|Zoosporangia of Batrachochytrium dendrobatidis growing on a freshwater arthropod (a) and on algae (b). The scale bars represent 30 µm.|
Longcore, Pessier & D.K. Nichols (1999)
Batrachochytrium dendrobatidis is a chytrid fungus that causes the disease chytridiomycosis. In the decade after it was first discovered in amphibians in 1998, the disease devastated amphibian populations around the world, in a global decline towards multiple extinctions, part of the Holocene extinction.
Some amphibian species appear to have an innate capacity to withstand chytridiomycosis infection. Even within species that generally succumb, some populations survive, possibly demonstrating that these traits or alleles of species are being subjected to evolutionary selection.
The generic name is derived from the Greek words batrachos (frog) and chytra (earthen pot), while the specific epithet is derived from the genus of frogs from which the original confirmation of pathogenicity was made (Dendrobates).
Batrachochytrium dendrobatidis is the single species of the monotypic genus Batrachochytrium. The initial classification of the pathogen as a chytrid was based on zoospore ultrastructure. DNA analysis of the ssu-rDNA has corroborated the view, with the closest match to Chytridium confervae.
B. dendrobatidis can grow within a wide temperature range (4-25°C), with optimal temperatures being between 17-25°C. The wide temperature range for growth, including the ability to survive at 4°C gives the fungus the ability to overwinter in its hosts, even where temperatures in the aquatic environments are low. The species does not grow well above temperatures of 25°C, and growth is halted above 28°C. Infected red-eyed treefrogs (Litoria chloris) recovered from their infections when incubated at a temperature of 37°C.
B. dendrobatidis infects the keratinized skin of amphibians. The fungus in the epidermis has a thallus bearing a network of rhizoids and smooth-walled, roughly spherical, inoperculate (without an operculum) sporangia. Each sporangium produces a single tube to discharge spores.
Zoospore structure 
Zoospores of B. dendrobatidis, which are typically 3-5 µm in size, have an elongate–ovoidal body with a single, posterior flagellum (19-20 µm long), and possess a core area of ribosomes often with membrane-bound spheres of ribosomes within the main ribosomal mass. A small spur has been observed, located at the posterior of the cell body, adjacent to the flagellum, but this may be an artifact in the formalin-fixed specimens. The core area of ribosomes is surrounded by a single cisterna of endoplasmic reticulum, two to three mitochondria, and an extensive microbody–lipid globule complex. The microbodies closely appose and almost surround four to six lipid globules (three anterior and one to three laterally), some of which appear bound by a cisterna. Some zoospores appear to contain more lipid globules (this may have been a result of a plane-of-sectioning effect, because the globules were often lobed in the zoospores examined). A rumposome has not been observed.
Flagellum structure 
A nonfunctioning centriole lies adjacent to the kinetosome. Nine interconnected props attach the kinetosome to the plasmalemma, and a terminal plate is present in the transitional zone. An inner ring-like structure attached to the tubules of the flagellar doublets within the transitional zone has been observed in transverse section. No roots associated with the kinetosome have been observed. In many zoospores, the nucleus lies partially within the aggregation of ribosomes and was invariably situated laterally. Small vacuoles and a Golgi body with stacked cisternae occurred within the cytoplasm outside the ribosomal area. Mitochondria, which often contain a small number of ribosomes, are densely staining with discoidal cristae.
Life cycle 
B. dendrobatidis has two primary life stages: a sessile, reproductive zoosporangium and a motile, uniflagellated zoospore released from the zoosporangium. The zoospores are known be active only for a short period of time, and can travel short distances of one to two centimeters. However, the zoospores are capable of chemotaxis, and can move towards a variety of molecules that are present on the amphibian surface, such as sugars, proteins and amino acids. B. dendrobatidis also contains a variety of proteolytic enzymes and esterases that help it digest amphibian cells and use amphibian skin as a nutrient source. Once the zoospore reaches its host, it forms a cyst underneath the surface of the skin, and initiates the reproductive portion of its life cycle. The encysted zoospores develop into zoosporangia, which may produce more zoospores that can reinfect the host, or be released into the surrounding aquatic environment. The amphibians infected with these zoospores are shown to die from cardiac arrest.
Varying forms 
B. dendrobatidis has occasionally been found in forms distinct from its traditional zoospore and sporangia stages. For example, before the 2003 European heatwave that decimated populations of the water frog Rana lessonae through chytridiomycosis, the fungus existed on the amphibians as spherical, unicellular organisms, confined to minute patches (80-120 micrometers across). These organisms, unknown at the time, were subsequently identified as B. dendrobatidis. Characteristics of the organisms were suggestive of encysted zoospores; they may have embodied a resting spore, a saprobe, or a parasitic form of the fungus that is non-pathogenic.
Habitat and relationship to amphibians 
The fungus grows on amphibian skin and produces aquatic zoospores. It is widespread and ranges from lowland forests to cold mountain tops. It is sometimes a non-lethal parasite and possibly a saprophyte. The fungus is associated with host mortality in highlands or during winter, and becomes more pathogenic at lower temperatures.
Chytridiomycosis prevalence 
It has been suggested that B. dendrobatidis originated in Africa and subsequently spread to other parts of the world by trade in African clawed frogs (Xenopus laevis). In this study, 697 archived specimens of three species of Xenopus, previously collected from 1879 to 1999 in southern Africa were examined. The earliest case of chytridiomycosis was found in a X. laevis specimen from 1938. The study also suggests that chytridiomycosis had been a stable infection in southern Africa from 23 years prior to finding any infected outside of Africa.
Bullfrogs (Rana catesbiana), also widely distributed, are also thought to be carriers of the disease due to their inherent low susceptibility to B. dendrobatidis infection. The bullfrog often escapes captivity and can establish feral populations where it may introduce the disease to new areas. It has also been shown that B. dendrobatidis can survive and grow in moist soil and on bird feathers, suggesting that B. dendrobatidis may also be spread in the environment by birds and transportation of soils. Infections have been linked to mass mortalities of amphibians in North America, South America, Central America, Europe and Australia. B. dendrobatidis has been implicated in the extinction of the sharp-snouted day frog (Taudactylus acutirostris) in Australia.
A wide variety of amphibian hosts have been identified as being susceptible to infection by B. dendrobatidis, including wood frogs (Rana sylvatica), the mountain yellow-legged frog (Rana muscosa) the southern two-lined salamander (Eurycea cirrigera), San Marcos Salamander (Eurycea nana) Texas Salamander (Eurycea neotenes) Blanco River Springs Salamander (Eurycea pterophila) Barton Springs Salamander (Eurycea sosorum) Jollyville Plateau Salamander (Eurycea tonkawae)  Ambystoma jeffersonianum, the western chorus frog (Pseudacris triseriata), the southern cricket frog (Acris gryllus), the eastern spadefoot toad (Scaphiopus holbrooki), the southern leopard frog (Rana sphenocephala), the Rio Grande Leopard frog (Lithobates berlandieri), and the Sardinian newt (Euproctus platycephalus).
Southeast Asia 
While most studies concerning B. dendrobatidis have been performed in various locations across the world, the presence of the fungus in Southeast Asia remains a relatively recent development. The exact process through which the fungus was introduced to Asia is not known, however, as mentioned above, it has been suggested transportation of asymptomatic carriers species (e.g. Lithobates catesbeianus, the American Bullfrog) may be a key component in the dissemination of the fungus, at least in China. Initial studies demonstrated the presence of the fungus on islands states/countries such as Hong Kong, Indonesia, Taiwan, and Japan. Soon thereafter, mainland Asian countries such as Thailand, South Korea, and China reported incidences of B. dendrobatidis among their amphibian populations. Much effort has been put into classifying herptofauna in countries like Cambodia, Vietnam, and Laos where new species of frogs, toads, and other amphibians and reptiles are being discovered on a frequent basis. Scientists simultaneously are swabbing herptofauna in order to determine if these newly discovered animals possess traces of the fungus.
In Cambodia, a study showed B. dendrobatidis to be prevalent throughout the country in areas near Phnom Penh (in a village <5 km), Sihanoukville (frogs collected from the local market), Kratie (frogs collected from streets around the town), and Siem Reap (frogs collected from a national preserve: Angkor Centre for Conservation of Biodiversity). Another study in Cambodia questioned the potential anthropological impact in the dissemination of B. dendrobatidis on local amphibian populations in 3 different areas in relation to human interaction: low (an isolated forest atop a mountain people rarely visit), medium (a forest road ~15km from a village that is used at least once a week), and high (a small village where humans interact with their environment on a daily basis). Using quantitative real-time PCR, evidence of B. dendrobatidis was found in all 3 sites with the highest percentage of amphibians positive for the fungus from the forest road (medium impact; 50%), followed by the mountain forest (low impact; 44%) and village (high impact; 36%). Human influence most likely explains detection of the fungus in the medium and high areas, however it does not provide an adequate explanation why even isolated amphibians were positive for B. dendrobatidis. This may go unanswered until more research is performed on transmission of the fungus across landscapes.
Immunity hypotheses 
Due to the fungus' immense impact on amphibian populations, considerable research has been undertaken to devise methods to combat its proliferation. Among the most promising is the revelation that amphibians in colonies that survive the passage of the chytrid epidemic tend to carry higher levels of the bacterium Janthinobacterium lividum. This bacterium produces antifungal compounds, such as indole-3-carboxaldehyde and violacein, that inhibit the growth of B. dendrobatidis even at low concentrations. Similarly, the bacterium Lysobacter gummosus found on the red-backed salamander (Plethodon cinereus), produces the compound 2,4-diacetylphloroglucinol that is inhibitory to the growth of B. dendrobatidis.
Understanding the interactions of microbial communities present on amphibians’ skin with fungal species in the environment can reveal why certain amphibians, such as the frog Rana muscosa, are susceptible to the fatal effects of B. dendrobatidis and why others, such as the salamander Hemidactylium scutatum, are able to coexist with the fungus. As mentioned before, the antifungal bacterial species Janthinobacterium lividum, found on several amphibian species, has been shown to prevent the effects of the pathogen even when added to another amphibian that lacks the bacteria (B. dendrobatidis-susceptible amphibian species). Interactions between cutaneous microbiota and B. dendrobatidis can be altered to favor the resistance of the disease, as seen in past lab studies concerning the addition of the violacein-producing bacteria J. lividum to amphibians that lacked sufficient violacein, allowing them to inhibit infection. Although the exact concentration of violacein (antifungal metabolite produced by J. lividum) needed to inhibit the effects of B. dendrobatidis is not fully confirmed, violacein concentration can determine whether or not an amphibian will experience morbidity (or mortality) caused by the chytrid fungus B. dendrobatidis. The frog Rana muscosa, for example, has been found to have very low concentrations of violacein on its skin, yet the concentration is so small that it is unable to facilitate increased survivability of the frog; furthermore, Janthinobacterium lividum has not been found to be present on the skin of Rana muscosa. This implies that the antifungal bacteria J. lividum (native to other amphibians' skin, such as Hemidactylium scutatum) is able to produce a sufficient amount of violacein to prevent infection by B. dendrobatidis and allow coexistence with the potentially deadly fungus.
Studies conducted by Dr. Reid Harris and colleagues of the Department of Biology of James Madison University in Virginia have shown that the addition of the anti-chytrid (antifungal) bacteria Janthinobacterium lividum to the skin of B. dendrobatidis-susceptible amphibians (i.e. Rana muscosa juveniles) increases the concentration of the antifungal metabolite violacein, which in turn decreases the mortality rate due to infection by B. dendrobatidis and also increases survivability. The removal of resident skin bacteria of the amphibians precedes the application of Janthinobacterium lividum and exposure to B. dendrobatidis zoospores (in the majority of experiments that have been previously been conducted), which reduces bacterial species on the amphibians' skin and also reduces possible interactions between J. lividum and other species of bacteria present on the skin. This allows for a standard condition of the amphibians’ skin that can then be compared to the J. lividum treatment of an experiment, thereby yielding simpler and more attributable survival/inhibition results concerning the newly added bacterial species (J. lividum). To reiterate, the majority of research done in this area has been concerned with prevention by applying J. lividum to amphibians before infection (by B. dendrobatidis) and after removal of their original skin bacteria. However, little research has been conducted to see if the addition of Janthinobacterium lividum without initial removal of the amphibians’ cutaneous microbiota is still as effective against the pathogen. Further research is needed to explore conditions and treatments that will include the original cutaneous bacterial species of the amphibians (that is to say, excluding the bacterial removal procedures commonly done before applying the antifungal bacteria) that will determine whether or not the addition of J. lividum will still increase survivability by inhibiting the fungus even without the initial removal of the resident skin bacteria. This would allow for a more practical method of bioaugmentation when treating a B. dendrobatidis-susceptible amphibian population in nature.
Effects of pesticides 
The hypothesis that pesticide use has contributed to declining amphibian populations has been suggested a several times in the literature. In 2007, this hypothesis was corroborated, as it was shown that sublethal exposure to the pesticide carbaryl (a cholinesterase inhibitor) increase susceptibility of foothill yellow-legged frogs (Rana boylii) to chytridomycosis. In particular, the skin peptide defenses were significantly reduced after exposure to cabaryl, suggesting that pesticides may inhibit this innate immune defense, and increase susceptibility to disease.
See also 
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|Wikispecies has information related to: Chytridiaceae|
|Wikimedia Commons has media related to: Chytridiomycota|
- Chytrid Fungi Online at University of Alabama
- Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America
- Emerging Infectious Diseases and Amphibian Population Declines
- Survival of Batrachochytrium dendrobatidis in Water: Quarantine and Disease Control Implications
- Index Fungorum
- Batrachochytrium dendrobatidis at the Encyclopedia of Life