Batrachochytrium dendrobatidis (Bd) is a chytrid fungus that was first reported as the cause of chytridiomycosis (an often deadly and highly infectious disease of amphibians) in wild and captive frogs collected in North and Central America and Australia. It was formally described as a species from captive South American frogs in 1999 and virtually everything known about it has been discovered in the context of its role in global amphibian declines. This fungus has now been found on all continents except Antarctica and has been detected in frog specimens collected as long ago as 1938. Chytrid fungi are typically aquatic and differ from other fungi in that they have a motile, flagellated zoospore. Many chytrid species have been described from aquatic habitats and soils as free-living or commensal organisms and as parasites of algae, invertebrates, fungi, and plants. Bd is one of only two chytrids known to parasitize vertebrates and it is the only one known to infect and develop within the keratinized epidermal cells of living amphibian skin. Bd infects an extraordinarily broad diversity of host species--in fact, it has the widest known host range of any pathogen. It is known to infect over 350 species of amphibians and has been been implicated in driving the decline or extinction of over 200 of these species. (Fisher et al. 2009; Kilpatrick et al. 2009)
Batrachochytrium dendrobatidis infects Anura
Fungus / infection vector
Batrachochytrium dendrobatidis is spread by Alytes obstetricans
In Great Britain and/or Ireland:
Fungus / infection vector
Batrachochytrium dendrobatidis is spread by Rana catesbeiana
The origin of the BD chytridiomycosis pandemic remains controversial. According to the "novel pathogen hypothesis", Bd has been recently introduced to areas where it is causing population declines. An alternative scenario, the "endemic pathogen hypothesis", suggests that Bd has been present in recently affected populations for a long time, with its newly devastating impact resulting from changes in host susceptibility, pathogen virulence, environmental changes, or some combination of these factors. Molecular data from sites broadly dispersed around the world seem to suggest a single origin of Bd with evidence of local and global spread; these findings lend strong support to the novel pathogen hypothesis. However, in some populations Bd appears to be present without causing subsequent declines and in other locations, it appears to have been present before widespread declines. Similarly, in some species that declined regionally but persisted in small populations, Bd is still present and infecting frogs without driving these populations to extinction. (Fisher et al. 2009 and references therein; Kilpatrick et al. 2009 and references therein; Lam et al. 2010) The possible role of natural or anthropogenic global climate cycles, or long-term changes, in driving outbreaks of amphibian chytridiomycosis is controversial (for discussion, see, e.g., Lips et al. 2008; Rohr et al. 2008; Rohr and Raffel 2010).
Chytridiomycosis is notable for an overall lack of obvious disease pathologies. Metamorphosed amphibians infected with Bd typically exhibit epidermal hyperplasia and hyperkeratosis, and possibly upregulated skin shedding, but only rarely do they exhibit any lesions visible to the naked eye. Larval anuran amphibians may exhibit visible deformities of keratinized mouthparts. Bd is hypothesized to produce lethal toxins either before or after infection, to interfere with water uptake and lead to death due to dehydration, or to cause sharp osmotic imbalances that may interfere with water regulation and/or neurological function. (Fisher et al. 2009) Voyles et al. (2009) studied Bd infection in the treefrog Litoria caerulea. They found that in diseased individuals electrolyte transport across the epidermis was inhibited by >50%, plasma sodium and potassium concentrations were reduced by ~20% and ~50%, respectively, and asystolic cardiac arrest resulted in death. Given the critical importance of the skin in maintaining amphibian homeostasis, disruption of cutaneous function may be the mechanism by which Bd produces morbidity and mortality across a wide range of phylogenetically distant amphibian taxa.
Fungicidal protocols have been developed that are generally very effective for captive frogs, making captive breeding an important component of frog conservation efforts. These approaches include the use of elevated temperature, formalin/malachite green, and standard veterinary antifungal drugs. Good results for clearing infection in captive colonies have suggested the possibility of clearing infection in natural populations using catch, treat, and release methods using inexpensive, mobile biocontainment laboratories. An important new development in research on mitigation is the recognition that some bacteria that occur naturally on amphibian skin produce antifungal metabolites. This finding suggests the possibility of a probiotic approach to Bd mitigation, i.e., intentionally spreading these protective bacteria. (Fisher et al. 2009 and references therein; Lam et al. 2010)
It remains unclear how Bd has dispersed to and persisted in remote pristine environments where anthropogenic introduction is unlikely. If Bd can survive independently of amphibian hosts, it must use non-amphibian organic materials as nutrient resources. Although Bd DNA has been detected in water bodies and on rocks, conclusive evidence of Bd persistence in the environment is lacking. Species, populations, and individuals vary widely in susceptibility to chytridiomycosis. Mortality rates in laboratory infection experiments can range from 0% to 100%, depending on the species, age of animals, and temperature regime. In the wild, some species and populations are extirpated. while others, those that survive initial declines, persist with various levels of infection. While the disease dynamics are undoubtedly influenced by local environmental conditions, particularly temperature, inherent differences in host susceptibility and behavior are also important. Colonization by Bd and subsequent disease development may be influenced by host defense mechanisms, such as secretions of antimicrobial peptides or bacterial commensals with anti-fungal properties. Some species-specific behavioral characteristics such as microhabitat selection, basking, aggregating in retreat sites, or association with water bodies may also affect the likelihood of infection and disease. (Rosenbaum et al 2010 and references therein)
Batrachochytrium dendrobatidis (or Bd for short) 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. Another explanation for such occurrences, explained below, could be that some forms of the fungus are not pathogenic.
The generic name is derived from the Greek words batracho (frog) and chytr (earthern pot), while the specific epithet is derived from the genus of frogs from which the original confirmation of pathogenicity was made (Dendrobates).
B. dendrobatidis is a monotypic species of the 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.
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.
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.
Bd 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. Bd 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.
Bd 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 Bd. 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 conditionally non-pathogenic . Once the heatwave began, the organisms assumedly changed into the more familiar disease-causing zoospores. This suggests that some populations afflicted with Bd may be free of chytridiomycosis not because of some inherent immunity, but because environmental conditions have not altered the organism into its more common, pathogenic form.
Habitat and relationship to amphibians
The fungus grows on amphibian skin and produces aquatic zoospores . It is widespread and ranges from deserts and 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 .
It has been suggested that Bd 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 Bd 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 Bd can survive and grow in moist soil and on bird feathers, suggesting that Bd 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. Bd 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 Bd, 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).
In 2008, the genomes of two Bd isolates were sequenced, and scientists have begun using this genetic information to help understand the molecular basis of the Bd life cycle and amphibian pathogenicity. Analysis of global gene expression using whole-genome arrays has revealed that greater than 55% of the approximately 9000 genes in the Bd genome undergo differential expression between the sessile sporangium stage and the infectious zoospore stage. Expression of a variety of metalloproteases (enzymes that can break down keratin-containing tissue, like amphibian tissue) is believed to contribute to pathogenicity by enabling cutaneous infection.
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 Bd 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 Bd.
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 Bd 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 (Bd-susceptible amphibian species). Interactions between cutaneous microbiota and Bd 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 Bd is not fully confirmed, violacein concentration can determine whether or not an amphibian will experience morbidity (or mortality) caused by the chytrid fungus Bd. 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 Bd 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 Bd-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 Bd and also increases survivability. The removal of resident skin bacteria of the amphibians precedes the application of Janthinobacterium lividum and exposure to Bd 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 Bd) 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 Bd-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.
- ^ Ellis, Richard (2004). No Turning Back: The Life and Death of Animal Species. New York: Harper Perennial. p. 187. ISBN 0-06-055804-0.
- ^ a b Longcore JE, Pessier AP, Nichols DK (1999). "Batrachochytrium Dendrobatidis gen. et sp. nov, a chytrid pathogenic to amphibians". Mycologia 91 (2): 219–227. doi:10.2307/3761366. http://jstor.org/stable/3761366.
- ^ a b Piotrowski JS, Annis S, Longcore JE (2004). "Physiology of Batrachochytrium dendrobatidis, a chytrid pathogen of amphibians". Mycologia 96 (1): 9–15. doi:10.2307/3761981. http://jstor.org/stable/3761981.
- ^ Woodhams DC, Alford RA, Marantelli G (June 2003). "Emerging disease of amphibians cured by elevated body temperature". Dis. Aquat. Org. 55 (1): 65–7. doi:10.3354/dao055065. PMID 12887256.
- ^ a b Garner TW, Perkins MW, Govindarajulu P, Seglie D, Walker S, Cunningham AA, Fisher MC (September 2006). "The emerging amphibian pathogen Batrachochytrium dendrobatidis globally infects introduced populations of the North American bullfrog, Rana catesbeiana". Biol. Lett. 2 (3): 455–9. doi:10.1098/rsbl.2006.0494. PMID 17148429. PMC 1686185. http://journals.royalsociety.org/openurl.asp?genre=article&doi=10.1098/rsbl.2006.0494.
- ^ Moss AS, Reddy NS, Dortaj IM, San Francisco MJ (2008). "Chemotaxis of the amphibian pathogen Batrachochytrium dendrobatidis and its response to a variety of attractants". Mycologia 100 (1): 1–5. doi:10.3852/mycologia.100.1.1. PMID 18488347.
- ^ Symonds EP, Trott DJ, Bird PS, Mills P (2008). "Growth characteristics and enzyme activity in Batrachochytrium dendrobatidis isolates". Mycopathologia 166 (3): 143–147. doi:10.1007/s11046-008-9135-y. PMID 18568420.
- ^ Berger L, Hyatt AD, Speare R, Longcore JE (December 2005). "Life cycle stages of the amphibian chytrid Batrachochytrium dendrobatidis". Dis. Aquat. Org. 68 (1): 51–63. doi:10.3354/dao068051. PMID 16465834.
- ^ Di Rosa, Ines et al. "The Proximate Cause of Frog Declines?" Nature 447.31 (2007) E4-E5.
- ^ Ron, S. R. Predicting the distribution of the amphibian pathogen Bd in the New World. Biotropica 37 209-221 (2005)
- ^ Daszak, P.; Cuningham, A. A. & Hyatt, A.D. Infection disease and amphibian population declines. Divers. Distrib. 9 141-150 (2003).
- ^ a b Weldon C, du Preez LH, Hyatt AD, Muller R, Spears R (December 2004). "Origin of the amphibian chytrid fungus". Emerging Infect. Dis. 10 (12): 2100–5. PMID 15663845. http://www.cdc.gov/ncidod/EID/vol10no12/03-0804.htm.
- ^ Kats LB, Ferrer RP (2003). "Alien predators and amphibian declines: review of two decades of science and the transition to conservation". Diversity and Distributions 9: 99–110. doi:10.1046/j.1472-4642.2003.00013.x.
- ^ Daszak P, Strieby A, Cunningham AA, Longcore JE, Brown CC, Porter D (2004). "Experimental evidence that the bullfrog (Rana catesbiana) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians". Herpetological Journal 14: 201–207.
- ^ Johnson ML, Speare R (July 2005). "Possible modes of dissemination of the amphibian chytrid Batrachochytrium dendrobatidis in the environment". Dis. Aquat. Org. 65 (3): 181–6. doi:10.3354/dao065181. PMID 16119886.
- ^ Lips KR (1999). "Mass mortality and population declines of anurans at an upland site in western Panama". Conservation Biology 13 (1): 117–125. doi:10.1046/j.1523-1739.1999.97185.x.
- ^ Daszak P, Cunningham AA, Hyatt AD (2003). "Infectious disease and amphibian population declines" (PDF). Diversity and Distributions 9: 141–50. doi:10.1046/j.1472-4642.2003.00016.x. http://www.conservationmedicine.org/papers/Infect.%20Dis.%20&%20Amphib%20Pop%20Declines%202003.pdf.
- ^ Herrera RA, Steciow MM, Natale GS (2005). "Chytrid fungus parasitizing the wild amphibian Leptodactylus ocellatus (Anura: Leptodactylidae) in Argentina". Diseases of Aquatic Organisms 64 (3): 247–52. doi:10.3354/dao064247. PMID 15997823.
- ^ Schloegel LM, Hero JM, Berger L, Speare R, McDonald K, Daszak P (2006). "The decline of the sharp-snouted day frog (Taudactylus acutiostris): the first documented case of extinction by infection in a free-ranging wildlife species?". EcoHealth 3: 35–40. doi:10.1007/s10393-005-0012-6.
- ^ Reeves MK (2008). "Batrachochytrium dendrobatidis in wood frogs (Rana sylvatica) from Three National Wildlife Refuges in Alaska, USA". Herpetological Review 39 (1): 68–70.
- ^ Andre SE, Parker J, Briggs CJ (2008). "Effect of temperature on host response to Batrachochytrium dendrobatidis infection in the mountain yellow-legged frog (Rana muscosa)". Journal of Wildlife Diseases 44 (3): 716–720. PMID 18689660.
- ^ Byrne MW, Davie EP, Gibbons JW (2008). "Batrachochytrium dendrobatidis occurrence in Eurycea cirrigera". Southeastern Naturlaist 7 (3): 551–555. doi:10.1656/1528-7092-7.3.551.
- ^ Gaertner JP, Forstner MRJ, O'Donnell L, Hahn D (2009). "Detection of Batrachochytrium dendrobatidis in endemic salamander species from Central Texas". EcoHealth 6 (1): 20–26. doi:10.1007/s10393-009-0229-x. PMID 19424755.
- ^ Brodman R, Briggler JT (2008). "Batrachochytrium dendrobatidis in Ambystoma jeffersonianum larvae in southern Indiana". Herpetological Review 39 (3): 320–321.
- ^ Lehtinen RM, Kam Y-C Richards CL (2008). "Preliminary surveys for Batrachochytrium dendrobatidis in Taiwan". Herpetological Review 39 (3): 317–318.
- ^ Lovich R, Ryan MJ, Pessier AP, CLaypool B (2008). "Infection with the fungus Batrachochytrium dendrobatidis in a non-native Lithobates berlandieri below sea level in the Coachella Valley, California, USA". Herpetological Review 39 (3): 315–317.
- ^ Bovero S, Sotgiu G, Angelini C, Doglio S, Gazzaniga E, Cunningham AA, Garner TWJ (2008). "Detection of chytridiomycosis caused by Batrachochytrium dendrobatidis in the endangered sardinian newt (Euproctus platycephalus) in Southern Sardinia, Italy". Journal of Wildlife Diseases 44 (3): 712–715. PMID 18689659.
- ^ Fisher MC (November 2008). "Molecular toolkit unlocks life cycle of the panzootic amphibian pathogen Batrachochytrium dendrobatidis". Proc. Natl. Acad. Sci. U.S.A. 105 (45): 17209–10. doi:10.1073/pnas.0809801105. PMID 18997006. PMC 2582274. http://www.pnas.org/cgi/pmidlookup?view=long&pmid=18997006. Retrieved 2009-01-05.
- ^ Rosenblum EB, Stajich JE, Maddox N, Eisen MB (November 2008). "Global gene expression profiles for life stages of the deadly amphibian pathogen Batrachochytrium dendrobatidis". Proc. Natl. Acad. Sci. U.S.A. 105 (44): 17034–9. doi:10.1073/pnas.0804173105. PMID 18852473. PMC 2566996. http://www.pnas.org/cgi/pmidlookup?view=long&pmid=18852473. Retrieved 2009-01-05.
- ^ (BBC News) Richard Black, " Bacteria could stop frog killer" Accessed 7 June 2008.
- ^ Brucker RM, Harris RN, Schwantes CR, Gallaher TN, Flaherty DC, Lam BA, Minbiole KP (November 2008). "Amphibian chemical defense: antifungal metabolites of the microsymbiont Janthinobacterium lividum on the salamander Plethodon cinereus". J. Chem. Ecol. 34 (11): 1422–9. doi:10.1007/s10886-008-9555-7. PMID 18949519.
- ^ Brucker RM, Baylor CM, Walters RL, Lauer A, Harris RN, Minbiole KP (January 2008). "The identification of 2,4-diacetylphloroglucinol as an antifungal metabolite produced by cutaneous bacteria of the salamander Plethodon cinereus". J. Chem. Ecol. 34 (1): 39–43. doi:10.1007/s10886-007-9352-8. PMID 18058176.
- ^ a b c d Harris R., Brucker R., Minbiole K., Walke J., Becker M., Schwantes C. et al. (2009). "Skin microbes on frogs prevent morbidity and mortality caused by a lethal skin fungus". ISME Journal 3 (7): 818–824. doi:10.1038/ismej.2009.27. PMID 19322245.
- ^ a b Becker M., Brucker R., Schwantes C., Harris R., Minbiole K. (2009). "The bacterially-produced metabolite violacein is associated with survival in amphibians infected with a lethal disease". AEM 21: 1294–1302.
- ^ Brucker R., Harris R., Schwantes C., Gallaher T., Flaherty D., Lam B. et al. (2008). "Amphibian Chemical Defense: Antifungal Metabolites of the Microsymbiont Janthinobacterium lividum on the Salamander Plethodon cinereus". Springer 34: 1422–1429.
- ^ a b Lam B., Walke J., Vredenburg V., Harris R. (2009). "Proportion of individuals with anti-Batrachochytrium dendrobatidis skin bacteria is associated with population persistence in the frog Rana muscosa". Biological Conservation 143: 529–531. doi:10.1016/j.biocon.2009.11.015.
- ^ Cohen, Nathan W.; Stebbins, Robert A. (1995). A Natural History of Amphibians. Princeton, N.J: Princeton University Press. ISBN 0-691-10251-1.
- ^ Daividson C, Shaffer HB, Jennings MR (2001). "Declines of the california red-legged frog: climate, UV-B, habitat, and pesticides hypotheses". Ecological Applications 11 (2): 464–479. doi:10.1890/1051-0761(2001)011[0464:DOTCRL]2.0.CO;2.
- ^ Hayes TB, Case P, Chui S, Chung D, Haeffele C, Haston K, Lee M, Mai VP, Marjuoa Y, Parker J, Tsui M (April 2006). "Pesticide mixtures, endocrine disruption, and amphibian declines: are we underestimating the impact?". Environ. Health Perspect. 114 Suppl 1: 40–50. PMID 16818245. PMC 1874187. http://www.ehponline.org/docs/2006/8051/abstract.html.
- ^ Davidson C, Benard MF, Shaffer HB, Parker JM, O'Leary C, Conlon JM, Rollins-Smith LA (March 2007). "Effects of chytrid and carbaryl exposure on survival, growth and skin peptide defenses in foothill yellow-legged frogs". Environ. Sci. Technol. 41 (5): 1771–6. doi:10.1021/es0611947. PMID 17396672.