctions in the region (Berger et al. 1998; Hines et al. 1999). Subsequent amphibian declines in central coastal Queensland (1985-86) and the Wet Tropics (1990-95) suggest that B. dendrobatidis spread north to its current northern limit at Big Tableland near Cooktown (Laurance et al. 1996; Berger et al. 1999a; Skerratt et al. 2010). In southern Australia, the spread of B. dendrobatidis is poorly documented but its distribution extends down the entire east coast to Tasmania (first detected in 2004) (Obendorf & Dalton 2006; Pauza et al 2007). Two separate foci occur in other states, one in southwest Western Australia, where the earliest record dates to 1985, and another around Adelaide in South Australia (earliest record 1995) (Berger et al. 2004, Murray et al. 2011).

Current situation Chytridiomycosis was listed on the OIE Wildlife Diseases List in 2008 (World Organisation for Health 2014).

Sixty-three (29%) of Australia’s 219 endemic frogs have positive records for infection. Species from three endemic families (Hylidae, , ) and one introduced (Bufonidae) family are affected (Murray et al. 2011) (Tables 1 and 2).

Batrachochytrium dendrobatidis is now endemic in Queensland, New South Wales, Australian Capital Territory, Victoria, Tasmania and Western Australia. Little is known about B. dendrobatidis in South Australia. Much of the continent is considered too hot and/or dry to sustain Bd and as such it is not endemic in all Australia. It has been found in wild amphibian populations on the east coast of Queensland and New South Wales on or between the Great Dividing Range and the coast, in the Australian Capital Territory, Victoria, Tasmania and in southwest Western Australia. The Northern Territory is currently considered amphibian chytrid free (Skerratt et al. 2008).

Bd has been directly implicated in the extinction of at least four native frog species and the dramatic decline of at least ten others, including Litoria nannotis (waterfall frog), L. rheocola (common mist frog), L. spenceri (spotted ) and Nycitmystes dayi (lace-eyed tree frog). The four species listed as extinct are from Queensland and include Rheobatrachus silus (southern gastric-brooding frog, last seen 1981), Rheobatrachus

WHA Fact sheet: Amphibian chytrid fungus in Australia | May 2014 | 2 vitellinus (northern gastric-brooding frog, 1985), Taudactylus acutirostris (sharp-snouted day frog, 1997) and T. diurnus (southern day frog, 1979). Many persisting species remain at lower abundance and smaller distributions than the levels recorded before the species were affected by Bd, some are continuing to decline and significant mortality from the disease is ongoing even decades after its introduction (Australian Government Department of Environment, Unpublished).

Table 1. Number of Australian amphibian species found with chytridiomycosis by state compared with total species of amphibians and total species tested in each state. State Species +ve Species Species % of tested % of all species for Bd tested species +ve +ve ACT 1 2 18 50.0 5.6 NSW 20 25 84 80.0 23.8 NT 0 6 47 0.0 0.0 QLD 34 69 123 49.3 27.6 SA 4 6 27 66.7 14.8 TAS 2 4 10 50.0 20.0 VIC 4 8 33 50.0 12.1 WA 16 30 77 53.3 20.8 Australia 63 115 219 54.8 28.8 From Murray et al. 2011

Table 2. Amphibian hosts (n=63) reported with chytridiomycosis in Australia. Family Genus Species First reference Bufonidae Rhinella marina (introduced; Berger (2001) formerly Bufo marinus) Hylidae Litoria adelaidensis Aplin & Kirkpatrick (2000) Hylidae Litoria aurea Berger (2001) Hylidae Litoria barringtonensis Berger (2001) Hylidae Litoria booroolongensis Hunter unpub. data Hylidae Litoria burrowsi Obendorf & Nelson (2004) Hylidae Litoria caerulea Berger (2001) Hylidae Litoria chloris Berger (2001) Hylidae Litoria citropa Mahony (2000) dayi (formerly Hylidae Litoria Nyctimystes) Berger (2001) Hylidae Litoria ewingii Berger (2001) Hylidae Litoria fallax Kriger & Hero (2007) Hylidae Litoria genimaculata Berger (2001) Hylidae Litoria gracilenta Berger & Speare (2004) Hylidae Litoria infrafrenata Berger (2001) Hylidae Litoria jungguy Berger (2001) Hylidae Litoria latopalmata Kriger & Hero (2007) Hylidae Litoria lesueurii Berger (2001) Hylidae Litoria lorica Puschendorf et al (2009) Hylidae Litoria moorei Aplin & Kirkpatrick (2000)

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Family Genus Species First reference Hylidae Litoria nannotis Berger et al (1998) Hylidae Litoria nasuta Donovan et al (1999) Hylidae Litoria pearsoniana Berger (2001) Hylidae Litoria peronii Berger (2001) Hylidae Litoria phyllochroa Mahony (2000) Hylidae Litoria raniformis Norman & Waldman (2000) Hylidae Litoria rheocola Berger (2001) Hylidae Litoria spenceri Berger (2001) Hylidae Litoria tyleri Berger (2001) Hylidae Litoria verreauxii Berger (2001) Hylidae Litoria wilcoxii Berger (2001) Phillott & McDonald unpub. Hylidae Litoria xanthomera data Adelotus brevis Speare & Berger (2004) Limnodynastidae australiacus Berger (2001) Limnodynastidae Heleioporus barycragus Speare website Limnodynastidae Heleioporus eyrei Aplin & Kirkpatrick (2000) Limnodynastidae fletcheri Berger (2001) Limnodynastidae dorsalis Aplin & Kirkpatrick (2000) Limnodynastidae Berger (2001) Limnodynastidae Limnodynastes peronii Berger (2001) Limnodynastidae Limnodynastes tasmaniensis Berger (2001) Limnodynastidae Limnodynastes terraereginae Berger (2001) Limnodynastidae kunapalari Berger (2001) Limnodynastidae Neobatrachus pelobatoides Speare website Microhylidae Cophixalus ornatus Kriger (2006)* Myobatrachidae Assa darlingtoni Kriger & Hero (2007)* Myobatrachidae Crinia georgiana Aplin & Kirkpatrick (2000) Myobatrachidae Crinia glauerti Aplin & Kirkpatrick (2000) Myobatrachidae Crinia insignifera Aplin & Kirkpatrick (2000) Myobatrachidae Crinia pseudinsignifera Aplin & Kirkpatrick (2000) Myobatrachidae Crinia subinsignifera Aplin & Kirkpatrick (2000) Myobatrachidae Crinia tasmaniensis Pauza & Driessen (2008) Myobatrachidae Geocrinia rosea Aplin & Kirkpatrick (2000) Myobatrachidae Geocrinia vitellina Aplin & Kirkpatrick (2000) Myobatrachidae Mixophyes fasciolatus Berger (2001) Myobatrachidae Mixophyes fleayi Berger (2001) Myobatrachidae Mixophyes iteratus Mahony (2000) Myobatrachidae Pseudophryne corroboree Speare & Berger (2004) Myobatrachidae Pseudophryne pengilleyi Berger et al. (2004) Myobatrachidae Taudactylus acutirostris Berger (2001) Myobatrachidae Taudactylus eungellensis Retallick et al. (2004) Myobatrachidae Uperoleia fusca Kriger & Hero (2007) Myobatrachidae Uperoleia laevigata Berger (2001)

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*Positive by qPCR from a single individual only. The specificity of the test used on these individuals may be less than 100%. From Murray et al. 2011

Epidemiology

Morbidity rate: When frogs show clinical signs, death usually follows within 2-3 days. Prevalence of infection in apparently aclinical frogs in infected populations in Australia can approach 100%.

Mortality rate: In captivity and in challenge and transmission experiments, some adult frogs are capable of surviving and clearing but mortality rates of up to 100% are common. Pathogenicity varies with host species, fungal strain, exposure dose and period, temperature and body size. High mortality or susceptibility to infection observed in the laboratory may not always occur in the wild and vice versa. Recently metamorphosed frogs appear most sensitive to the disease in some species. Infections of tadpoles are limited to their keratinized mouthparts and often appear to have no negative effects (implicating them as potential disease reservoirs), although some evidence suggests that infected tadpoles of some species may lose body condition and suffer reduced survival. infections can be carried through metamorphosis and cause high metamorph and juvenile mortality. Presence and prevalence of Bd in the wild varies with species, life- stage, season, altitude and latitude and may be broadly governed by temperature.

It appears that amphibian populations develop some measure of resistance to the fungus over time, or the fungus becomes less pathogenic, or both, as frog populations have been identified that declined dramatically when the fungus first arrived but now appear to have stabilized and co-exist with the fungus (Retallick et al 2004).

Studies have also shown that individual frogs develop an acquired immune response allowing them to clear subsequent infections more rapidly (Richmond et al 2009).

Incubation period: time to clinical signs and death usually ranging between 14 and 70 days post-exposure

Transmission: Via zoospore, waterborne.

Sources of agent

• Shedding of zoospores from infected skin. Zoospores leave host via discharge papillae projecting through surface of epithelial cell. Zoospores require water to survive, although a film is adequate. • Skin on frogs and mouthparts on tadpoles are the only infective tissues • Zoospore invades stratum corneum to infect new host • Frogs are subclinical for the majority of the duration of the infection. B. dendrobatidis is not an obligate parasite and can exist and grow in moisture in the laboratory. However, it is easily outcompeted by environmental microorganisms. Hence, frogs can be infected from water containing zoospores generated either from frogs or potentially from non-parasitic growth that might occur under certain conditions (has not been demonstrated to date).

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Image of infected skin from a B. dendrobatidis in culture Scanning electron micrograph of scanning electron microscope. zoosporangia with open discharge Fungal discharge tubes are tubes, and rhizoids at the base protruding through the surface (Images courtesy Lee Berger) (arrow)

Clinical signs

In the majority of infected for most of the time, clinical signs are absent. The period of showing signs is typically short and limited to those amphibians that will mostly die. Central nervous system signs predominate; behavioural change, slow and uncoordinated movement, abnormal sitting posture, tetanic spasms, loss of righting reflex, paralysis. Skin changes in chytridiomycosis are typically microscopic and not detectable at the clinical level with any degree of confidence although abnormal skin shedding occurs (skin shed more frequently and in smaller amounts) and erythema may be seen.

Figure 1. Great barred frog (Mixophyes fasciolatus), a lethargic frog with shedding skin accumulating on the body (Courtesy Lee Burger).

Diagnosis

Chytridiomycosis is diagnosed by detecting B. dendrobatidis in the skin of amphibians. This is done by 1) light microscopy, or 2) PCR.

• Light microscopy: Zoosporangia detected in stratum corneum. There are two routine tests: 1) Examination of skin slough with or without staining, or 2) examination of histological sections stained

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with haematoxylin and eosin (Berger et al. 1999b; Pessier et al. 1999), with silver stain (Green et al 2002), or with an immunoperoxidase stain using a polyclonal antibody against B. dendrobatidis (Berger et al. 2002). • Molecular tests: The most common test is the real-time Taqman PCR which can quantify the amount of DNA in the sample (Boyle et al. 2004; Hyatt et al. 2007). A newly developed duplex PCR can be used to detect both B. dendrobatidis and B. salamandrivorans (Blooi et al 2013).

Clinical pathology

No consistent pattern although electrolyte changes have been seen in blood in some species (Voyles et al. 2007).

Pathology

Gross lesions: In most cases nil. Occasional cases have increased sloughing of skin, but this is rarely detectable with the unaided eye.

Histology/ microbiology: For complete description see Berger et al (1999) available online at http://www.jcu.edu.au/school/phtm/PHTM/frogs/histo/chhisto.htm. Summary: Local hyperkeratosis of infected and adjacent cells with presence of sporangia inside cells. Epithelial cells in the layer beneath the superficial layer undergo dissolution, often leading to sloughing of the most superficial layer. Usually no associated inflammatory reaction in dermis. Sites of predilection are the feet, hands and ventral surfaces, but in heavy infections other sites on the body are infected. There are no consistent lesions in other organs.

Differential diagnoses

Using histopathology: Other fungal infections of skin. Artefacts of skin are capable of being confused with sporangia by inexperienced diagnosticians.

Laboratory diagnostic specimens

For histopathology: Skin of feet or toe tips are often adequate but whole frog for necropsy is best to aid diagnosis and rule out other diseases, Fixed in 10% buffered neutral formalin or 70% ethanol.

For PCR: Swab of skin from feet, hands and ventral body surface.

Laboratory procedures

• Light microscope examination of pieces of stratum corneum, unstained or stained with Congo Red (Briggs & Burgin 2003) or Diff Quick. • Histopathology: haematoxylin and eosin (Berger et al. 1999b; Pessier et al. 1999) or silver stain (Green et al 2002). • PCR: Taqman real-time PCR. See Boyle et al (2004) and Hyatt et al. (2007).

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Treatment

The terminal disease has not been successfully treated. The early phases of infection have been treated successfully. However, results are variable and 100% cure cannot at the moment be guaranteed. In many frogs, treatment will control and suppress the infection, but not eliminate it. Frogs remain infected but at a lower level. The infection will then build up over time and can result in death. However, some frogs will be cured by treatment.

At the current time no treatment is 100% effective for Australian amphibians. Chytridiomycosis can be suppressed in almost all cases, but cured in a smaller percentage. Some treatments have been successful overseas.

Bathing in 0.01% suspension for 5 minutes a day for 11 days was reported to successfully treat chytridiomycosis in Dendrobates tinctorius (Nichols & Lamirande 2000) and several other dendrobatid frogs (Forzan et al. 2008). A commercial solution of 25 ppm formalin and 0.10 mg/l malachite green was used for 24 hours every other day four times to successfully treat Xenopus tropicalis (Parker et al. 2002).

Raising the temperature of experimentally infected red eyed tree frogs, Litoria chloris, a native Australian species, to 37°C resulted in cure of chytridiomycosis. All members of a group of 10 experimentally infected L. chloris were cured after being held at 37°C for 2 periods of 8 hours 24 hours apart (Woodhams et al. 2003). However, when this regime was used in other Australian species, it was not 100% successful.

Topical treatment with treated Bd effectively in 12 Archey’s frogs ( archeyi) in New Zealand (Bishop et al. 2009), but further testing is required to confirm the safety of chloramphenicol for other species. A concentration of 1.25 mg/l voriconazole sprayed once daily for seven days onto poison arrow frogs and Iberian midwife toads (Alytes cisternasii) successfully cleared their chytrid infections (Martel et al 2011).

F10, at a concentration of 1:2000, was used to treat two chytrid positive painted reed frogs (Hyperolius marmoratus). The frogs were bathed daily for five minutes for 12 days and subsequently tested negative for B. dendrobatidis by PCR (Barrows et al 2010).

Sodium chloride (NaCl) was used to treat Peron’s tree frogs (Litoria peronei). While the doses of NaCl used did not affect the survival of the pathogen concentrations greater than 3 ppt significantly reduced the growth and motility of the chytrid fungus. Concentrations of 1-4 ppt NaCl were also associated with significantly lower host infection loads while infected hosts exposed to 3 and 4 ppt NaCl were found to have significantly higher survival rates (Stockwell et al 2012).

Prevention and control

Temperature: B. dendrobatidis stops growth at ~28C in vitro and continues growing slowly at 10C (Piotrowski et al. 2004). B. dendrobatidis is highly susceptible to temperatures above 32C. Cultures of B. dendrobatidis die at 32C after 4 days and at 37C after 4 hrs (Berger 2001; Johnson et al 2003). This is in contrast to B. salamandrivorans which has an optimal growth temperature between 10 and 150C and dies above 250C (Martel et al 2013). Life-history trade-offs that see increased zoospore production as sporangia maturation rate slows appears to maintain population growth at sub-optimal temperatures (Woodhams et al. 2008).

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B. dendrobatidis can grow over a wide range of pH (4-9), but optimum is 6-7 (Piotrowski et al. 2004). The amphibian chytrid fungus is susceptible in vitro to many standard disinfectants (Table 3), but is not killed by sterilising ultraviolet light.

Table 3. Disinfection techniques that will kill 100% of Bd zoospores and zoosporangia. From Johnson et al (2003), Webb et al (2007) and Gold et al (2013). RH = relative humidity.

Physical techniques Temp / concentration Minimum time of exposure

Heat 60C 5 min Desiccation 25C RH 70% 3 hr Disinfectants Ethanol 70% 5 min solution 1% 5 min Virkon 0.1%/ 1% 5 min/ 1 min Sodium hypochlorite (bleach) 3% 1 min Didecyl dimethyl ammonium chloride 1x10-3 5 min Benzalkonium chloride 1% 5 min TriGene Virucidal Disinfectant Cleaner 0.1 ml l–1 1 min F10 Super Concentrate Disinfectant 0.33 ml l–1 1 min Betadine Antiseptic Liquid 100 ml l–1 1 min Chlorhexidine 0.75% 1 min Terbinafine 1 mg/ml 5 min

TriGene is the most effective disinfectant yet to be found, and both TriGene and F10 are more effective than previously tested disinfectants (Webb et al. 2007).

B. dendrobatidis will survive and grow in the external environment; it is not an obligate parasite and will survive in tap water for three weeks, deionized water for four weeks, and lake water for seven weeks (Johnson and Speare 2003). It can be grown on a range of keratin supplemented sterile media, including frog skin, snake skin and feather meal agars (Symonds et al. 2008), and it can survive in moist river sand and on bird feathers (Johnson & Speare 2005). Bare human skin has a fungicidal effect on B. dendrobatidis, but this killing effect is reduced by repeated washing with water and ethanol. Nitrile gloves kill B. dendrobatidis on contact, but washing in water decreases this effect. Latex and polyethylene gloves have no killing effect (Mendez et al. 2008).

Surveillance and management

Infection of amphibians with the amphibian chytrid fungus has been listed as a Key Threatening Process in Australia by the Commonwealth Department of Environment and Heritage and a Threat Abatement Plan (TAP) prepared (Department of the Environment and Heritage 2006; Speare 2006). Surveillance for chytridiomycosis especially in currently chytrid-free zones is proposed in this TAP and sampling design has commenced (Skerratt et al. 2008; Murray et al. 2011; Skerratt et al. 2010).

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Chytridiomycosis is not currently included in AUSVETPLAN. Threat Abatement Plan recommendation 1.1.3 proposes that a risk-based approach be used for chytridiomycosis using AUSVETPLAN as a model (Department of the Environment and Heritage 2006b), which is being progressed. Nation-wide mapping protocols and disease risk models have been developed and could serve as the basis for cost-sharing arrangements between states (Murray et al. 2011; Skerratt et al. 2010). Implementing this step remains a priority.

Risk analysis performed by Biosecurity Australia in “Quarantine requirements for the importation of amphibians or their eggs into zoological facilities” (Animal Biosecurity Policy Memorandum 2003/26) did not list chytridiomycosis as a risk since it is endemic in Australia. However, this disregards the risk of importation into chytrid free areas. Although chytridiomycosis is not specifically mentioned, the general hygiene strategies recommended will prevent the release of imported strains of B. dendrobatidis during the initial two years. After two years the amphibians can be released without testing for B. dendrobatidis. However, if being released into a chytrid free area, the same requirements imposed on Australian bred amphibians under the Threat Abatement Plan would apply.

Amphibian chytridiomycosis was listed on the OIE Wildlife Diseases List and has been declared an international notifiable disease (World Organisation for Animal Health 2014). Australia now reports to OIE on its chytrid status via Aquatic Animal Health Committee (AAHC).

Statistics

The most complete dataset currently available on chytrid in Australia is managed by the Amphibian Diseases Group, School of Public Health and Tropical Medicine, James Cook University. Reports of chytrid are also captured in the National Wildlife Health Surveillance Database (eWHIS).

Wildlife disease surveillance in Australia is coordinated by Wildlife Health Australia. The National Wildlife Health Information System (eWHIS) captures information from a variety of sources including Australian government agencies, zoo and wildlife parks, wildlife carers, universities and members of the public. Coordinators in each of Australia's States and Territories report monthly on significant wildlife cases identified in their jurisdictions. NOTE: access to information contained within the National Wildlife Health Information System dataset is by application. Please contact [email protected].

Research

Key research questions:

• What can be done to mitigate the impact of chytridiomycosis where it is endemic? • What can be done to prevent further spread of chytridiomycosis? More detailed research questions that may help answer the above:

• Why do infected amphibians die? • What areas in Australia are chytrid free? • Can B. dendrobatidis spread to and establish in these disease-free areas? • In populations where chytridiomycosis is endemic what determines the impact on the frog population? • Can resistance to infection or clinical disease caused by B. dendrobatidis be selected for? • Can acquired immunity protect amphibians?

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• Does B. dendrobatidis exist as a free-living organism in suitable habitats, particularly natural water bodies and moist substrate in the absence of amphibians? • Can detection of B. dendrobatidis in water bodies be used as a technique to map contaminated and chytrid-free areas? • How do environmental characteristics of natural water bodies (pH, pO2, ion content, nitrate, organic content) and climate (e.g., temperature, rainfall) affect the biology and survival of B. dendrobatidis? • What density of zoospores in natural water bodies can infect susceptible species of amphibians and what is the role of natural water bodies in transmission? • Does the density of zoospores in natural water bodies correlate with intensity of infection of amphibian populations living in those water bodies, and with the level of clinical chytridiomycosis? Can the density of zoospores in natural water bodies or on amphibians be used to predict periods of high risk for amphibian populations? • How does B. dendrobatidis spread between water bodies and amphibian populations? • Are there more non-amphibian vectors of B. dendrobatidis? • Can B. dendrobatidis be eradicated from ponds or small standing water bodies? Recent work indicates that the zooplankton Daphnia magna, readily consumes Batrachochytrium zoospores and may have potential as a biological control agent (Buck et al 2011) • Can amphibian populations be treated or vaccinated? • Can amphibian antichytrid skin bacteria be used to protect susceptible species (Harris et al 2006)?

Known research activities:

• Testing of protocols for mapping regions with unknown chytrid status. • Investigation of pathogenicity and epidemiology including monitoring of amphibian populations that have survived initial epidemic invasion to look for evidence of recovery or ongoing impact and to decipher the host-pathogen relationship in the wild. • Assessing effectiveness of management options such as supplementing critically endangered populations by captive-breeding and reintroduction. • Determining whether innate immunity can be used to improve reintroduction success. • Assessing the potential of selection for innate immunity in protecting amphibian populations. • Testing of hygiene protocols used to reduce the risk of spread. • Assessing the effectiveness of treatment regimes. • Predictive climatic and environmental modelling for risk of impact and spread.

Human health implications

Nil. B. dendrobatidis will not grow above 28C and dies if held at 37C for 4 hours. Homeotherms are thus considered unsuitable hosts.

Conclusions

For control at the national level we need to confirm whether chytridiomycosis is absent in areas predicted as unsuitable by species distribution models (e.g., the Northern Territory), whether there are any areas that are suitable that are currently free, the most effective strategies for monitoring these chytrid-free suitable areas and the best methods to prevent spread of chytridiomycosis to these areas. We need emergency response

WHA Fact sheet: Amphibian chytrid fungus in Australia | May 2014 | 11 plans in case of spread to these areas and for species currently threatened with extinction (underway). We need to determine whether chytridiomycosis can be eradicated from small contaminated water bodies and whether these can be kept disease free. Understanding the medium- to long-term consequences of endemic chytridiomycosis for amphibians is critical for future management in the medium to long-term. We need research to enable us to better mitigate the effects of chytridiomycosis in the short to long-term in affected populations.

Acknowledgements

The following people have had input into this document: Kris Murray, Richard Speare, Lee Skerratt and the WHA coordinators. Peter Holz reviewed the latest version.

A lot of the good work on chytrid has been performed by Australians, most of whom are WHA members and are very willing to share their knowledge, experience and expertise. We suggest that if you are interested in this condition you contact us ([email protected]) and we can put you in touch with them.

References and other information

Barrows M, Koeppel K, Drake G. (2010). The use of F10 as a treatment for bacterial and fungal disease in anurans. In: Proceedings Association of Reptilian and Amphibian Veterinarians Annual Conference, pp. 21-22.

Berger L., Hyatt A.D., Olsen V., Hengstberger S.G., Boyle D., Marantelli G., Humphreys K. & Longcore J.E. (2002). Production of polyclonal antibodies to Batrachochytrium dendrobatidis and their use in an immunoperoxidase test for chytridiomycosis in amphibians. Dis Aquat Org, 48, 213-220.

Berger L., Speare R., Daszak P., Green D.E., Cunningham A.A., Goggin C.L., Slocombe R., Ragan M.A., Hyatt A.D., McDonald K.R., Hines H.B., Lips K.R., Marantelli G. & Parkes H. (1998). Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc Natl Acad Sci U S A, 95, 9031-9036.

Berger L., Speare R. & Hyatt A. (1999a). Chytrid fungi and amphibian declines: Overview, implications and future directions. In: Declines and Disappearances of Australian Frogs. (ed. Campbell A). Environment Australia Canberra, pp. 23-33.

Berger L., Speare R. & Kent A. (1999b). Diagnosis of chytridiomycosis of amphibians by histological examination. Zoos Print Journal, 15, 184-190.

Bishop P.J., Speare R., Poulter R., Butler M., Speare B.J., Hyatt A., Olsen V. & Haigh A. (2009). Elimination of the amphibian chytrid fungus Batrachochytrium dendrobatidis by Archey's frog Leiopelma archeyi. Dis Aquat Org, 84, 9-15.

Blooi M, Pasmans F, Longcore JE, Spitzen-van der Sluijs A, Vercammen F, Martel A. (2013). Duplex real-time PCR for rapid simultaneous detection of Batrachochytrium dendrobatidis and Batrachochytrium salamandrivorans in amphibian samples. J. Clin. Microbiol. 51, 4173-4177.

Boyle D.G., Boyle D.B., Olsen V., Morgan J.A.T. & Hyatt A.D. (2004). Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Dis Aquat Org, 60, 141-148.

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Briggs C. & Burgin S. (2003). A rapid technique to detect chytrid infection in adult frogs. Herpetol Rev, 34.

Buck JC, Truong L, Blaustein AR. (2011) Predation by zooplankton on Batrachochytrium dendrobatidis: biological control of the deadly amphibian chytrid fungus? Biodivers Conserv, 20, 3549-3553.

Department of the Environment and Heritage (2006). Threat abatement plan forinfection of amphibians with chytrid fungus resulting in chytridiomycosis. URL http://www.environment.gov.au/biodiversity/threatened/publications/tap/chytrid.html.

Doherty-Bone TM, Gonwouo NL, Hirschfeld M, Ohst T, Weldon C,Perkins M, Kouete MT, Browne RK, Loader SP, Gower DJ, Wilkinson MW, Rödel MO, Penner J, Barej MF, Schmitz A, Plötner J, Cunningham AA. (2013). Batrachochytrium dendrobatidis in amphibians of Cameroon, including first records for caecilians. Dis Aquat Org, 102, 187-194.

Forzan M.J., Gunn H. & Scott P. (2008). Chytridiomycosis in an aquarium collection of frogs: Diagnosis, treatment, and control. J Zoo Wildl Med, 39, 406-411.

Gold KK, Reed PD, Bemis DA, Miller DL, Gray MJ, Souza MJ. (2013). Efficacy of common disinfectants and terbinafine in inactivating the growth of Batrachochytrium dendrobatidis in culture. Dis Aquat Org, 107, 77- 81.

Gower DJ, Doherty-Bone T. Loader SP, Wilkinson M, Kouete MT, Tapley B, Orton F, Daniel OZ, Wynne F, Flach E, Müller H, Menegon M, Stephen J, Browne RK, Fisher MC, Cunningham AA, Garner TWJ. (2013). Batrachochytrium dendrobatidis infection and lethal chytridiomycosis in caecilian amphibians (Gymnophiona). EcoHealth, 10, 173-183.

Harris RN, James TY, Lauer A, Simon MA, Patel A. (2006) Amphibian pathogen Batrachochytrium dendrobatidis is inhibited by the cutaneous bacteria of amphibian species. EcoHealth, 3, 53-56.

Hines H., Mahony M. & McDonald K. (1999). An assessment of frog declines in wet subtropical Australia In: Declines and Disappearances of Australian Frogs (ed. Campbell A). Environment Australia Canberra, pp. 44-63.

Hyatt A.D., Boyle D.G., Olsen V., Boyle D.B., L. B., Obendorf D., Dalton A., Kriger K., Hero J.-M., Hines H., Phillott R., Campbell R., Marantelli G., Gleason F. & Colling A. (2007). Diagnostic assays and sampling protocols for the detection of Batrachochytrium dendrobatidis. Dis Aquat Org, 73, 175-192.

Johnson M.L. & Speare R. (2005). Possible modes of dissemination of the amphibian chytrid Batrachochytrium dendrobatidis in the environment. Dis Aquat Org, 65, 181-186.

Johnson ML and Speare R. (2003). Survival of Batrachochytrium dendrobatidis in water: quarantine and disease control implications. Emerg Infect Dis, 9, 922-925.

Johnson M, Berger L, Philips L, Speare R (2003). Fungicidal effects of chemical disinfectants, UV light, desiccation and heat on the amphibian chytrid, Batrachochytrium dendrobatidis. Dis Aquat Org 57, 255–260.

Laurance W., McDonald K. & Speare R. (1996). Catastrophic declines of Australian rain forest frogs: support for the epidemic disease hypothesis. Conserv Biol, 10, 406-413.

Longcore J.E., Pessier A.P. & Nichols D.K. (1999). Batrachochytrium dendrobatidis gen et sp nov, a chytrid pathogenic to amphibians. Mycologia, 91, 219-227.

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Martel A, Van Rooij P, Vercauteren G, Baert K, Van Waeyenberghe L, Debacker P, Garner TWJ, Woeltjes T, Ducatelle R, Haesebrouck F, Pasmans F. (2010). Developing a safe antifungal treatment protocol to eliminate Batrachochytrium dendrobatidis from amphibians. Medical Mycology, 2011, 49, 143-149.

Martel A, Spitzen-van der Sluijs A, Blooi M, Bert W, Ducatelle R, Fisher MC, Woeltjes A, Bosman W, Chiers K, Bossuyt F, Pasmans F. (2013). Batrachochytrium salamandrivorans sp. nov. causes lethal chytridiomycosis in amphibians. Proc. Natl. Acad. Sci. U. S. A. 110, 15325–15329.

McMahon TA, Brannelly LA, Chatfield MWH, Johnson PTJ, Joseph MB, McKenzie VJ, Richards-Zawacki CL, Venesky MD, Rohr JR. (2013). Chytrid fungus Batrachochytrium dendrobatidis has nonamphibian hosts and releases chemicals that cause pathology in the absence of infection. Proc. Natl. Acad. Sci. U.S.A. 110, 210-215.

Mendez D., Webb R., Berger L. & Speare R. (2008). Survival of the amphibian chytrid fungus Batrachochytrium dendrobatidis on bare hands and gloves: hygiene implications for amphibian handling. Dis Aquat Org, 82, 97- 104.

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Pessier A., Nichols D., Longcore J. & Fuller M. (1999). Cutaneous chytridiomycosis in poison dart frogs (Dendrobates spp.) and White's tree frogs (Litoria caerulea). Journal of Veterinary Diagnostic Investigation, 11, 194-199.

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Retallick RWR, McCallum H, Speare R. (2004). Endemic infection of the amphibian chytrid fungus in a frog community post-decline. PLoS Biol, 2(11): e351, 1965-1971.

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Skerratt L.F., Berger L., Hines H.B., McDonald K.R., Mendez D. & Speare R. (2008). Survey protocol for detecting chytridiomycosis in all Australian frog populations. Dis Aquat Org, 80, 85-94.

Skerratt L.F., McDonald K.R., Hines H.B., Berger L., Mendez D., Phillott A., Cashins S.D., Murray K.A. & Speare R. (2010). Application of the survey protocol for chytridiomycosis to Queensland, Australia. Dis Aquat Org 92, 117-129.

Stockwell MP, Clulow J, Mahony MJ. (2012). Sodium chloride inhibits the growth and infective capacity fo the amphibian chytrid fungus and increases host survival rates. PLoS ONE, 7(5):e36942. doi:10.1371/journal.pone.0036942.

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Voyles J., Berger L., Young S., Speare R., Webb R., Warner J., Rudd D., Campbell R. & Skerratt L.F. (2007). Electrolyte depletion and osmotic imbalance in amphibians with chytridiomycosis. Dis Aquat Org, 77, 113- 118.

Webb R., Mendez D., Berger L. & Speare R. (2007). Additional disinfectants effective against the amphibian chytrid fungus Batrachochytrium dendrobatidis. Dis Aquat Org, 74, 13-16.

Woodhams D.C., Alford R.A., Briggs C.J., Johnson M. & Rollins-Smith L.A. (2008). Life-history trade-offs influence disease in chaning climates: strategies of an amphibian pathogen. Ecology, 89, 1627-1639.

Woodhams D.C., Alford R.A. & Marantelli G. (2003). Emerging disease of amphibians cured by elevated body temperature. Dis Aquat Org, 55, 65-67.

World Organisation for Animal Health (2013). Aquatic Animal Code 2013. URL http://www.oie.int/eng/normes/fcode/en_sommaire.htm

Other information sources:

Speare R. Amphibian diseases home page. http://www.jcu.edu.au/school/phtm/PHTM/frogs/ampdis.htm

Updated: 7 May 2014.

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We are interested in hearing from anyone with information on this condition in Australia, including laboratory reports, historical datasets or survey results that could be added to the National Wildlife Health Information System. If you can help, please contact us at [email protected].

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Disclaimer

This fact sheet is managed by Wildlife Health Australia for information purposes only. Information contained in it is drawn from a variety of sources external to Wildlife Health Australia. Although reasonable care was taken in its preparation, Wildlife Health Australia does not guarantee or warrant the accuracy, reliability, completeness, or currency of the information or its usefulness in achieving any purpose. It should not be relied on in place of professional veterinary consultation. To the fullest extent permitted by law, Wildlife Health Australia will not be liable for any loss, damage, cost or expense incurred in or arising by reason of any person relying on information in this fact sheet. Persons should accordingly make and rely on their own assessments and enquiries to verify the accuracy of the information provided.

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