Encephalitozoon cuniculi in cats

 

Updated July 2021

 

These guidelines were published by D.D. Addie et al. in the Journal of Feline Medicine and Surgery 22 (2020), 1084-1088. The present guidelines were updated by D.D. Addie with input from Petra Benz (Dipl ECVO, Leonding, Austria).

 

Overview / Abstract

Encephalitozoon (E.) cuniculi is a common obligate intracellular microsporidian parasite of rabbits (Oryctolagus cuniculus), which is increasingly recognised as a pathogen of cats and other mammalian species.

 

Objectives: to review the literature on E. cuniculi in cats and provide recommendations for feline clinical conditions in which E. cuniculi should be considered as a possible differential diagnosis.

 

Conclusions:  E. cuniculi should be considered in cases of feline uveitis and cataract formation. It is not significantly associated with either chronic kidney disease or meningoencephalitis. E. cuniculi infection is more common in stray or feral cats than in pet cats.

Serological tests for antibody detection in blood are easy to perform and can be useful for diagnosis, but their specificity is low as antibodies have been found in apparently healthy cats. PCR appears more sensitive than histopathology for diagnosis, and is more sensitive when performed on cataractous lenses compared to aqueous humour, although ease of sampling is an obvious limitation. Treatment is with fenbendazole for 3 weeks and phacoemulsification to remove microsporidia from cataractous lenses.

 

E. cuniculi is a potential zoonotic infection, posing a special risk to immunocompromised humans from infected rabbits. Spore shedding has been infrequently identified in cats, so care should be taken around infected cats.

 

Introduction

Encephalitozoon cuniculi is a common obligate intracellular microsporidian parasite of rabbits (Oryctolagus cuniculus), which is increasingly recognised as a pathogen of cats and other mammals. These unicellular microsporidia were previously considered "primitive" protozoa; however, more recent insight gained through molecular phylogenetic analysis is indicating  that these organisms are not primitive but instead degenerate, and that microsporidia are related to the fungal Kingdom, either as a basal branch of the Fungi or as a sister group (Han and Weiss, 2017). Four species have been identified by PCR and sequencing: strain I is the rabbit strain; strain II is the mouse strain,  strain III is from the dog (Didier et al, 1995) and strain IV is from the cat (Benz et al, 2011 and Benz personal communication) .

 

In rabbits, E. cuniculi can infect all organs, but specifically causes chronic kidney and central nervous system disease (Nast et al., 1996; Harcourt-Brown and Holloway, 2003; Künzel et al., 2008; Valencakova et al., 2008; Csokai et al., 2009a, 2009b; Rodriguez-Tovar et al., 2017) as well as cataract formation (Ashton et al., 1976) with lens capsule rupture and phacoclastic uveitis (Stiles et al., 1997; Felchle and Sigler, 2002; Giordano et al., 2005; Künzel et al., 2008; Csokai et al., 2009b; Morsy et al., 2020). Infected rabbits shed spores in urine (Cox et al., 1980; Csokai et al., 2009b; Abu-Akkada and Oda, 2016; Rodriguez-Tovar et al., 2017) and faeces (Valencakova et al., 2008).

 

The susceptibility of cats to E.cuniculi infection was first reported in 1985, in an experimental infection of feline leukaemia virus-infected kittens (Pang and Shadduck, 1985).

 

Epidemiology

Kvac et al. (2017) detected E. cuniculi spores in the faeces of one pet and eight strays among 255 cats sampled in central Europe, and Piekarska et al. (2017) found spores in the faeces of one of 44 Polish cats.  No E. cuniculi spores were detected in the faeces of 40 and 26 cats in two studies in Iran, although Enterocytozoon bieneusi spores were found in the faeces of 3 /40 and 3/26 cats, respectively (Jamshidi et al., 2012; Askari et al., 2015).  No E. cuniculi spores were found in the faeces of ten Spanish cats tested (Lores et al., 2002).

 

Halánová et al. (2003) found antibodies to E. cuniculi in 17/72 cats in eastern Slovakia using an indirect immunofluorescence antibody test (IFAT).  In the same study, anti-E. cuniculi antibodies were found in 26/456 (5.7%) human sera samples examined. The highest occurrence of anti-microsporidial antibodies was found in a group of 24 immunocompromised patients: 37.5% (9/24) (Halánová et al., 2003).

 

Stray (Kvac et al., 2017) and feral (Tsukada et al., 2016) cats are more likely to be exposed or infected than pet cats.

 

A summary of prevalence data is shown in Table 1.

 

Table 1. Antibody / spores prevalence of E. cuniculi infection in the feline populations in various countries

Country/regionWhat was detectedNumber of catsPrevalence (%)Reference
AustriaAntibodies1002.0Benz et al., 2011
Central EuropeSpores2553.5Kvac et al., 2017
IranSpores
Spores
40
26
0
0
Jamshidi et al., 2012
Askari et al., 2015
JapanAntibodies2956.1Tsukada et al., 2016
PolandSpores442.3Piekarska et al., 2017
Slovakia (Eastern)Antibodies7223.6Halánová et al., 2003
SpainSpores100Lores et al., 2002
UKAntibodies270Meredith et al., 2015
USA (Virginia)Antibodies2326.5Hsu et al., 2011

 

Transmission

Mice appear to be the major reservoir of infection for cats (Benz et al, 2011). Cats are most likely to become infected by ingestion of mice rather than rabbits, since of 11 infected cats in Austria, seven were infected with the mouse strain (strain II), and the remaining four were infected with what is now known as the cat strain (strain IV) (Benz et al, 2011). 

 

Cats, like humans, may also be infected by consuming water or food contaminated with infective spores (Wang et al., 2018).  Oral and nasal transmission has been described in rabbits (Harcourt-Brown and Holloway, 2003), but it is unknown if direct transmission occurs in cats. Two uninfected cats that had been in direct contact with infected ones tested negative for blood antibodies in one study (Benz et al., 2011).  In utero infection is seen in rabbits (Baneux and Pognan, 2003), but it is unknown if transmission by this route occurs in the cat (Benz et al., 2011). Rebel-Bauder et al. (2011) reported a case of generalised encephalitozoonosis in a kitten with cerebellar hypoplasia, which could have been related to in utero infection. Benz et al (2011) speculate that vertical transmission is necessary for cataract formation in the cat because vertical transmission is presumed to play an important role in the mechanisms by which the microsporidium enters the lens in rabbits and mink: the presumption is that the organism would be unable to infect a lens which already had a fully formed capsule.

 

Clinical signs

In cats, ocular signs have been associated with E. cuniculi infection (Fig. 1).

 

Fig. 1: Ocular signs in a cat with confirmed E. cuniculi infection: keratic precipitates, which is evidence of anterior uveitis, in the right eye; cataract in the left eye. © Barbara Nell, University of Veterinary Medicine Vienna, Austria

 

Anterior uveitis and cataracts

Benz et al. (2011) reported a study of 19 eyes from 11 European Shorthair cats (median age 3.5 years) in Austria.  Nine of these cats had bilateral cataracts, with 12/19 eyes having focal anterior cortical cataracts and 7/19 eyes having mature cataracts. In 14/19 eyes, anterior uveitis was present. All cats had antibody titres in the blood (titre 1:80–1:10,000) for E. cuniculi (Benz et al., 2011).  E. cuniculi DNA was detected by PCR (Csokai et al., 2010) and sequencing in 18/19 lenses and in 10/19 aqueous humour samples (Benz et al., 2011).

 

Conditions not associated with E. cuniculi infection

Chronic kidney disease (CKD): 4/36 cats with CKD tested positive for E. cuniculi antibodies in blood but this prevalence was not significantly different (P>0.05) from cats without CKD (Hsu et al., 2011).

 

Meningoencephalitis (ME): Künzel et al. (2017) concluded that E. cuniculi was unlikely to be directly associated with (non-suppurative and/or granulomatous) ME in cats in Austria; none of 30 affected cats examined by immunohistochemistry were positive.

 

Diagnosis

Serology

Detection of antibodies in blood by Western blot or IFAT remains the major means of pre-mortem clinical diagnosis in animals. As the IFAT is quick and easy to perform, it is recommended for routine use in the diagnosis of feline encephalitozoonosis (Künzel et al., 2014). However, antibodies have been detected in cats that appeared to be clinically healthy, which has to be borne in mind when interpreting positive results: a positive result supports a diagnosis of encephalitozoonosis, but is not confirmatory.

 

PCR

Encephalitozoon cuniculi DNA was detected by PCR (Csokai et al., 2010) and sequencing in 18/19 lenses (liquefied lens material) and in 10/19 aqueous humour samples from 11 cats with cataracts (Benz et al., 2011).

 

Histopathology and cytology

Histopathology and cytology are aided by immunohistochemistry.  Five tentative positive results were achieved by cytological examination of material removed from cataractous lenses (Benz et al., 2011). Spores were detected in 15 of 19 samples of cataractous lens material with immunohistochemical staining (Benz et al., 2011).

 

E. cuniculi spores are difficult to observe when the samples are stained with haematoxylin and eosin, particularly when there is an inflammatory reaction and tissue damage. The spores are easily mistaken for other microorganisms, such as fungi (yeasts), protozoa and bacteria. Modified trichrome stain (MTS) and Gram stain, detected by light microscopy, and calcofluor white stain, detected by ultraviolet light microscopy, are the best stains for detecting spores of E. cuniculi in paraffin-embedded tissues. These stains were superior to Warthin–Starry, Ziehl–Neelsen, Giemsa and periodic acid–Schiff reaction for identifying spores without background ‘noise’ or monochromatic interference. In addition, these stains allow individual spores to be discerned in paraffin-embedded tissues. MTS allows observation of the polar tube, polaroplast and posterior vacuole, the most distinctive parts of the spore (Rodriguez-Tovar et al., 2017).

 

Leipig et al. (2013) recommended that confirmation of pathogenic E. cuniculi infection in rabbits should include standard histology of the predilection sites in combination with a specific aetiological assay, preferably real-time PCR. Presumably the same is true for diagnosis of E. cuniculi infection in cats.

 

Treatment

Fenbendazole is used to treat E. cuniculi infection in cats at a dose of 20 mg/kg q24 h for three weeks (Benz et al., 2011).  Cataracts can be successfully treated by phacoemulsification alongside medical treatment for E. cuniculi and symptomatic therapy for uveitis (e.g., ointment or drops containing dexamethasone), as reported by Benz et al. (2011).

 

Prevention

There is no commercially available vaccine to prevent E. cuniculi infection in rabbits or cats. It is noteworthy, however, that an experimental vaccine containing inactivated spores was shown to induce a long-lasting antibody response in rabbits (Sobottka et al., 2001). However, it is unknown whether antibodies are protective in this infection.

 

Where cats and rabbits are kept together, the main method of prevention of infection is by maintenance of excellent hygiene. Heat or steam cleaning will be the most effective means of eliminating E. cuniculi spores.  Rabbits suspected to be infected should be tested and treated.

 

The safest option for individuals - both cats and humans – that consume rabbit meat is for E. cuniculi-free sources to be used.  However, the prevalence of E. cuniculi is extremely high in rabbits kept for meat: 100% of 13 rabbit farms in Italy contained seropositive rabbits (Lonardi et al., 2013), and active E. cuniculi infections were determined in 85.9% and 56.3% of rabbits in commercial and household farms, respectively, in the Czech and Slovak Republics (Neumayerová et al., 2014). Where rabbit meat is prepared for feline or human consumption, it should be well cooked. Microsporidian spores in fish were shown to be inactivated by heating to 60°C for 10 min or by microwaving at 750 W, for 20 s (Leiro et al., 2012); similar treatment is likely to be effective for rabbit meat.  However, Graczyk et al. (2007) found microwaving to be ineffective against the spores of E. bieneusi and E. intestinalis in sewage sludge, and pasteurisation failed to inactivate spores in milk (Kváč et al., 2016) so more work is needed to determine appropriate conditions to inactivate E. cuniculi spores.  Microsporidian spores in fish were also inactivated by freezing at -20°C for more than 48 hours (Leiro et al., 2012).

 

Any area used to prepare rabbit meat should be thoroughly cleaned then disinfected with sodium hypochlorite (household bleach), ensuring a contact time of at least 16 min (Wolk et al., 2000), followed by rinsing with boiling water or steam cleaning. Similar precautions are recommended for pig meat, which Sak et al. (2019) recently reported can also contain E. cuniculi spores. Exposure to 70% ethanol for 15 min inactivated microsporidian spores in fish (Leiro et al., 2012), and so is likely to be effective for cleaning hands and utensils following the preparation of rabbit or pig meat (although use of disposable gloves would be more practical in the case of hands).

 

The ABCD recommends that immunosuppressed cats should not have any contact with infected rabbits or their urine and faeces.

 

Zoonotic risk

Since E. cuniculi is a zoonotic infection (Mathis et al., 2005), veterinary surgeons and nurses should wear gloves when dealing with infected cats or rabbits.  Calcium oxide (quicklime, burnt lime) was 100% effective in inactivating microsporidian spores in landfill leachate and sewage sludge (Graczyk et al., 2007), so could be used in the disposal of infected cadavers.

 

Acknowledgement

The ABCD thank Barbara Nell (Specialised Clinic for Ophthalmology at the Clinical Unit of Small Animal Surgery, University of Veterinary Medicine, Vienna) for providing the photo for Figure 1. The authors are deeply grateful to Dr Petra Benz for helpful comments which improved this guideline.

ABCD Europe gratefully acknowledges the support of Boehringer Ingelheim (the founding sponsor of the ABCD) and Virbac. 

 

References

Abu-Akkada SS, Oda SS (2016): Prevention and treatment of Encephalitozoon cuniculi infection in immunosuppressed rabbits with fenbendazole. Iran J Vet Res 17, 98-105.

 

Ashton N, Cook C, Clegg F (1976): Encephalitozoonosis (nosematosis) causing bilateral cataract in a rabbit. Br J Ophthalmol 60, 618-631.

 

Askari Z, Mirjalali H, Mohebali M, Zarei Z, Shojaei S, Rezaeian T, Rezaeian M (2015)Molecular Detection and Identification of Zoonotic Microsporidia Spore in Fecal Samples of Some Animals with Close-Contact to Human.  Iran J Parasitol 10(3), 381-388.

 

Baneux PJR, Pognan F (2003): In utero transmission of Encephalitozoon cuniculi strain type I in rabbits. Lab Anim 37, 132-138.

 

Benz P, Maass G, Csokai J, Fuchs-Baumgartinger A, Schwendenwein I, Tichy A, Nell B (2011): Detection of Encephalitozoon cuniculi in the feline cataractous lens.  Vet Ophthalmol 14 Suppl 1, 37-47.

 

Cox JC, Pye D, Edmonds JW, et al (1980): An investigation of Encephalitozoon cuniculi in the wild rabbit Oryctolagus cuniculus in Victoria, Australia. J Hyg (Lond) 84, 295-300. DOI: 10.1017/s0022172400026796.

 

Csokai J, Fuchs-Baumgartinger A, Maaß G et al (2010): Detection of Encephalitozoon cuniculi-infection (strain II) by PCR in a cat with anterior uveitis. Wien Tierärztl Monat – Vet Med Austria 97, 210–215.

 

Csokai J, Gruber A, Künzel F, et al (2009a): Encephalitozoonosis in pet rabbits (Oryctolagus cuniculus): pathohistological findings in animals with latent infection versus clinical manifestation. Parasitol Res 104, 629-635.

 

Csokai J, Joachim A, Gruber A, et al (2009b): Diagnostic markers for encephalitozoonosis in pet rabbits. Vet Parasitol 163, 18-26.

 

Didier ES, Vossbrinck CR, Baker MD et al (1995): Identification and characterization of three Encephalitozoon cuniculi strains.  Parasitology 11, 411–421.

 

Felchle L, Sigler RL (2002): Phacoemulsification for the management of Encephalitozoon cuniculi-induced phacoclastic uveitis in a rabbit. Vet Ophthalmol 5, 211-215.

 

Giordano C, Weigt A, Vercelli A, et al (2005): Immunohistochemical identification of Encephalitozoon cuniculi in phacoclastic uveitis in four rabbits. Vet Ophthalmol 8, 271-275.

 

Graczyk TKKacprzak MNeczaj ETamang LGraczyk HLucy FEGirouard AS (2007): Human-virulent microsporidian spores in solid waste landfill leachate and sewage sludge, and effects of sanitization treatments on their inactivation. Parasitol Res 101(3), 569-575.

 

Halánová MCisláková LValencákova ABálent PAdam JTrávnicek M (2003): Serological screening of occurrence of antibodies to Encephalitozoon cuniculi in humans and animals in Eastern Slovakia. Ann Agric Environ Med 10(1), 117-120.

 

Han B, Weiss LM (2017): Microsporidia: obligate intracellular pathogens within the fungalkingdom. Microbiol Spectr 5(2); doi: 10.1128/microbiolspec.FUNK-0018-2016.

 

Harcourt-Brown FM, Holloway HKR (2003): Encephalitozoon cuniculi in pet rabbits. Vet Rec 152, 427–431.

 

Hsu V, Grant DC, Zajac AM, Witonsky SG, Lindsay DS (2011): Prevalence of IgG antibodies to Encephalitozoon cuniculi and Toxoplasma gondii in cats with and without chronic kidney disease from Virginia. Vet Parasitol 176(1), 23-26.

 

Jamshidi ShTabrizi ASBahrami MMomtaz H (2012): Microsporidia in household dogs and cats in Iran; a zoonotic concern. Vet Parasitol 185(2-4), 121-123.

 

Künzel F, Gruber A, Tichy A, et al (2008): Clinical symptoms and diagnosis of encephalitozoonosis in pet rabbits. Vet Parasitol 151, 115-124.

 

Künzel FPeschke RTichy AJoachim A (2014): Comparison of an indirect fluorescent antibody test with Western blot for the detection of serum antibodies against Encephalitozoon cuniculi in cats. Parasitol Res 113(12), 4457-4462.

 

Künzel F, Rebel-Bauder B, Kassl C, Leschnik M, Url A (2017): Meningoencephalitis in cats in Austria: a study of infectious causes, including Encephalitozoon cuniculi. J Feline Med Surg 19(2), 171-176.

 

Kvac M, Hofmannova L, Ortega Y, Holubova N, Horcickova M, Kicia M, Hlaskova L, Kvetonova D, Sak B, McEvoy J (2017):  Stray cats are more frequently infected with zoonotic protists than pet cats. Folia Parasitol (Praha) 6, 64.

 

Kváč M, Tomanová V, Samková E, et al (2016): Encephalitozoon cuniculi in raw cow’s milk remains infectious after pasteurization. Foodborne Pathog Dis 13, 77-79.

 

Leipig M,  Matiasek K, Rinder H, Janik D, Emrich D, Baiker K, Hermanns W (2013): Value of histopathology, immunohistochemistry, and real-time polymerase chain reaction in the confirmatory diagnosis of Encephalitozoon cuniculi infection in rabbits. J Vet Diagn Invest 25(1), 16–26.

 

Leiro JMPiazzon CDomínguez BMallo NLamas J (2012): Evaluation of some physical and chemical treatments for inactivating microsporidian spores isolated from fish. Int J Food Microbiol 156(2), 152-160.

 

Lonardi C, Grilli G, Ferrazzi V, et al (2013): Serological survey of Encephalitozoon cuniculi infection in commercially reared rabbit does in Northern Italy. Res Vet Sci 94, 295-298.

 

Lores Bdel Aguila CArias C (2002): Enterocytozoon bieneusi (microsporidia) in faecal samples from domestic animals from Galicia, Spain. Mem Inst Oswaldo Cruz 97(7), 941-945.

 

Mathis A, Weber R, Deplazes P (2005): Zoonotic potential of the microsporidia. Clin Microbiol Rev 18, 423-445.

 

Meredith ALCleaveland SCBrown JMahajan AShaw DJ (2015): Seroprevalence of Encephalitozoon cuniculi in wild rodents, foxes and domestic cats in three sites in the United Kingdom. Transbound Emerg Dis 62(2), 148-156.

 

Morsy EA, Salem HM, Khattab MS et al (2020): Encephalitozoon cuniculi infection in farmed rabbits in Egypt. Acta Vet Scand 62, 11. DOI: 10.1186/s13028-020-0509-6.

 

Nast R, Middleton DM, Wheler CL (1996): Generalized encephalitozoonosis in a Jersey wooly rabbit. Can Vet J 37, 303-305.

 

Neumayerová H, Juránková J, Jeklová E, et al (2014): Seroprevalence of Toxoplasma gondii and Encephalitozoon cuniculi in rabbits from different farming systems. Vet Parasitol 204, 184-190.

 

Pang VFShadduck JA (1985): Susceptibility of cats, sheep, and swine to a rabbit isolate of Encephalitozoon cuniculiAm J Vet Res 46(5), 1071-1077.

 

Piekarska JKicia MWesołowska MKopacz ŻGorczykowski MSzczepankiewicz BKváč MSak B (2017): Zoonotic microsporidia in dogs and cats in Poland. Vet Parasitol 246, 108-111.

 

Rebel-Bauder BLeschnik MMaderner AUrl A (2011): Generalized encephalitozoonosis in a young kitten with cerebellar hypoplasia. J Comp Pathol 145(2-3), 126-131.

 

Rodriguez-Tovar LE, Villarreal-Marroquin A, Nevarez-Garza AM, et al (2017): Histochemical study of Encephalitozoon cuniculi spores in the kidneys of naturally infected New Zealand rabbits. J Vet Diagn Invest 29, 269-277.

 

Sak B, Vecková T, Brdíčková K, et al (2019): Experimental Encephalitozoon cuniculi infection acquired from fermented meat products. Foodborne Pathog Dis 16, 394-398.

 

Sobottka IIglauer FSchüler TSchmetz CVisvesvara GSAlbrecht HSchwartz DAPieniazek NJBartscht KLaufs RSchottelius J (2001): Acute and long-term humoral immunity following active immunization of rabbits with inactivated spores of various Encephalitozoon species. Parasitol Res 87(1), 1-6.

 

Stiles J, Didier E, Ritchie B, et al (1997): Encephalitozoon cuniculi in the lens of a rabbit with phacoclastic uveitis: conformation and treatment. Progress in Veterinary & Comparative Ophthalmology 7, 233-238.

 

Tsukada R, Osaka Y, Takano T, Sasaki M, Inose M, Ikadai H (2016): Serological survey of Encephalitozoon cuniculi infection in cats in Japan.  J Vet Med Sci 78(10), 1615-1617.

 

Valencakova A, Balent P, Petrovova E, et al (2008): Encephalitozoonosis in household pet Nederland Dwarf rabbits (Oryctolagus cuniculus). Vet Parasitol 153, 265-269.

 

Wang S, Yao Z, Li L, Pan Y, Li P, Nan X, Xie Q, Zhang Z (2018): Seroprevalence of Toxoplasma gondii and Encephalitozoon cuniculi among domestic rabbits in central China.  Parasite 25, 9.

 

Wolk DM, Johnson CH, Rice EW, Marshall MM, Grahn KF, Plummer CB, Sterling CR (2000): A spore counting method and cell culture model for chlorine disinfection studies of Encephalitozoon syn. Septata intestinalis. Appl Environ Microbiol 66(4), 1266-1273.

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