Cryptosporidium sp. is a coccidian parasite that belongs to Cryptosporidiae family. Initially, only one species was recognised (C. parvum), which was then divided in to two genotypes: genotype 1 for humans and genotype 2 for animals. After reviewing the literature it shoved that species status was based on four components or characteristics: oocyst morphology, host specicifity, genetic characterisation and international nomenclature concordation (International Committee for Zoological Nomenclature).

Currently, more than 30 species of Cryptosporidium have been recognised, including C. muris, C. andersoni, C. parvum, C. hominis, C. wrairi, C. felis, and C. canis in mammals; C. baïleyi, C. meleagridis, and C. galli in birds; and C. molnari in fish. Because of the low host specificity the number of species remains a matter of discussion. 

Cryptosporidium has been found in around 60 species of reptile including 40 species of snakes,, 15 lizards and 2 tortoises at the time of writing. These include C. serpentis found in snakes, C. varanii (previously C. saurophilum) found in lizards but also snakes, and C ducimarci and C. tortoise genotype in tortoises including Pancake, Russian and Hermann's tortoises (Ritcher et al., 2012). The author has identified unknown Cryptosporidium sp. in a Russina Tortoise with clinical signs that was ultimately euthanased (not published).

Cryptosporidium saurophilum was named by Koudela and Modry in 1998 from a Schneiders skink. C varanii (saurophilum) was elevated as a proper species of intestinal pathogen of lizards (Fayer, 2010; Pavlasek and Ryan, 2008). A paper was presented by Pavlasek and Ryan showing biological as well as genetic analysis of Cryptosporidium varanii at the 18S rRNA and actin loci and show that it is genetically identical to C. saurophilum. As C. varanii was described prior to C. saurophilum, it takes precedence over C. saurophilum and therefore C. saurophilum should be considered a junior synonym of C. Varanii. 

Although there are reports of Cryptosporidium in reptiles since the 70's, C. Serpentis has been genetically described relatively recently (Xiao et al., 1999; Kimbell et al., 1999). 

Cryptosporidium has been identified in tortoises over the last 20 years, including species from testudo, Indotestudo, Geochelone and Gopherus genres (Xiao et al., 2004; Graczyk et al., 1998; Alves et al., 2005; Griffin et al., 2010; Bonnie et al., 1997). Over the last few years there has been the suggestion of a new Cryptosporidium species affecting tortoises (Traversa et al., 2008; Pedraza et al., 2009; Griffin et al., 2010; Richter and Rasim, 2012). Traversa in 2010 proposed  that the  species of Cryptosporidium be named as C. ducismarci but due to the lack of a traditional formal description associated with naming a new parasite including oocyst description C. ducismarci cannot be considered a valid species and should be referred to as tortoise genotype II. Tortoise genotype I has been identified in gastric mucosa in a Russian tortoise and tortoise genotype II has been identified in intestinal lesions in a Pancake tortoise and a Russian tortoise (Griffin et al., 2010). Phylogenetically, tortoise genotype I is closer to C. serpentis and tortoise genotype II is closer to C. varanii (Griffin et al., 2010). Tortoise genotype I has been detected in a Ball Python and tortoise genotype II has been identified in a chameleon (Pedraza Diaz et al., 2009).
Update: Cryptosporidium genotype I has been established as a new species known as Cryptosporidium Testudinis sp. (Article)
Genetic characterisation proved that Cryptosporidium isolates from a group of 5 tortoises in the Testudo genre were identical to the zoonotic Cryptosporidium pestis (C. parvum, bovine genotype), which indicates a potential zoonotic risk (Traversa et al., 2008).
Those concerning us are C. serpentis and C. varanii. These species are found mainly in the gastrointestinal sytem; C. serpentis in the stomach and C. varanii in the small intestine (Lihua Xiao et al. 2004), although they can also be founs in other locations including ear canal (Fitzgerald et al. 1998) and pharynx (Uhl et al. 2001).
C. Saurophilum has been detected in Corn snakes (Plutzer and Karanis, 2007).
Cryptosporidium serpentis has also been detected in cattle (Azami et al., 2007; Chen and Qiu, 2012).

C. hominis and C. parvum are frequently detected in humans. C. parvum does not require contact with farm animals and transmission between humans has been proven. C. felis, C. canis and C. meleagridis have been found in humans with adquired immunodeficiency syndrome (AIDS).


All phases in the cycle are intracellular and on excretion the infective sporulated oocyst contains 4 sporozoites. After ingestion and excystation in the host, each sporozoite adheres to the lumenal surface of a epithelial cell. Microvilli surround the sporozoite gradually becoming intracellular and extracytoplasmatic. Multiple fission (merogonyschizogony) ocurrs, forming 8 merozoites within the meront. Merozoites are released and further Type I meronts are formed. Type II meronts are also formed containing 4 merozoites and they undergo the sexual phase forming macrogametes and microgametes, and ultimately the zygote. Meiosis ocurrs and 4 sporozoites are formed (sporogony). Approximately 20% of the oocysts fail to form a wall and are often termed "thin walled oocysts". It is believed that these infect new cells in the gut. The remaining oocysts are then passed in to the enviroment via the faeces measuring between 4-6um.


Infections in humans are normally associated to trips abroad, contact with farm animals, contact in pools or care centres. Over 400,000 people were infected in Milwaukee after Cryptosporidium found a way in to the water supply. In reptiles, infection is a major concern in large collections and especially in leopard geckos and specific snake species. In reptiles transmission is faecal-oral, either via direct contact or through fomites or vehicles, including us whilst cleaning out encloosures. With routine disinfections not killing oocysts, a disease not treatable with "routine" drugs and high mortality rate, this disease can be considered somewhat complicated.

Detection of oocysts may prove challenging for the following reasons: 1) excretion is intermittent and therefore a single test may show a false negative, and 2) the presence of asymptomatic carriers.


In mammals infection often is autolimiting and is considered important due to financial losses in farm animals and serious illness in immunodepressed humans. Also, in other animals Cryptosporidium infections are selflimiting in immunocompetent individuals, whereas in reptiles it is often chronic and sometimes fatal in snakes.

The most common symptoms in reptiles include: apathy, missing feeds, regurgitation, diarrhoea, maldigestion-malabsorption syndrome, anorexia, cachexia, and death. Leopard geckos produce foul smelling faeces and snakes may present with a mid-abdomen dilation. Asymptomatic animals are also common.


Gold standard for ante-mortem and post-mortem diagnosis of cryptosporidiosis is histopathology, rendering gastric hyperplasia in snakes and proliferative enteritis in lizards with the presence of cryptosporidium on mucosal surface.

Presumptive diagnosis may be based on history, symptoms and species. Faecal analysis or gastric lavage (in snakes) and specialised staining techniques are common tests to perform in live animals. More specialised tests are readily available now and include polymerase chain reaction (PCR) and immunological tests ie ELISA. A negative diagnosis from a Modified Kinyoun-stained preparation does not preclude the presence of Cryptosporidium. At least three consecutive specimens may be required (

In live reptiles samples are sent to the laboratory either fresh or stored in 5% or 10% formaline, or sodium acetate-acetic acid–formalin (SAF).

Diagnostic techniques include:

a) Direct wet preparation: Due to their small size and similarity to yeast this may prove difficult.

b) Specific Stains: Modified Ziehl-Neelsen (mZN) (Kin Youn) and auramine/rhodamine stain. mZN will stain oocysts red with a blue (methylen blue) or green (malachite green) counter-stain and is considered the standard test according to some studies (Macpherson et al., 1993). Auramine stain is to be used with fluorescence microscopy in complete blackout. Control samples are recommended.

Modified Ziehl-Neelson faecal stain from a Leopard Gecko 400X

Modified Ziehl-Neelson faecal stain from a Boa Constrictor 1000X

Modified Ziehl-Neelson faecal stain from a Green Iguana 1000X

Oocysts on direct wet mount of a Bearded Dragon 400X

Modified Ziehl-Neelson faecal stain from a Bearded Dragon 1000X


c) Immunochromatographic tests: monoclonal antibodies re directed at Cryptosporidium parvum antigen. Although has been used in reptiles, still has to be validated with adequate trials for detection of C. varanii/serpentis. We are currently running tests with these and results should be available in 2016.

d) Enzyme linked immunoassay (ELISA): sensitivity varies between 93-97% and false positives increase with the presence of blood. These have not been validated in cryptosporidium species affecting reptiles.

e) Polymerase chain reaction (PCR): becoming readily available at laboratories.

f) Immunofluorescence (IF): an antibody is used that is chemicaly linked to a fluorophore (direct) or a specifica antibody detects the antigen and then a second antibody that carries the fluorophore recognises the primary antibody (indirect).

A study performed in 2009 by Pedraza-Diaz, Cryptosporidium was detected in 38.6% and 25.1% of the samples analysed using IF and PCR, respectively. If we have a positive faecal result and a negative IF then the reptile maybe shedding non-reptilian oocysts or it is too soon to have formed antibodies. If we have a positive IF result and a negative faecal (or faecals) then the reptile is not shedding oocysts (or numbers are very reduced).

In mammals Cryptosporidium antibodies around 10 days post-infection, whereas in snakes, this takes around 10 weeks. Anti-cryptosporidium immunoglobulins are passed through the egg and are detected in juveniles up to 2 months of age (not yet published at time of writing).

Many laboratories use modified ZN and auramine staining techniques for primary diagnosis and a modified ZN test is considered very effective and according to some studies is the elected diagnostic test (Macpherson et al., 1993).

Detection of Cryptosporidium during quarantine periods in Leopard Gecko collections may prove challenging due to intermittent shedding and therefore examination of up to 5-7 samples has been suggested (Graczyk and Cranfield, 1996).


Many different drugs have been used to treat cryptosporidiosis with not much success.

Ionophores (alborixin, halofuginone, lasalocid, maduramicin) have historically been used in animals and, although effective, cause severe hepato- and nephrotoxicity in snakes (Graczyk et al., 1996). Aprinocid and azithromycin have been used (Current, 1999, Coke et al., 2003). Metronidazole, spiramycin combined with Paromomycin,  Trimetoprim/sulphonamide combinations have all been used with varied and disappointing results (Bone, 1992; Cranfield and Graczyk, 1996; Frye, 1991; Grosset et al., 2011; Pantchev et al., 2008).

Paromomycin has also been widely used and seemed to render good results at high doses (100mg-360mg/kg) (Cranfield and Graczyk, 2006; Grosset et al., 2011; Jacobson, 1988; Pare et al., 1997; Wilson and Carpenter, 1996; ). A 800mg/kg dose has also been successful in Leopard Geckos (Coke et al., 1998). The author has seen good control of clinical signs with precise stomach feeding plans and paromomycin every 48h at 300mg/kg in Leopard Gecko collections. Renal failure has been described in cats (Gookin et al., 1999) so a potential risk should be considered.

Treatment with hyperimmune bovine calostrum (HBC) has shown promising results in snakes, leopard geckos and Savannah monitors (Cranfield et al., 1999; Graczyk et al., 1998b; 1999; 2000). Snakes administered HBC at 1% bodyweight once weekly showed clearance of the parasite on histology and reduced oocysts from stomach washes and faecal  samples in clinically affected individuals.  The use of the commercially available bovine serum product Colostrix is therefore a possible treatment option.

General hygiene precautions should always be followed. Prevention of transmission between different groups of reptiles in mixed collections is essential. Infections in chelonians and lizards may be subclinical, allowing potential fatal transmission to snakes.

In collections quarantine of incoming reptiles and sequenced testing with a removal/euthanasia policy may be advisable.

Disinfection of enclosures is essential whether it is a hospital environment or vivaria of a reptile collection. Only ammonia (5%) and formol saline (10%) have been proven effective at low temperatures (Cranfield and Graczyk, 1995). Efficient ways to disrupt the parasite cycle are steam cleaning, freezing and desiccation.


Alves M, Xiao L, Lemos V, Zhou L, Cama V, da Cunha MB, Matos O, Antunes F. 2005. Occurrence and molecular characterization of Cryptosporidium spp. in mammals and reptiles at the Lisbon Zoo. Parasitol Res. 97:108–112. 

Antinoff N. 2000. Cryptosporidium in a Green Iguana. Proceedings of the Association of Reptilian and Amphibian Veterinarians 2000, pp.15-18

Barnard S M and Upton S J. 1994. A Veterinary Guide to the Parasites of Reptiles. Volume 1. Protozoa. Krieger Publishing C o, Malabar,FL.

Biron K. 2008. Cryptosporidiosis in reptiles: diagnosis and therapy. Proceedings ARAV, 18-19.

Bonnie RL, Calle PP, Gottdenker N, James S, Linn WJ, McNamara T, Cook RA. 1997. Proceedings of the Annual meeting of the American Association of Zoo Veterinarians: 26-30 October 1997; Houston, Texas. AAZV; 1997. Clinical significance of Cryptosporidia in captive and free-ranging chelonians; pp. 19–20.

Brownstein DG. 1997. Cryptosporidium in snakes with hypertrophic gastritis. Journal of Veterinary Pathology 14, 606-617

Coke RL and T ristan T E. 1998. Cryptosporidium infection in a colony of leopard geckos, Eublepharis macularius, in Proceedings of the Association of Reptilian and Amphibian Veterinarians 5th Annual Meeting, Kansas City, MO, 157–165.

Cranfield MR, Graczyk TK. 1996. Cryptosporidiosis. In DR Mader, Reptile Medicine and Surgery, WB Saunders, Philadelphia, p. 359-363.

"Cryptosporidiosis Laboratory Case Definition (LCD)". Accesed 26th March 2014.

"Cryptosporidium parvum." In Foodborne Pathogenic Microorganisms and Natural Toxins Handbook. U.S. Food & Drug Administration, Center for Food Safety & Applied Nutrition, Feb 2002. 10 Oct 2002.

da Silva D.C., Paiva P.R., Nakamura A.A., Homem C.G., de Souza M.S., Grego K.F., Meireles M.V. The detection of Cryptosporidium serpentis in snake fecal samples by real time PCR. Vet. Parasitol. 2014;204:134–138. [PubMed]

Donoghue PJ. 1995. Cryptosporidium and cryptosporidiosis in man and animals. Internat J Parasitol 25:139–195.

Fayer R. 2010. Taxonomy and species delimitation in Cryptosporidium. Exp Parasitol. 124:90–97. 

Fitzgerald S D, Molsan PG, and B ennett R. 1998. A ural polyp associated with cryptosporidiosis in an iguana (Iguana iguana). JVet Diagn Invest 10:179–180.

Frye FL, Garman RH, Graczyk T K, B oyer T H, and Miller H. 1999. Atypical non-alimentary cryptosporidiosis in three lizards, in Proceedings of the Association of Reptilian and Amphibian Veterinarians 6th Annual Conference, Columbus, OH, pp.43–48.

Graczyk TK, Cranfield MR, Fayer R 1995. A comparative assessment of direct fluorescence antibody modified acid fast stain, and sucrose flotation techniques for detection of Cryptosporidium serpentis oocysts in snakes fecal specimens. J Zoo Wildl Med 26: 396-402.

Graczyk TK, Cranfield MR 1997. Detection of Cryptos-poridium-specific serum immunoglobulins in captive snakes by a polyclonal antibody in the direct Elisa. Vet Res 28: 131-142.

Graczyk T K, B alazs GH, Work T , A guirre AA , Ellis DM, Murakawa SKK, and Morris R. 1997. Cryptosporidium sp. infections in green turtles, Chelonia mydas, as a potential source of marine waterborne oocysts in the Hawaiian I slands. Appl Environ Micro 63:2925–2927.

Graczyk T K, C ranfield MR, Mann J, and S trandberg JD. 1998. Intestinal Cryptosporidium sp. infection in the Egyptian tortoise, Testudo kleinmanniInt J Parasitol 28:1885–1888.

Graczyk T K and Cranfield MR. 1998. Experimental transmission of Cryptosporidium oocyst isolates from mammals, birds and reptiles to captive snakes. Vet Res 29:187–195.

Graczyk T K, Cranfield MR, and B ostwick EF. 1999. Hyperimmune bovine colostrums treatment of moribund leopard geckos (Eublepharis macularius) infected with Cryptosporidium sp. Vet Res 30:377–382.

Graczyk T K and C ranfield MR. 2000. Cryptosporidium serpentis oocysts and microsporidian spores in feces of captive snakes. J Parasitol 86:413–414.

Griffin C, Reavill DR, Stacy BA, Childress AL, Wellehan JFX. 2010. Cryptosporidiosis caused by two distinct species in Russian tortoises and a Pancake tortoise. Vet Par. 

Grosset C, Villeneuve A, Brieger A, and Lair S. 2011. Cryptosporidiosis in Juvenile Bearded Dragons (Pogona vitticeps): Effects of Treatment with Paromomycin. Journal of Herpetological Medicine and Surgery. 2011;21(1):10-15

Henriksen A, Pohlenz JFL 1981. Staining of Cryptosporidium by a modified Ziehl-Neelsen technique. Acta Vet Scand 22: 594-596.

Heuschele WP, O osterhuis J, Janssen D, Robinson PT, Ensley PK, Meier E, O lson T , A nderson MP, and B enirschke K. 1986. Cryptosporidial infections in captive wild animals. J Wildl Dis 22:493–496.

Juranek, D. "Cryptosporidiosis: Sources of Infection and Guidelines for Prevention." Centers for Disease Control and Prevention (CDC), 2000. 10 Oct 2002.

Kimbell LM, Miller DL, Chavez W, Altman N. 1999. Molecular analysis of the 18S rRNA gene of Cryptosporidium serpentis in a wild-caught corn snake (Elaphe guttata guttata) and a five-species restriction fragment length polymorphism-based assay that can additionally discern C. parvum from C. wrairi. Appl Environ Microbiol. 65:5345–5349.

Klingenberg RJ (1996) Enteric cryptosporidiosis in a colony of Indigo Snakes, Drymarchon corais, a Panther Chameleon, Chameleon pardalis, and a Savannah monitor, Varanus exhanthematicus. Bulletin of the Association of Amphibian and reptilian veterinarians, 6 (1), 5-9

Koudela B and Modry D. 1998. N ew species of Cryptosporidium (Apicomplexa: Cryptosporidiidae). Folia Parasitol 45:93–100.

Morgan UM, Pallant L, Dwyer BW, Forbes DA, Rich G, Thompson RCA. 1998. Comparison of PCR and Microscopy for Detection of Cryptosporidium parvum in Human Faecal Specimens: Clinical Trial J. Clin. Microbiol. 36:995-998.

"Material Safety Data Sheet - Cryptosporidium parvum." January 2001 Canadian Laboratory Centre for Disease Control, February 2000. 10 October 2002.

O’Donoghue PJ. 1995. Cryptosporidium and cryptosporidiosis in man and animals. Int J Parasitol 25:139–195.

Pavlasek I, Ryan U. Cryptosporidium varanii takes precedence over C. saurophilum. Exp Parasitol. 2008;118:434–437.

Pedraza-Díaz S, Ortega-Mora LM, Carrión BA, Navarro V, Gómez-Bautista M. Molecular characterisation of Cryptosporidium isolates from pet reptiles. Vet Parasitol. 2009;160:204–210.

Plutzer, J, Karanis, P (2007) Molecular identification of a Cryptosporidium saurophilum from corn snake (Elaphe guttata guttata). Parasitol Res 101: pp. 1141-1145

Richter, B, Nedorost, N (2011) Detection of Cryptosporidium species in feces or gastric contents from snakes and lizards as determined by polymerase chain reaction analysis and partial sequencing of the 18S ribosomal RNA gene. J Vet Diagn Invest 23: pp. 430-435

Richter, B, Rasim, R (2012) Cryptosporidiosis outbreak in captive chelonians (Testudo hermanni) with identification of two Cryptosporidium genotypes. J Vet Diagn Invest 24: pp. 591-595

Taylor MA, Geach MR, and C ooley WA. 1999, C loacal and pathological observations on natural infections of cryptosporidiosis and flagellate protozoa in leopard geckos (Eublepharis macularius). Vet Rec 145:695–699.

Terrell S P, Uhl EW, and Funk RS. 2003. Proliferative enteritis in leopard geckos (Eublepharis macularius) associated with Cryptosporidium sp. infection. J Zoo Wildl Med 34:69–75.

Tilley, M, Upton, SJ (1990) A comparative study of the biology of Cryptosporidium serpentis and Cryptosporidium parvum (Apicomplexa:Cryptosporidiidae). J Zoo Wildlife Med 21: pp. 463-467

Traversa D, Iorio R, Otranto D, Modrý D, Slapeta J. Cryptosporidium from tortoises: Genetic characterisation, phylogeny and zoonotic implications. Mol Cell Probes. 2008;22:122–128.

Traversa, D (2010) Evidence for a new species of Cryptosporidium infecting tortoises: Cryptosporidium ducismarci. Parasit Vectors 3: pp. 21-23

Uhl EW, Jacobson E, B artick T E, Micinilio J, and S chmidt R. 2001. Pharyngeal polyps associated with Cryptosporidium infection in three iguanas (Iguana iguana). Vet Pathol 38:239–242.

Upton S J. 1990. Cryptosporidium spp. in lower vertebrates, in Cryptosporidiosis of Man and Animals, Dubey JP, S peer CA , and Fayer R (Eds.), CRC Press, Boca Raton, FL, 149–156.

Wright, K. (1997). Cryptosporidium controversy: when do you consider a reptile crypto-free? Association of Reptilian and Amphibian Veterinarians Conference Proceedings 1997:169-173.

Xiao L, Escalante L, Yang C, Sulaiman I, Escalante AA, Montali RJ, Fayer R, Lal AA. 1999. Phylogenetic analysis of Cryptosporidium parasites based on the small-subunit rRNA gene locus. Appl Environ Microbiol. 65:1578–1583.

Xiao L , Ryan UM, Graczyk T K, L imor J, L i L , Kombert M, Junge R, Sulaiman I M, Zhou L , A rrowood MJ, Koudela B , Modry D, and L al AA . 2004. Genetic diversity of Cryptosporidium spp. in captive reptiles. Appl Environ Microbiol 70:891–899.


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