Paracoccidioides brasiliensis: Infectious substances pathogen safety data sheet

Section I – Infectious agent

Name

Paracoccidioides brasiliensis

Agent type

Fungus

Taxonomy

Family

Ajellomycetaceae

Genus

Paracoccidioides

Species

Brasiliensis

Synonym or cross-reference

Zymonema brasiliense; Mycoderma brasiliense; Coccidioides brasiliensis; Monilia brasiliensis; Blastomyces brasiliensis; Aleurisma brasiliense; paracoccidioidomycosis; South American blastomycosis; Lutz's mycosis^{Footnote 1}^{Footnote 2}.

Characteristics

Brief description

Paracoccidioides brasiliensis is the causative agent of a systemic mycosis termed paracoccidioidomycosis (PCM). P. brasiliensis is a thermally dimorphic fungus: it grows as a white mould at 4 to 25 °C and a yeast at 35 to 37 °C^{Footnote 3}. The saprophytic mycelial form is composed of septate hyphae that produce infectious propagules called conidia, which undergo a thermally regulated transition to the yeast form upon infection of host tissues^{Footnote 3}^{Footnote 4}. Yeast cells are round or oval-shaped and vary in size from 4 to 40 µm^{Footnote 3}. Genomic DNA ranges from 26 to 36 Mbp in size and is arranged in four or five chromosomes^{Footnote 5}^{Footnote 6}.

Properties

The P. brasiliensis species complex comprises at least five phylogenetic groups: S1a, S1b, PS2, PS3, and PS4^{Footnote 7}^{Footnote 8}^{Footnote 9}. Reclassification of the cryptic species PS2, PS3, and PS4 as species P. americana, P. restrepiensis, and P. venezuelensis, respectively, was recently proposed^{Footnote 8}^{Footnote 9}.

Yeast forms of P. brasiliensis reproduce asexually by multiple budding. Mother cells are typically surrounded by multiple daughter cells (i.e., blastoconidia) in a configuration resembling a "pilot wheel"^{Footnote 3}^{Footnote 4}. Genetic evidence suggests that a sexual stage may also occur in the P. brasiliensis life cycle, but this teleomorph has not been isolated^{Footnote 10}^{Footnote 11}.

Section II – Hazard identification

Pathogenicity and toxicity

Disease associated with P. brasiliensis infection can range from asymptomatic to severe systemic illness^{Footnote 7}. Infection results from inhalation of P. brasiliensis conidia, which migrate to pulmonary alveoli where transformation to the yeast form occurs^{Footnote 12}. In the lungs, P. brasiliensis infection is controlled by the host cellular immune response in most individuals, resulting in asymptomatic or mild, self-limiting respiratory infection^{Footnote 13}^{Footnote 14}. In a small percentage of infected individuals, inability to control the infection results in progression to clinical disease.

Acute clinical manifestation includes lymph node enlargement in nearly all cases, cutaneous or mucous membrane lesions (17-20% of cases), and pulmonary involvement (5-10% of cases), often accompanied by fever and weight loss^{Footnote 4}^{Footnote 7}. Acute PCM progresses over weeks and can disseminate to other parts of body^{Footnote 4}^{Footnote 7}^{Footnote 14}. Acute PCM can be moderate or severe and usually occurs in children, adolescents, and young adults^{Footnote 4}^{Footnote 13}. The estimated relapse rate was 7.8% and the mortality rate was 5.7% for acute PCM^{Footnote 15}. Latent P. brasiliensis cells that were not cleared during initial infection can reactivate years later resulting in a chronic form of PCM, which can be mild, moderate, or severe^{Footnote 4}^{Footnote 7}^{Footnote 13}. Chronic PCM usually occurs in adults over 30 years of age and predominantly affects males^{Footnote 3}^{Footnote 16}. Disease develops slowly and symptoms usually persist for more than six months^{Footnote 7}. Clinical manifestations include lesions in lungs (90% of cases), mucosa of the upper aerodigestive tract, and skin near the mouth and nose, accompanied by malaise, anorexia, and nausea^{Footnote 4}^{Footnote 7}^{Footnote 13}. Dissemination to other organs, including the adrenal glands, central nervous system (CNS), and bones, can occur in cases of severe chronic PCM^{Footnote 7}. It is estimated that 3-5% of PCM cases are fatal^{Footnote 3}^{Footnote 13}.

PCM sequelae include dysphonia, adrenal impairment in adults, pulmonary fibrosis, and emphysema^{Footnote 4}^{Footnote 11}^{Footnote 17}. In disseminated PCM cases involving the CNS, neurological sequelae are common^{Footnote 7}. Scarring of skin and oral mucosa can be severe.

There are relatively few reports of clinical disease associated with P. brasiliensis in animals, despite serological and molecular evidence of P. brasiliensis infection in many mammalian species^{Footnote 4}. PCM has been reported in dogs^{Footnote 18}^{Footnote 19}^{Footnote 20}, cats^{Footnote 21}, and a two-toed sloth^{Footnote 22}. A novel strain of P. brasiliensis was isolated from bottlenose dolphins with cutaneous granulomas^{Footnote 23}.

Epidemiology

P. brasiliensis is endemic to southern Mexico, Central America, and parts of South America^{Footnote 13}^{Footnote 24}. P. brasiliensis genotypes S1a and S1b are found in Southeast Brazil, Argentina and Paraguay; PS2 (P. americana) in parts of Brazil and Venezuela; PS3 (P. restrepiensis) in Colombia; and PS4 (P. venezuelensis) in Venezuela^{Footnote 4}^{Footnote 7}^{Footnote 8}^{Footnote 9}. In PCM-endemic regions, prevalence of P. brasiliensis infection in the population ranges from 4 to 47%^{Footnote 13}^{Footnote 25}^{Footnote 26}. Approximately 2% of individuals exposed to P. brasiliensis develop PCM^{Footnote 4}. Acute PCM and chronic PCM comprise 5-25% and 75-95% of cases, respectively^{Footnote 7}^{Footnote 13}. Approximately 10-15% of PCM cases occur in children and adolescents^{Footnote 3}^{Footnote 7}. Although infection rates are similar among males and females^{Footnote 4}, approximately 75% to 95% of PCM cases in adults are men^{Footnote 7}^{Footnote 13}^{Footnote 16}.

Approximately 80% of reported PCM cases occur in Brazil^{Footnote 7}^{Footnote 13}. Annual PCM incidence in endemic areas of Brazil is estimated at 0.71 to 3.7 cases per 100,000 inhabitants^{Footnote 13}. In hyperendemic areas of Brazil, annual incidence can be up to 40 cases per 100,000 inhabitants^{Footnote 27}. In Colombia, estimated annual incidence is less than 0.24 cases per 100,000 inhabitants^{Footnote 28}.

Approximately 80 PCM cases have been reported in non-endemic regions, affecting individuals who previously visited or resided in PCM-endemic areas^{Footnote 3}^{Footnote 13}^{Footnote 29}.

While intradermal testing indicates that P. brasiliensis infection rates are similar among males and females, PCM predominantly affects males^{Footnote 3}^{Footnote 4}^{Footnote 7}. Females produce 17β-estradiol, which inhibits transformation of conidia to yeast, thereby promoting resistance to disease^{Footnote 12}. Approximately 73% of PCM patients have a history of agriculture-related occupations^{Footnote 3}. Smoking and alcohol consumption are associated with development of chronic PCM^{Footnote 30}^{Footnote 31}. PCM mortality rate is higher for immunosuppressed individuals^{Footnote 32}.

Host range

Natural host(s)

Humans and armadillos are the primary hosts of P. brasiliensis^{Footnote 9}^{Footnote 13}. There are a few reported cases of PCM in dogs^{Footnote 18}^{Footnote 19}^{Footnote 20}, a cat^{Footnote 21}, squirrel monkey^{Footnote 33}, and two-toed sloth^{Footnote 22}. Serological, histological, and molecular evidence indicates that P. brasiliensis can infect a wide range of hosts including bats^{Footnote 34}, rodents^{Footnote 35}, equines^{Footnote 36}, sheep^{Footnote 37}, cattle^{Footnote 38}, pigs^{Footnote 39}, goats^{Footnote 40}, chickens^{Footnote 41}, rabbits^{Footnote 42}, opossums^{Footnote 43}, fish^{Footnote 44}, guinea pigs^{Footnote 45}, raccoons^{Footnote 45}, porcupines^{Footnote 45}, grison^{Footnote 45}, and anteaters^{Footnote 46}.

Other host(s)

Experimental hosts include hamsters^{Footnote 47} and moths (i.e., Galleria mellonella)^{Footnote 48}.

Infectious dose

Unknown.

Incubation period

Acute PCM develops weeks to months after exposure to P. brasiliensis^{Footnote 7}^{Footnote 14}. Chronic PCM develops years to decades after exposure to P. brasiliensis^{Footnote 4}^{Footnote 7}.

Communicability

P. brasiliensis infection occurs primarily through inhalation of airborne conidia^{Footnote 3}^{Footnote 7}. Infection resulting from traumatic inoculation of skin or mucous membranes is possible, yet rare^{Footnote 49}^{Footnote 50}. P. brasiliensis has been transmitted via solid organ transplantation; however, the recipient of the P. brasiliensis-infected organ did not develop clinical PCM after one year of follow up^{Footnote 51}. Human-to-human transmission has not been reported.

Section III – Dissemination

Reservoir

Armadillos are known reservoirs of P. brasiliensis^{Footnote 4}^{Footnote 9}^{Footnote 52}.

Zoonosis

None.

Vectors

None.

Section IV – Stability and viability

Drug susceptibility/resistance

P. brasiliensis is susceptible to sulfonamide derivatives (e.g., co-trimoxazole, sulfadiazine, sulfamethoxypyridazine, sulfadoxine)^{Footnote 53}, azoles (e.g., ketoconazole, fluconazole, itraconazole, posaconazole, voriconazole, isavuconazole)^{Footnote 7}, amphotericin B^{Footnote 7}, and terbinafine^{Footnote 54}^{Footnote 55}.

Susceptibility to disinfectants

P. brasiliensis is susceptible to sodium hypochlorite (5%)^{Footnote 56}. Other fungi in the Ajellomycetaceae family are susceptible to phenolic compounds and glutaraldehyde^{Footnote 57}.

Physical inactivation

Gamma irradiation (6.5 kGy) inhibits growth of P. brasiliensis^{Footnote 58}. Yeast and mycelial forms of P. brasiliensis are inactivated at pH 3 and 1.4, respectively^{Footnote 59}. Other fungi in the Ajellomycetaceae family are inactivated by moist heat treatment at 121 °C for 30 min^{Footnote 57}.

Survival outside host

In nature, the saprophytic mycelial form of P. brasiliensis presumably occupies soil in PCM endemic regions; however, few attempts to isolate the fungus from soil have been successful^{Footnote 60}. P. brasiliensis suspended in water at room temperature can survive for several months^{Footnote 61}^{Footnote 62}.

P. brasiliensis survival was adversely affected at -20°C and 45°C in soil extracts and by oxygen deprivation in liquid medium^{Footnote 59}^{Footnote 63}.

Section V – First aid/medical

Surveillance

P. brasiliensis is identified in clinical samples by direct visualisation of multiple-budding cells in a "pilot's wheel" configuration, which are characteristic of the yeast form of P. brasiliensis^{Footnote 4}^{Footnote 64}. P. brasiliensis can be isolated from clinical specimens and cultured but growth is slow (20 to 30 days)^{Footnote 3}. P. brasiliensis antibodies can be detected in serum using double agar gel immunodiffusion test, enzyme-linked immunosorbent assay (ELISA), and immunoblot^{Footnote 4}^{Footnote 7}.

P. brasiliensis antigen can be detected in bronchoalveolar lavage fluid, serum, and cerebrospinal fluid using ELISA^{Footnote 3}^{Footnote 4}. Molecular methods including PCR and sequencing can be used to detect P. brasiliensis in clinical samples^{Footnote 4}^{Footnote 65}^{Footnote 66}.

Note: The specific recommendations for surveillance in the laboratory should come from the medical surveillance program, which is based on a local risk assessment of the pathogens and activities being undertaken, as well as an overarching risk assessment of the biosafety program as a whole. More information on medical surveillance is available in the Canadian Biosafety Handbook (CBH).

First aid/treatment

PCM can be treated with an appropriate antifungal agent. Prognosis improves with early diagnosis and treatment. Duration of antifungal treatment for mild and moderate forms varies from 9 to 18 months (average 12 months)^{Footnote 7}. Treatment of severe forms involves induction therapy with amphotericin B for 2 to 4 weeks followed by maintenance treatment with oral antifungal medication for up to 24 months^{Footnote 7}. Oral lesions have been successfully treated with photodynamic therapy consisting of topical toluidine blue dye and low-level irradiation^{Footnote 67}^{Footnote 68}. Dogs with PCM have also been treated successfully with itraconazole^{Footnote 18}^{Footnote 19}.

Note: The specific recommendations for first aid/treatment in the laboratory should come from the post-exposure response plan, which is developed as part of the medical surveillance program. More information on the post-exposure response plan can be found in the CBH.

Immunization

No vaccine is currently available. Vaccine research is ongoing^{Footnote 69}^{Footnote 70}.

Note: More information on the medical surveillance program can be found in the CBH, and by consulting the Canadian Immunization Guide.

Prophylaxis

Post-exposure prophylaxis with itraconazole for 30 days is recommended^{Footnote 7}.

Note: More information on prophylaxis as part of the medical surveillance program can be found in the CBH.

Section VI – Laboratory hazard

Laboratory-acquired infections

A researcher developed localized infection after an accidental needlestick injury with a syringe containing the yeast form of P. brasiliensis^{Footnote 50}. The patient responded favourably to antifungal treatment and was discharged from treatment after 3 months^{Footnote 50}.

Note: Please consult the Canadian Biosafety Standard (CBS) and CBH for additional details on requirements for reporting exposure incidents. A Canadian biosafety guideline describing notification and reporting procedures is also available.

Sources/specimens

Specimens may include bronchoalveolar lavage fluid, sputum, scrapings of skin and mucosal lesions, lymph node aspirate, tissue biopsy, serum, urine, and cerebrospinal fluid^{Footnote 4}^{Footnote 7}.

Primary hazards

Exposure to P. brasiliensis via inhalation of airborne infectious material or traumatic inoculation (e.g., needlestick injury)^{Footnote 50}.

Special hazards

None.

Section VII – Exposure controls/personal protection

Risk group classification

P. brasiliensis is a Risk Group 3 human pathogen and Risk Group 1 animal pathogen^{Footnote 71}^{Footnote 72}.

Containment requirements

Containment Level 3 facilities, equipment, and operational practices, as outlined in the CBS, are required for work involving infectious or potentially infectious materials, animals, or cultures.

Protective clothing

The applicable Containment Level 3 requirements for personal protective equipment and clothing as outlined in the CBS, are to be followed. At minimum, it is recommended to use full body coverage dedicated protective clothing, dedicated protective footwear and/or additional protective footwear, gloves when handling infectious materials or animals, face protection when there is a known or potential risk of exposure to splashes or flying objects, respirators when there is a risk of exposure to infectious aerosols, and an additional layer of protective clothing prior to work with infectious materials or animals.

Note: A local risk assessment will identify the appropriate hand, foot, head, body, eye/face, and respiratory protection equipment required. The personal protective equipment requirements for the containment zone must be documented.

Other precautions

All activities involving open vessels of infectious material are to be performed in a certified biological safety cabinet (BSC) or other appropriate primary containment device. The use of needles, syringes, and other sharp objects should be strictly limited. Additional precautions should be considered with work involving animals or large scale activities.

Section VIII – Handling and storage

Spills

Allow aerosols to settle. Wearing protective clothing, gently cover the spill with absorbent paper towel and apply suitable disinfectant, starting at the perimeter and working towards the centre. Allow sufficient contact time with disinfectant before clean up (CBH).

Disposal

All materials/substances that have come in contact with the infectious agent should be completely decontaminated before they are removed from the containment zone. This can be achieved by using decontamination technologies and processes that have been demonstrated to be effective against the infectious material, such as chemical disinfectants, autoclaving, irradiation, incineration, an effluent treatment system, or gaseous decontamination (CBH).

Storage

The applicable Containment Level 3 requirements for storage outlined in the CBS are to be followed. Containers of infectious material or toxins stored outside the containment zone must be labelled, leakproof, impact resistant, and kept in locked storage equipment and within an area with limited access (CBH).

Section IX – Regulatory and other information

Canadian regulatory information

Controlled activities with P. brasiliensis require a Human Pathogens and Toxins licence, issued by the Public Health Agency of Canada^{Footnote 71}.

The following is a non-exhaustive list of applicable designations, regulations, or legislations:

Last file update

2020

Prepared by

Centre for Biosecurity, Public Health Agency of Canada.

Disclaimer

The scientific information, opinions, and recommendations contained in this Pathogen Safety Data Sheet have been developed based on or compiled from trusted sources available at the time of publication. Newly discovered hazards are frequent and this information may not be completely up to date. The Government of Canada accepts no responsibility for the accuracy, sufficiency, or reliability or for any loss or injury resulting from the use of the information.

Persons in Canada are responsible for complying with the relevant laws, including regulations, guidelines and standards applicable to the import, transport, and use of pathogens in Canada set by relevant regulatory authorities, including the Public Health Agency of Canada, Health Canada, Canadian Food Inspection Agency, Environment and Climate Change Canada, and Transport Canada. The risk classification and related regulatory requirements referenced in this Pathogen Safety Data Sheet, such as those found in the Canadian Biosafety Standard, may be incomplete and are specific to the Canadian context. Other jurisdictions will have their own requirements.

References:

Footnote 1

Species Fungorum. Paracoccidioides brasiliensis (Splend.) F.P. Almeida.

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Footnote 2

Negroni, R. 1993. Paracoccidioidomycosis (South American blastomycosis, Lutz's mycosis). Int. J. Dermatol. 32:847-859.

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Footnote 3

Restrepo, A. M., A. M. Tobón Orozco, B. L. Gómez, and G. Benard. 2015. Paracoccidioidomycosis, p. 225-236. D. R. Hospenthal and M. G. Rinaldi (eds.), Diagnosis and Treatment of Fungal Infections. Springer International Publishing, Switzerland.

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Footnote 4

Mendes, R. P., R. d. S. Cavalcante, S. A. Marques, M. E. A. Marques, J. Venturini, T. F. Sylvestre, A. M. M. Paniago, A. C. Pereira, J. d. F. da Silva, A. T. Fabro, S. d. M. G. Bosco, E. Bagagli, R. C. Hahn, and A. D. Levorato. 2017. Paracoccidioidomycosis: Current perspectives from Brazil. The Open Microbiology Journal. 11:224-282.

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Footnote 5

Almeida, A. J., D. R. Matute, J. A. Carmona, M. Martins, I. Torres, J. G. McEwen, A. Restrepo, C. Leão, P. Ludovico, and F. Rodrigues. 2007. Genome size and ploidy of Paracoccidioides brasiliensis reveals a haploid DNA content: Flow cytometry and GP43 sequence analysis. Fungal Genetics and Biology. 44:25-31.

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Footnote 6

Desjardins, C. A., M. D. Champion, J. W. Holder, A. Muszewska, J. Goldberg, A. M. Bailão, M. M. Brigido, M. E. da Silva Ferreira, A. M. Garcia, M. Grynberg, S. Gujja, D. I. Heiman, M. R. Henn, C. D. Kodira, H. León-Narváez, L. V. G. Longo, L. Ma, I. Malavazi, A. L. Matsuo, F. V. Morais, M. Pereira, S. Rodríguez-Brito, S. Sakthikumar, S. M. Salem-Izacc, S. M. Sykes, M. M. Teixeira, M. C. Vallejo, M. E. M. T. Walter, C. Yandava, S. Young, Q. Zeng, J. Zucker, M. S. Felipe, G. H. Goldman, B. J. Haas, J. G. McEwen, G. Nino-Vega, R. Puccia, G. San-Blas, C. M. A. de Soares, B. W. Birren, and C. A. Cuomo. 2011. Comparative genomic analysis of human fungal pathogens causing paracoccidioidomycosis. PLoS Genetics. 7:e1002345-e1002345.

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Footnote 7

Shikanai-Yasuda, M. A., R. P. Mendes, A. L. Colombo, F. d. Q. Telles, A. Kono, A. M. M. Paniago, A. Nathan, A. C. F. d. Valle, E. Bagagli, G. Benard, M. S. Ferreira, M. d. M. Teixeira, M. L. S. Vergara, R. M. Pereira, R. d. S. Cavalcante, R. Hahn, R. R. Durlacher, Z. Khoury, Z. P. d. Camargo, M. L. Moretti, and R. Martinez. 2018. Brazilian guidelines for the clinical management of paracoccidioidomycosis. Epidemiologia e Serviços De Saúde. 27:e0500001-e0500001.

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Footnote 8

Turissini, D. A., O. M. Gomez, M. M. Teixeira, J. G. McEwen, and D. R. Matute. 2017. Species boundaries in the human pathogen Paracoccidioides. Fungal Genetics and Biology. 106:9-25.

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Footnote 9

Hrycyk, M. F., H. Garcia Garces, S. d. M. G. Bosco, S. L. de Oliveira, S. A. Marques, and E. Bagagli. 2018. Ecology of Paracoccidioides brasiliensis, P. lutzii and related species: Infection in armadillos, soil occurrence and mycological aspects. Medical Mycology (Oxford). 56:950-962.

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Footnote 10

Teixeira, M. M., R. C. Theodoro, G. Nino-Vega, E. Bagagli, and M. S. S. Felipe. 2014. Paracoccidioides species complex: Ecology, phylogeny, sexual reproduction, and virulence. PLoS Pathogens. 10:e1004397-e1004397.

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Footnote 11

Teixeira, M., R. C. Theodoro, S. Derengowski L, A. M. Nicola, E. Bagagli, and M. S. Felipe. 2013. Molecular and morphological data support the existence of a sexual cycle in species of the genus Paracoccidioides. Eukaryot. Cell. 12:380-389.

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Footnote 12

Shankar, J., A. Restrepo, K. V. Clemons, and D. A. Stevens. 2011. Hormones and the resistance of women to paracoccidioidomycosis. Clin. Microbiol. Rev. 24:296-313.

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Footnote 13

Martinez, R. 2017. New trends in paracoccidioidomycosis epidemiology. Journal of Fungi (Basel). 3:1.

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Footnote 14

R. Buccheri, Z. Khoury, L. C. B. Barata, and G. Benard. 2016. Incubation period and early natural history events of the acute form of paracoccidioidomycosis: Lessons from patients with a single Paracoccidioides spp. exposure. Mycopathologia. 181:435-439.

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Footnote 15

Romaneli, M. T. d. N., N. R. Tardelli, A. T. Tresoldi, A. M. Morcillo, and R. M. Pereira. 2019. Acute‐subacute paracoccidioidomycosis: A paediatric cohort of 141 patients, exploring clinical characteristics, laboratorial analysis and developing a non‐survival predictor. Mycoses. 62:999-1005.

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Footnote 16

Blotta, M., R. L. Mamoni, S. J. Oliveira, S. A. Nouer, P. Papaiordanou, A. Goveia, and Z. P. d. Camargo. 1999. Endemic regions of paracoccidioidomycosis in Brazil: a clinical and epidemiologic study of 584 cases in the southeast region. Am. J. Trop. Med. Hyg. 61:390-394.

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Footnote 17

Pereira, R. M., G. Guerra-Júnior, and A. T. Tresoldi. 2006. Adrenal function in 23 children with paracoccidioidomycosis. Revista do Instituto De Medicina Tropical De São Paulo. 48:333-336.

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Footnote 18

De Farias, M. R., L. A. Z. Condas, M. G. Ribeiro, Bosco, Sandra de Moraes Gimenes, M. D. Muro, J. Werner, R. C. Theodoro, E. Bagagli, S. A. Marques, and M. Franco. 2011. Paracoccidioidomycosis in a dog: case report of generalized lymphadenomegaly. Mycopathologia. 172:147-152.

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Footnote 19

Headley, S. A., L. G. Pretto-Giordano, G. W. Di Santis, L. A. Gomes, R. Macagnan, D. F. Da Nóbrega, K. M. Leite, B. K. de Alcântara, E. N. Itano, and A. A. Alfieri. 2017. Paracoccidioides brasiliensis-associated dermatitis and lymphadenitis in a dog. Mycopathologia. 182:425-434.

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Footnote 20

Ricci, G., F. Mota, A. Wakamatsu, R. Serafim, R. Borra, and M. Franco. 2004. Canine paracoccidioidomycosis. Sabouraudia. 42:379-383.

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Footnote 21

Gonzalez, J. F., N. A. Montiel, and R. L. Maass. 2010. First report on the diagnosis and treatment of encephalic and urinary paracoccidioidomycosis in a cat. Journal of Feline Medicine and Surgery. 12:659-662.

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Footnote 22

Trejo-Chávez, A., R. Ramírez-Romero, J. Ancer-Rodríguez, A. Nevárez-Garza, and L. Rodriguez-Tovar. 2011. Disseminated paracoccidioidomycosis in a Southern two-toed sloth (Choloepus didactylus). J. Comp. Pathol. 144:231-234.

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Footnote 23

Bossart, G. D., P. Fair, A. M. Schaefer, and J. S. Reif. 2017. Health and Environmental Risk Assessment Project for bottlenose dolphins Tursiops truncatus from the southeastern USA. I. Infectious diseases. Dis. Aquat. Org. 125:141-153.

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Footnote 24

López‐Martínez, R., F. Hernández‐Hernández, L. J. Méndez‐Tovar, P. Manzano–Gayosso, A. Bonifaz, R. Arenas, M. d. C. Padilla‐Desgarennes, R. Estrada, and G. Chávez. 2014. Paracoccidioidomycosis in Mexico: clinical and epidemiological data from 93 new cases (1972–2012). Mycoses. 57:525-530.

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Footnote 25

Magalhães, Evandro Monteiro de Sá, C. d. F. Ribeiro, C. S. Dâmaso, L. F. L. Coelho, R. R. Silva, E. B. Ferreira, M. R. Rodrigues, Z. P. d. Camargo, T. R. G. Velloso, and L. C. C. Malaquias. 2014. Prevalence of paracoccidioidomycosis infection by intradermal reaction in rural areas in Alfenas, Minas Gerais, Brazil. Revista do Instituto De Medicina Tropical De São Paulo. 56:281-285.

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Footnote 26

Marques, Ana Paula da C, S. M. V. Oliveira, G. R. Rezende, D. A. Melo, S. M. Fernandes-Fitts, E. R. J. Pontes, M. da Glória Bonecini-Almeida, Z. P. Camargo, and A. M. Paniago. 2013. Evaluation of Paracoccidioides brasiliensis infection by gp 43 intradermal test in rural settlements in Central-West Brazil. Mycopathologia. 176:41-47.

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Footnote 27

Vieira, G. d. D., T. d. C. Alves, Lima, Sônia Maria Dias de, L. M. A. Camargo, and C. M. d. Sousa. 2014. Paracoccidioidomycosis in a western Brazilian Amazon State: clinical-epidemiologic profile and spatial distribution of the disease. Rev. Soc. Bras. Med. Trop. 47:63-68.

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Footnote 28

Torrado, E., E. Castañeda, F. de la Hoz, and A. Restrepo. 2000. Paracoccidioidomicocis: definición de las áreas endémicas de Colombia. Biomedica. 20:327-334.

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Footnote 29

Buitrago, M. J., L. Bernal‐Martínez, M. V. Castelli, J. L. Rodríguez‐Tudela, and M. Cuenca‐Estrella. 2011. Histoplasmosis and paracoccidioidomycosis in a non‐endemic area: a review of cases and diagnosis. Journal of Travel Medicine. 18:26-33.

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Footnote 30

Martinez, R., and M. J. Moya. 1992. The relationship between paracoccidioidomycosis and alcoholism. Rev. Saude Publica. 26:12-16.

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Footnote 31

dos Santos, W. A., B. M. da Silva, E. D. Passos, E. Zandonade, and A. Falqueto. 2003. Association between smoking and paracoccidioidomycosis: a case-control study in the State of Espírito Santo, Brazil. Cadernos De Saude Publica. 19:245-253.

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Footnote 32

de Almeida, J. N., P. M. Peçanha, and A. L. Colombo. 2019; 2018. Paracoccidioidomycosis in immunocompromised patients: A literature review. Journal of Fungi (Basel). 5:2.

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Footnote 33

Johnson, W., and C. Lang. 1977. Paracoccidioidomycosis (South American blastomycosis) in a squirrel monkey (Saimiri sciureus). Vet. Pathol. 14:368-371.

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Footnote 34

da Paz, G. S., B. M. V. Adorno, V. B. Richini‐Pereira, S. M. Bosco, and H. Langoni. 2018. Infection by Histoplasma capsulatum, Cryptococcus spp. and Paracoccidioides brasiliensis in bats collected in urban areas. Transboundary and Emerging Diseases. 65:1797-1805.

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Footnote 35

Losnak, D. O., F. R. Rocha, B. S. Almeida, K. Z. Batista, S. L. Althoff, J. Haupt, L. S. Ruiz, L. Anversa, S. B. Lucheis, and L. M. Paiz. 2018. Molecular detection of fungi of public health importance in wild animals from Southern Brazil. Mycoses. 61:455-463.

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Footnote 36

Albano, A. P. N., G. B. Klafke, T. M. Brandolt, V. P. Da Hora, C. E. W. Nogueira, M. O. Xavier, and M. C. A. Meireles. 2015. Seroepidemiology of Paracoccidioides brasiliensis infection in horses from Rio Grande do Sul, Brazil. Brazilian J. Microbiol. 46:513-517.

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Footnote 37

Oliveira, G. G., I. T. Navarro, R. L. Freire, D. R. Belitardo, L. H. Silveira, Z. P. Camargo, E. N. Itano, and M. A. Ono. 2012. Serological survey of paracoccidioidomycosis in sheep. Mycopathologia. 173:63-68.

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Footnote 38

Silveira, L., R. Paes, E. Medeiros, E. Itano, Z. Camargo, and M. Ono. 2008. Occurrence of antibodies to Paracoccidioides brasiliensis in dairy cattle from Mato Grosso do Sul, Brazil. Mycopathologia. 165:367-371.

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Footnote 39

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Ferreira, J. B., I. T. Navarro, R. L. Freire, G. G. Oliveira, A. M. Omori, D. R. Belitardo, E. N. Itano, Z. P. Camargo, and M. A. Ono. 2013. Evaluation of Paracoccidioides brasiliensis infection in dairy goats. Mycopathologia. 176:95-99.

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Oliveira, G. G., L. H. Silveira, E. N. Itano, R. M. Soares, R. L. Freire, M. A. Watanabe, Z. P. Camargo, and M. A. Ono. 2011. Serological evidence of Paracoccidioides brasiliensis infection in chickens from Paraná and Mato Grosso do Sul States, Brazil. Mycopathologia. 171:197-202.

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Belitardo, D. R., A. S. Calefi, M. R. Sbeghen, G. G. de Oliveira, M. A. E. Watanabe, Z. P. de Camargo, and M. A. Ono. 2014. Paracoccidioides brasiliensis infection in domestic rabbits (Oryctolagus cuniculus). Mycoses. 57:222-227.

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de Souza Suguiura, Igor Massahiro, R. Macagnan, A. M. Omori, E. L. Buck, J. A. Scarpassa, L. G. Pretto-Giordano, L. A. Vilas-Boas, Z. P. de Camargo, E. N. Itano, and M. A. Ono. 2020. First report of Paracoccidioides brasiliensis infection in fish. Medical Mycology. 58:737-743.

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Richini-Pereira, V. B., De Moraes Gimenes Bosco, Sandra, J. Griese, R. Cordeiro Theodoro, Assis Da Graça Macoris, Severino, R. José Da Silva, L. Barrozo, Morais E Silva Tavares, Patrícia, R. Maria Zancopé-Oliveira, and E. Bagagli. 2008. Molecular detection of Paracoccidioides brasiliensis in road-killed wild animals. Medical Mycology. 46:35-40.

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Richini-Pereira, V. B., S. M. Bosco, R. C. Theodoro, L. Barrozo, S. C. Pedrini, P. S. Rosa, and E. Bagagli. 2009. Importance of xenarthrans in the eco-epidemiology of Paracoccidioides brasiliensis. BMC Research Notes. 2:1-6.

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Essayag, S., M. Landaeta, C. Hartung, S. Magaldi, L. Spencer, R. Suárez, F. García, and E. Pérez. 2002. Histopathologic and histochemical characterization of calcified structures in hamsters inoculated with Paracoccidioides brasiliensis. Mycoses. 45:351-357.

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de Lacorte Singulani, J., L. Scorzoni, A. M. Fusco-Almeida, and M. J. S. Mendes-Giannini. 2016. Evaluation of the efficacy of antifungal drugs against Paracoccidioides brasiliensis and Paracoccidioides lutzii in a Galleria mellonella model. Int. J. Antimicrob. Agents. 48:292-297.

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Restrepo, A., M. Robledo, R. Giraldo, H. Herna, F. Sierra, F. Gutie, F. London, R. Lo, and G. Calle. 1976. The gamut of paracoccidioidomycosis. Am. J. Med. 61:33-42.

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Batista, M. V., P. K. Sato, L. C. Pierrotti, F. J. De Paula, G. F. Ferreira, D. S. Ribeiro-David, W. C. Nahas, M. I. S. Duarte, and M. A. Shikanai-Yasuda. 2012. Recipient of kidney from donor with asymptomatic infection by Paracoccidioides brasiliensis. Med. Mycol. 50:187-192.

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Corredor, G. G., J. H. Castaño, A. Paralta, S. Díez, M. Arango, J. McEween, and A. Restrepo. 1999. Isolation of Paracoccidioides brasiliensis from the nine-banded armadillo Dasypus novemcinctus, in an endemic area for paracoccidioidomycosis in Columbia. Rev. Iberoam. Micol. 16:216-220.

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Shikanai-Yasuda, M. A. 2015. Paracoccidioidomycosis treatment. Revista do Instituto De Medicina Tropical De São Paulo. 57:31-37.

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Shikanai-Yasuda, M. A., R. T. Taguchi, M. K. Sato, N. T. Melo, C. M. Assis, R. C. Nigro, E. E. Camargo, C. S. Lacaz, V. Amato Neto, and A. Sesso. 1991. In vitro action of some disinfectants on Paracoccidioides brasiliensis yeast forms. Revista do Instituto De Medicina Tropical De São Paulo. 33:37-43.

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Kowalski, W. J. 2012. Hospital airborne infection control. CRC Press, Boca Raton, FL.

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Restrepo, A., L. H. Moncada, and M. Quintero. 1969. Effect of hydrogen ion concentration and of temperature on the growth of Paracoccidioides brasiliensis in soil extracts. Sabouraudia. 7:207-215.

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de Bedout, C., L. E. Cano, A. M. Tabares, M. Van de Ven, and A. Restrepo. 1986. Water as a substrate for the development of Paracoccidiodes brasiliensis mycelial form. Mycopathologia. 96:123-130.

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Restrepo, A., B. E. Jiménez, and C. de Bedout. 1981. Survival of Paracoccidioides brasiliensis yeast cells under microaerophilic conditions. Sabouraudia. 19:301-305.

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Rocha-Silva, F., S. M. de Figueiredo, La Santrer, Emanoelle Fernandes Rutren, A. S. Machado, B. Fernandes, C. B. Assunção, A. M. Góes, and R. B. Caligiorne. 2018. Paracoccidioidomycosis: Detection of Paracoccidioides brasiliensis´ genome in biological samples by quantitative chain reaction polymerase (qPCR). Microb. Pathog. 121:359-362.

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Ribeiro, C. M., C. A. Caixeta, M. L. de Carli, F. F. Sperandio, de Sá Magalhães, Evandro Monteiro, A. A. C. Pereira, and J. A. C. Hanemann. 2017. Photodynamic inactivation of oral paracoccidioidomycosis affecting woman with systemic lupus erythematosus: an unusual case report. Photodiagnosis and Photodynamic Therapy. 17:160-163.

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Dos Santos, L. F., N. B. Melo, M. L. de Carli, A. C. S. Mendes, G. M. A. Bani, L. M. Verinaud, E. Burger, Gabriel de Oliveira, I. Moraes, A. A. Pereira, and M. R. Brigagão. 2017. Photodynamic inactivation of Paracoccidioides brasiliensis helps the outcome of oral paracoccidiodomycosis. Lasers in Medical Science. 32:921-930.

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Oliveira, A. F., and P. S. R. Coelho. 2017. Yeast expressing GP43 protein as a vaccine against Paracoccidioides brasiliensis infection, p. 213-224. Vaccines for Invasive Fungal Infections. Springer New York, NY.

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Public Health Agency of Canada. 2019. Human Pathogens and Toxins Act (HPTA) (S.C. 2009, c.24).

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