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Schilliger, L.; Paillusseau, C.; François, C.; Bonwitt, J. Fungal Pathogens. Encyclopedia. Available online: https://encyclopedia.pub/entry/42511 (accessed on 19 July 2025).
Schilliger L, Paillusseau C, François C, Bonwitt J. Fungal Pathogens. Encyclopedia. Available at: https://encyclopedia.pub/entry/42511. Accessed July 19, 2025.
Schilliger, Lionel, Clément Paillusseau, Camille François, Jesse Bonwitt. "Fungal Pathogens" Encyclopedia, https://encyclopedia.pub/entry/42511 (accessed July 19, 2025).
Schilliger, L., Paillusseau, C., François, C., & Bonwitt, J. (2023, March 24). Fungal Pathogens. In Encyclopedia. https://encyclopedia.pub/entry/42511
Schilliger, Lionel, et al. "Fungal Pathogens." Encyclopedia. Web. 24 March, 2023.
Fungal Pathogens
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This entry is adapted from the peer-reviewed paper 10.3390/pathogens12030429

Emerging infectious diseases (EIDs) are caused by pathogens that have undergone recent changes in terms of geographic spread, increasing incidence, and expanding host range, or by previously unknown pathogens that are being discovered thanks to advances in surveillance and research, particularly in the field of laboratory diagnostics.

reptile amphibian emerging infectious diseases nannizziomycosis

1. Reptiles

Most fungal diseases of reptiles were originally grouped under a fungal complex named Chrysosporium anamorph of Nannizziopsis vriesii (CANV). Many of the pathogens that constituted this group have since been identified. The CANV denomination was abandoned in the 2010s for a new classification, including three genera belonging to the order Onygenales: Nannizziopsis spp., Paranannizziopsis spp. (Family: Nannizziopsiaceae), and Ophidiomyces spp. (Family Onygenaceae) [1][2][3].
Nannizziopsis spp. infection in reptiles was formerly known as “yellow fungus disease” (YFD), now replaced by nannizziomycosis (for Nannizziopsis spp. infection) and paranannizziomycosis (for Paranannizziopsis spp. infection) [4]. Similarly, Ophidiomyces ophidiicola (Oo) infection in snakes, formerly known as “snake fungal disease” (SFD), has been replaced by ophidiomycosis [4]. SFD is still used to describe a broad set of clinical signs, whereas ophidiomycosis should be restricted to confirmed Oo infection. These dermatomycoses can be grouped under the term “onygenalean dermatomycosis” [4]. Researcers focus on nannizziomycosis and ophidiomycosis as the most common fungal diseases of captive and wild reptiles worldwide [4][5][6].

1.1. Nannizziopsis spp.

Nannizziomycosis was first described in 1991 in day geckos (Phelsuma sp.) and later in 1997 in three different species of captive chameleons (Calumma parsonii, Chamaeleo lateralis, and C. jacksoni), with CANV attributed as the causative agent [7]. The disease was later named YFD after being reported in three captive inland bearded dragons (Pogona vitticeps) with deep granulomatous dermatomycosis and yellow discoloration of the epidermis [8]. Nannizziomycosis has since been reported in several species of lizards and crocodilians, and it is now recognized as an emerging disease in both wild and captive animals [9]. Cases of Nannizziopsis spp. infection in captive reptiles have been reported in Africa, Asia, Europe, North America, Australia, and New Zealand [1][2][10][11]. The most commonly affected species in captivity are bearded dragons, although other agamids and iguanids are also susceptible [2]. Nannizziopsis guarroi is the most frequently reported Nannizziopsis species in bearded dragons, but infection with N. chlamydospora, N. draconii, and N. barbatae have also been reported as a cause of nannizziomycosis in bearded dragons [10][12][13]. Several cases of cutaneous and systemic infection involving lungs and kidneys attributed to N. dermatidis have been reported in chameleons and geckos [10]. Nannizziopsis crocodili was first isolated in 1994 and 1997 in saltwater crocodiles (Crocodylus porosus) farms; forty-eight hatchlings died in two outbreaks, suggesting a possible age predisposition [14]. More recently, N. crocodili was identified from biopsied tissue in a captive juvenile Johnston’s crocodile (C. johnstoni) during an outbreak of severe multifocal dermatitis, affecting four of five crocodiles, in which lesions progressed from superficial ulcerations to black pigmentation and localized edema [15]. Of major concern has been the first report of N. barbatae in wild animals in 2020, involving four lizard species in Australia, all of which were found dead [11].
Nannizziomycosis begins with initial hyphae proliferation in the outer epidermal stratum corneum, with subsequent invasion of the deeper epidermal strata and dermis. A spectrum of lesions is usually observed, ranging from liquefactive necrosis of the epidermis to granulomatous inflammation in the dermis [16]. In bearded dragons, clinical signs include crusting dermatitis of the face, ventral surface of the limbs, and pericloacal region [13][17][18]. The crusts present with a yellow to brown appearance (Figure 1). Nonspecific clinical signs include molting retention, lethargy, and anorexia. Infection eventually leads to granulomatous inflammation and visceral dissemination, resulting in a poor prognosis [14].
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Figure 1. Nannizziomycosis in a green iguana (Iguana iguana) presenting with widespread cutaneous lesions of the flank.
The thermotolerance of reptile-associated Nannizziopsis spp. infection is highly dependent on the species involved. N. chlamydospore, for example, is moderately inhibited at 35 °C, whereas N. barbatae does not grow above 35 °C, and N. guarroi grows faster at 35 °C than at 30 °C [2][10][11][14]. Definitive diagnosis of nannizziomycosis is based on both demonstration of fungal elements in affected tissue (via histopathology) and identification of the organism on culture, PCR, or whole genome sequencing [14]. Fungal culture can take up to three weeks, and precise species identification can be challenging.
No standard treatment protocols exist, and the prognosis is guarded because recurrence is commonly encountered. Antifungal susceptibility of reptile-associated Nannizziopsis species has been poorly described and is often limited to case reports [14]. Voriconazole and terbinafine show good activity against the main Nannizziopsis species, even though resistance is being increasingly described [2][14][19][20][21][22]. Nannizziomycosis treatment consists of wound trimming, systemic appropriate antifungal treatment, and administration of analgesics associated with strict disinfection procedures (materials and enclosure) [2][17][18][20][21]. Animals with confirmed nannizziomycosis or compatible clinical signs should be quarantined and only reintroduced into a captive collection upon a negative PCR and histopathological test. These tests, however, have a low sensitivity in asymptomatic animals, meaning that the introduction of carriers in a naive population cannot be fully excluded. There is a paucity of information regarding environmental disinfection for Nannizziopsis sp., but a minimum of two minutes contact with 10% bleach seems to be effective against N. guarroi [23]. When handling animals with suspected or confirmed infection, it is recommended to dip gloved hands into disinfectant between handling, allowing sufficient contact time to the disinfectant, followed by rinsing with clean water [24].

1.2. Ophidiomyces ophidiicola

Ophidiomyces ophidiicola (Oo) was first described in 2009 as Chrysosporium ophidiicola in a black rat snake (Pantherophis obsoletus) presenting severe subcutaneous facial swelling, causing displacement of cranial anatomical features [25]. Several cases of snake dermatomycosis were retrospectively identified as ophidiomycosis, including a fatal case in 1990 in wild-caught brown tree snakes (Boiga irregularis) imported from Guam to Maryland, United States [26] and free-ranging pygmy rattlesnakes (Sistrurus catenatus) in Florida, United States in the 1990s [27]. The presence of Oo has been retrospectively demonstrated in museum specimens in the United States as early as 1945 [28][29] and in Europe as early as 1959 [30]. Molecular-based investigations suggest that strains of Oo in the eastern United States are primarily represented by four clonally expanded lineages or hybrids between those lineages, and the ancestors of these clonal lineages arrived in the region relatively recently, probably via the animal trade and via human-mediated transmission [31][32]. Ophidiomyces ophidiicola is considered responsible for wild population declines of Eastern Massasaugas rattlesnakes (Sistrurus catenatus) in Illinois and timber rattlesnakes (Crotalus horridus) in Massachusetts, United States [33][34]. Ophidiomycosis is thought to only affect snakes, with cases reported worldwide in colubrid snakes (e.g., Pantherophis sp., Nerodia sp., Natrix sp., Lampropeltis sp., Thamnophis sp.), viperids (e.g., Agkistrodon sp., Crotalus sp., Sistrurus sp.), Acrochordidae, and boids, elapids, and pythonids [14][35]. Ophidiomycosis is likely overreported in colubrids and viperids in comparison to other species, no doubt because of the important surveillance and research effort in North American snakes; the true number of susceptible species is likely more important than currently reported [10][29][31][33][36][37][38][39][40]. Ophidiomycosis was reported for the first time in Europe in 1985 in a captive ball python (Python regius) [10], in captive snakes in Japan [41], and in wild snakes in Hong Kong in 2019 [40] and in Taiwan in 2021 [39]. In North America (including Puerto Rico), Oo infection has been reported in at least 49 native snake species and in three non-native species [5].
Ophidiomyces ophidiicola is a saprobe with a particular tropism for keratinized environments. Growth is inhibited below 7 °C and above 35 °C, with an optimal temperature of 25 °C [37]. Lesions can be found anywhere along the body, but they initially present as pustular and then crusting dermatitis, involving the face, precorneal scales, thermosensitive dimples, ventral body surface, and the pericloacal region [18] (Figure 2). Regional swelling, edema, and vesiculation may be visible, eventually leading to ulceration. Lesions result in dysecdysis and increased molting frequency. Nonspecific signs include lethargy and anorexia. Infection can progress to underlying tissues including bones, muscle, and viscera. Granulocytic inflammation, edema, and necrosis of the epidermis extending to the dermis are visible on histology. In the wild, ophidiomycosis typically causes a pustular dermatosis in snakes emerging from brumation [2][36][39]. Shedding can reduce or even clear skin lesions [42], which may result in asymptomatic carriers [43][44]. Other fungal infections, such as Paranannizziopsis spp., can be confused with Oo infection, which can complicate the diagnosis [5][45]. Ophidiomycosis can be classified as possible, apparent, or confirmed, depending on clinical signs, laboratory testing, and demonstration of fungal hyphae on histopathology [45]. Transmission occurs via direct contact with infected individuals or via fomites [38]. Vertical transmission has also been documented [46].
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Figure 2. Ophidiomycosis in a ball python (Python regius) presenting severe ventral lesions of the epidermis.
As with nannizziomycosis, the diagnosis of ophidiomycosis is based on both demonstration of tissue involvement and pathogen identification [14]. Medical management is the same as for nannizziomycosis; Oo is susceptible to itraconazole, voriconazole, and terbinafine [2][34]. Subcutaneous terbinafine implants with a release over five weeks are being studied for managing venomous snakes [47]. Nebulized terbinafine is also of interest, with therapeutic plasma concentration possibly reached between half an hour and four hours, although efficacy studies are needed [47]. In infections involving captive animals, the environment can be disinfected with common disinfectants, including 3 to 10% bleach or 70% ethanol due to shedding of spores in the environment [24][48]. To avoid contaminating native fauna, snakes should never be released into the wild without first confirming freedom from infection, although this can be challenging because of the relatively low test sensitivity in asymptomatic snakes [43].

2. Amphibians

2.1. Chytridiomycosis

Chytridiomycosis is a fungal disease of amphibians attributed to two pathogenic species of the Chytridiomycete class: Batrachochytrium dendrobatidis (Bd) and B. salamandrivorans (Bsal) [49][50][51][52]. Both species differ mainly in terms of their lifecycle (especially temperature and pH requirements), host species, and clinical signs [49][50][53].
Batrachochytrium spp. are primitive fungi that inhabit wetlands and aquatic environments [49][50][51][52]. Both species multiply asexually and have an evolutionary cycle consisting of two stages, the motile infectious stage (zoospores) and the immotile reproductive stage (thallus) [52]. The zoospores can move at a speed of about two cm per day in stagnant water, a speed that can be greatly increased in the presence of water currents [54][55]. A second type of non-motile and floating zoospore is produced by Bsal. These spores remain infective for over 30 days in pond water and up to 48 h in soil [56]. Following adhesion to the host integument, the zoospore flagellum is resorbed, and a cystic wall is formed [52]. The lifecycle of Bsal is complete within five days at 15 °C [57]. The encysted zoospores of Bd mature in the zoosporangium and then the thallus for a period of four to five days (at 22 °C in vitro) [58]. A notable difference between the two chytrid species is that Bsal continues to divide in the encysted zoospore stage, thereby releasing a large number of zoospores from the thallus [50]. Bd is a non-obligate parasite that can survive as a saprobionte in water and moist soil for up to several months [50]. Some isolates of Bsal are able to synthesize molecules that enable it to survive as a saprophyte, conferring it the ability to withstand prolonged periods without hosts [59]. Batrachochytrium spp. are extremely vulnerable to desiccation.
Amphibian chytrids colonize the keratinized layers of the epidermis (stratum corneum and stratum granulosum) in adults (Bsal and Bd) and the mouthparts in tadpoles (Bd only), thereby disrupting osmoregulation, respiration, and foraging activity [49][52][60]. Once transcutaneous ion exchange is impeded, metabolite imbalances (hyponatremia, hypokalemia, hypochloremia, and hypocalcemia) cause decreased plasma osmolarity, cardiac pathologies, and death [49][50].
Diagnosis is obtained by PCR (or qPCR) of skin swabs or skin biopsies. Cytological examination of skin scrapping or histological examination of skin tissues are also possible, but are less sensitive than PCR [61]. Surveillance in wildlife can be achieved using PCR on environmental DNA (eDNA) [62][63][64][65][66]. Multiple treatment regimens have been described in captivity, including treating water with chloramphenicol malachite green or methylene blue [67], altering the skin microbiota by using probiotics [68][69][70], or increasing the environmental temperature [71][72][73][74][75]. Antifungal therapy (e.g., itraconazole, voriconazole, polymyxin E, or terbinafine) paired with non-steroidal anti-inflammatories are another option [50][51]. Additional microbiological testing can be required to treat superinfections [51]. Zoospores can easily be eliminated by desiccation, UV exposure, heat (4 h at 37 °C), or 5% sodium chloride [76]. Prevention in amphibians destined for trade involves strict quarantine of captive hosts for 60 days with entry and exit PCR testing [77][78]. Outdoor areas are particularly difficult to protect from disease incursion because zoospores can travel in moving water [54] or be transmitted through fomites, for example on the feathers and interdigital skin of aquatic birds [56][79][80] or fomite transmission via anthropogenic activities (e.g., movements of vehicles and equipment, or via footwear) [81][82]. Strict hygiene procedures are therefore required to prevent pathogen translocation [24]; these include cleaning and disinfection of footwear, tires, and other potentially contaminated surfaces [24]. Contact with 70% ethanol (one minute contact time) or 5% bleach (5–15 min contact time) is sufficient to inactivate Bd [24]. Non-powdered or vinyl gloves used to handle infected individuals can be disinfected using the same protocol described for Oo [24].

Batrachochytrium dendrobatidis

Bd was first described in 1998 from dead wild anurans collected in Australia in 1993 and Panama in 1994 [83]. Recent studies suggests that East Asia could be the original source of the panzootic lineage [84]. Retrospective investigations of archival samples have revealed its presence in the United States (since 1888) [85], Brazil (since 1894) [86], Asia (since 1902) [87], Africa (since 1933) [88], Canada (since 1961) [89], and Europe (in 1997) [90]. Bd has since been reported worldwide in over 1375 species, including in anurans, caudates, and caecilians [50][52][91][92][93]. Molecular investigations suggest that it appears to have been stable in wild populations for many decades, after which it spread globally, most likely as a consequence of the global trade in wild animals [85][94], notably that of the African clawed frog (Xenopus laevis) and bullfrog (Lithobates catesbeianus) [95][96][97][98][99]. Bd is responsible for the decline of amphibian populations around the world, mostly in tropical regions of Africa and South America, but also in Australia and southern Europe [100]. The harlequin frog (Atelopus varius), for example, has undergone >90% population declines over the past 10 years, while other species, such as the golden toad (Incilius periglenes) and Panamanian golden frog (Atelopus zeteki), have gone extinct in the wild [101].
Bd can be classified according to lineages, including the very high pathogenic Bd-GPL (global pandemic lineage) and endemic lineages, including Bd-Cape, Bd-Brazil, Bd-Asia, and Bd-CH [102]. This classification is constantly evolving as new lineages and genotypes are being discovered, including recent evidence of hybridization [103][104].
Growth is inhibited below 10 °C or above 28 °C, with an optimal temperature of 17–25 °C and a pH of 6–7 [50]. In tadpoles, infection is usually limited to the beak, manifested by depigmentation of the mouth and its periphery [83]. This results in decreased food intake and growth, as well as limited swimming ability [60]. In contrast to adults, infection in tadpoles is rarely life-threatening because their integument contains very little keratin. In adults, signs are highly variable and nonspecific. Affected animals may be asymptomatic, lethargic, anorexic, or present with neurological disorders (e.g., abolition of the reversal reflex, ataxia, convulsive, and seizures). Sudden death without overt signs can occur [50]. Dermatologic signs include increased molting frequency associated with hyperkeratosis, hyperplasia (up to 30 times the normal thickness), erythema, and discoloration of the skin [50]. The lesions are typically located on the ventral body surface (mainly on the pelvic patch), hindlimbs, and fingers in anurans (Figure 3) [50][58].
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Figure 3. Typical chytridiomycosis lesions in a green tree frog (Dryopsophus caeruleus). (a) Thickened skin on the ventrum; (b) excessive skin shedding on the feet. Photos courtesy K. Wright, In: Mader and Divers (eds). Current Therapy in Reptile Medicine and Surgery, Elsevier, 2014.

Batrachochytrium salamandrivorans

Bsal was first described in the Netherlands in 2013 and is responsible for the decline of >99% of wild fire salamanders (Salamandra salamandra) in some areas of Europe [49][57][105]. Bsal has been detected in captive newts and salamanders in Germany, Spain, and the United Kingdom [106][107][108]. It appears to be limited to European salamanders and newts [50][105][109], while native wild Asian urodeles are suspected to act as asymptomatic reservoirs [105][110][111][112]. A list of amphibian species according to Bsal susceptibility (susceptible, asymptomatic carrier, resistant) and geographic location is available elsewhere [113]. Bsal chytridiomycosis is, however, not limited to urodele species. Although laboratory studies have demonstrated the inability of Bsal to infect caecilians when placed in contact with 10,000 zoospores for 24 h [49][105], common midwife toads (Alytes obstetricans) from Europe are susceptible to Bsal when exposed to high loads of zoospores (contact with 100,000 zoospores for 24 h), suggesting a potential role of anurans in the pathogen lifecycle [56][114]. Cuban treefrogs can be infected with Bsal and, surprisingly, chytridiomycosis can develop in animals at the two highest zoospore dose exposures [115]. Moreover, different strains of Bsal might account for variations in susceptible species and epidemic profile, as was hypothesized following the isolation of Bsal in wild small-webbed fire-bellied toads (Bombina microdeladigitora) from Vietnam [110][116]. Current bans on amphibian transport that largely focus on halting the trade of urodele species may, therefore, be insufficient to prevent translocation of Bsal, especially as anurans constitute 99% of global amphibian trade [117].

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