The opportunistic pathogen Encephalitozoon cuniculi in wild living Murinae and Arvicolinae in Central Europe

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ScienceDirect European Journal of Protistology 69 (2019) 14–19

SHORT COMMUNICATION

The opportunistic pathogen Encephalitozoon cuniculi in wild living Murinae and Arvicolinae in Central Europe Agnieszka Perec-Matysiaka,∗ , Kinga Le´snia´nskaa , Katarzyna Bu´nkowska-Gawlika , b,c , Bohumil Sakc , Martin Kvᡠˇ ˇ Sárka Condlová cb,c , Duˇsan Rajsk´yd , Joanna Hildebranda a

Department of Parasitology, Institute of Genetics and Microbiology, University of Wrocław, Przybyszewskiego 63, 51-148 Wrocław, Poland ˇ ˇ Faculty of Agriculture, University of South Bohemia in Ceské Budˇejovice, Studentská 13, 370 05 Ceské Budˇejovice, Czech Republic c ˇ Institute of Parasitology, the Biology Centre of the Czech Academy of Sciences, Braniˇsovská 31, 370 05 Ceské Budˇejovice, Czech Republic d Faculty of Forestry, Department of Forest Protection and Wildlife Management, Technical University in Zvolen, Zvolen, Slovak Republic b

Received 9 November 2018; received in revised form 16 January 2019; accepted 8 February 2019 Available online 14 February 2019

Abstract Encephalitozoon spp. is an obligate intracellular microsporidian parasite that infects a wide range of mammalian hosts, including humans. This study was conducted to determine the prevalence of Encephalitozoon spp. in wild living rodents from Poland, the Czech Republic and Slovakia. Faecal and spleen samples were collected from individuals of Apodemus agrarius, Apodemus flavicollis, Apodemus sylvaticus, and Myodes glareolus (n = 465) and used for DNA extraction. PCR, targeting the ITS region of the rRNA gene was performed. The overall prevalence of microsporidia was 15.1%. The occurrence of Encephalitozoon cuniculi in the abovementioned host species of rodents has been presented for the first time, with the highest infection rate recorded for A. flavicollis. Sequence analysis showed that the most frequent species was E. cuniculi genotype II (92.5%). E. cuniculi genotypes I (1.5%) and III (6.0%) were also identified. © 2019 Elsevier GmbH. All rights reserved. Keywords: Encephalitozoon cuniculi; Microsporidia; Molecular analysis; Small rodents

Introduction Encephalitozoon spp., a member of the phylum Microsporidia, is an obligate intracellular parasite. The three species in this genus, which infect both human and animal hosts, are Encephalitozoon cuniculi, Encephalitozoon intestinalis and Encephalitozoon hellem (Didier et al. 2004). Encephalitozoon infections in animals usually disseminate and can affect almost every organ. Encephalitozoon cuniculi ∗ Corresponding

author. E-mail address: [email protected] (A. Perec-Matysiak). https://doi.org/10.1016/j.ejop.2019.02.004 0932-4739/© 2019 Elsevier GmbH. All rights reserved.

has been detected in a wide range of mammals including rodents, lagomorphs, carnivores, ruminants, nonhuman primates, and humans (Didier and Weiss 2011; Hinney et al. 2016). Molecular techniques confirmed the presence of four different E. cuniculi genotypes (I–IV): E. cuniculi genotype I primarily isolated from rabbits; E. cuniculi genotype II initially obtained from mice and blue foxes (Åkerstedt et al. 2002; Mathis et al. 1996); E. cuniculi genotype III obtained from dogs, humans and nonhuman primates (Didier et al. 1995, 1996; Juan-Salles et al. 2006); and E. cuniculi genotype IV identified in a human and a dog (Talabani et al. 2010; unpublished GenBank Acc. No. JQ340013). Although all genotypes are considered zoonotic, humans are

A. Perec-Matysiak, K. Le´snia´nska, K. Bu´nkowska-Gawlik, et al. / European Journal of Protistology 69 (2019) 14–19

mostly infected with E. cuniculi genotype II (Didier 2005; Didier and Weiss 2011; Mathis et al. 2005; Sokolova et al. 2011). Encephalitozoon intestinalis has been reported in a wider range of mammals but has so far been detected only sporadically in wild animals (Bornay-Llinares et al. 1998; Graczyk et al. 2002; Hinney et al. 2016; Murphy et al. 2007). Encephalitozoon hellem is mainly distributed amongst birds and some mammals (Childs-Sanford et al. 2006; Hinney et al. 2016). Although the presence of Encephalitozoon spp. has been generally studied among humans and domestic animals, there is still information needed on the role of wild living animals, including rodents, which may be a potential source of environmental contamination with this zoonotic microsporidia species. Microsporidial spores can be released into the environment via faeces, urine or respiratory secretions of infected individuals (Si´nski 2003). Infection with microsporidia has been confirmed in rodents with the use of the serological and the molecular methods (Hersteinsson et al. 1993; Meredith et al. 2015; Müller-Doblies et al. 2002; Sak et al. 2011). Molecular epidemiology surveys on the contribution of small rodents to spore spreading in the environment have so far been limited and concern rodents obtained from Austria, the Czech Republic–Germany border, Slovakia, and Japan (Daniˇsová et al. 2015; Fuehrer et al. 2010; Sak et al. 2011; Tsukada et al. 2013). However, representatives of wild living Murinae and Arvicolinae, i.e. Apodemus agrarius, Apodemus flavicollis, Apodemus sylvaticus, and Myodes glareolus, which are widely distributed and abundant in Central Europe, have not yet been examined as potential reservoirs of Encephalitozoon spp. The purpose of this survey was therefore the molecular determination of the pathogen in rodents as reservoir hosts obtained from selected Central European countries.

Material and Methods Area and specimens studied From 2011 to 2014 (from summer to autumn each year), wild rodents—namely the striped field mouse (A. agrarius), the yellow-necked mouse (A. flavicollis), the longtailed field mouse (A. sylvaticus) and the bank vole (M. glareolus)—were trapped using Sherman and wooden traps baited with a piece of fried bread with peanut butter and a piece of apple. This occurred in eight locations across the Czech Republic, Slovakia and Poland, spanning a range of environments (Fig. 1). Areas were selected due to the co-occurrence of the different species of rodents, meaning that the biotope of the locations was similar but they differed with regard to anthropopressure and biodiversity. There were two sites selected in the Czech Republic. The first was a nature reserve (CZ1), situated to the northwest ˇ of Ceské Budˇejovice, which includes four large ponds and vast areas of adjacent wetlands and meadows, and is characterised by rich and unique fauna and flora. The second, a

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suburban recreational area (CZ2) to the northeast of Brno, comprises meadows, forests and residential areas. The two rural areas selected in Slovakia—one close to the Danube river to the south of Bratislava (SK1) and one east of Koˇsice (SK2)—were both characterised by fields enclosed by forests. In Poland, there were four sites, all of which were located in the Lower Silesian region. Two were rural areas, the “Milicz Ponds” ornithological reserve (PL1) and ´ ez˙ a Landscape Park (PL4), and two were suburban areas, Sl˛ a water distribution area (PL2) and irrigation fields outside the Wroclaw agglomeration (PL3). Approximately 100 single traps were used, set out at 3 m intervals along transects. The trapping period lasted 2–3 days at each site; traps were inspected in the morning and just before dusk. The captured animals were brought to the laboratory, anaesthetised (Isofluranum) and euthanised by cervical dislocation (Gajda et al. 2017). Individual faecal and spleen samples were collected and stored in screw-cap microtubes containing 70% EtOH. Each sample was removed using a different pair of sterile dissection tools. Prior to DNA isolation, the tubes were centrifuged (16,000g; 10 min), supernatants were drained, and the remaining ethanol evaporated in a thermostat incubator at 60 ◦ C overnight. Molecular and phylogenetic analysis Total DNA was extracted from ∼200 mg of faeces or a whole spleen through bead disruption, using a mixture of 0.5 mm and 2.0 mm glass and zirconium beads for faecal samples and a mixture of 0.5 mm, 2.0 mm and 5.0 mm ® glass and zirconium beads for tissue samples in a FastPrep 24 Instrument (MP Biomedicals, CA, USA) (Sak et al. 2011), followed by isolation and purification using commercially available kits in accordance with the manufacturer’s instructions (GeneMatrix Stool DNA Purification Kit and GeneMATRIX Bio-Trace DNA Purification Kit; EURx, Gda´nsk, Poland). All DNA isolates were stored at −20 ◦ C until further usage. Nested PCR amplification was performed on two sets of primers, amplifying the ITS region of the rRNA gene of Encephalitozoon spp.: INT 580F and INT 580R; MSP3 and MSP4a with cycling parameters, according to KatzwinkelWladarsch et al. (1996). Each PCR mixture contained 1 × PCR buffer, 1.0 mM MgCl2 , 0.2 mM dNTPs, 2.5U Taq polymerase (Fermentas, Waltham, MA, USA), 1 ␮l BSA (10 mg/ml) and 0.25 M of each primer; BSA was excluded in the secondary step. For all PCR reactions, negative and positive controls were performed, with sterile water and reference DNA, respectively. Secondary PCR products were subjected to electrophoresis on a 1.0% agarose gel and stained with Midori Green (EURx). Samples that produced positive results were selected for sequencing, purified by using QIAquick PCR Purification Kit (Qiagen, Hilden, Germany), and then sequenced on Applied Biosystems ABI 3500 Sequencer (SEQme, Dobˇríˇs, Czech Republic).

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Fig. 1. Geographical distribution of samples obtained from wild living rodents. PL1 – “The Milicz Ponds” ornithological reserve (51◦ 31 N 17◦ 19 E); PL2 – water distribution area (51◦ 4 N 17◦ 6 E); PL3 – irrigation fields for ´ ez˙ a Landscape Park (50◦ 50 N 16◦ 44 E); CZ1 – Ceské ˇ the Wrocław agglomeration (51◦ 9 N 16◦ 58 E); PL4 – Sl˛ Budˇejovice (49◦ 00 N 14◦ 25 E); CZ2 – Brno (49◦ 13 N 16◦ 41 E); SK1 – Bratislava (48◦ 06 N 17◦ 10 E); SK2 – Koˇsice (48◦ 43 N 21◦ 18 E).

The nucleotide sequences obtained in this study were edited using DNA Baser Sequence Assembly software (Heracle BioSoft SRL, Romania). BLAST searches were conducted in order to elucidate any homologies with the previously deposited sequences in GenBank. The consensus sequences of Encephalitozoon spp. obtained from the examined animals were deposited in GenBank under accession numbers KX189630–KX189632. Statistical analysis Contingency tables were used to compare prevalence between the host species and the sampling areas using the chisquare test; p < 0.05 was considered statistically significant ® (STATISTICA 8, StatSoft Polska).

Results and Discussion Rodents are commonly found in both rural and urban areas where they have frequent opportunity to come into contact with domestic and wild animals, livestock, and humans. Wild living rodents act as reservoir hosts for microsporidia species. This was previously shown for Enterocytozoon bieneusi (Perec-Matysiak et al. 2015; Sak et al. 2011). Therefore, they represent a significant source of microsporidial spores in the environment (Sak et al. 2011). The spores are highly resistant to environmental factors and ingestion of contaminated food and water is the most common transmission route (Li et al. 2003; Meredith et al. 2015). There have been a few molecular studies which concern Encephalitozoon spp. in wild rodents. The rationale for this study was to examine if these hosts

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Table 1. Presence of specific DNA of Encephalitozoon cuniculi (Ec) in feacal samples and spleen tissues of wild living striped field mice (Apodemus agrarius), yellow-necked mice (Apodemus flavicollis), long-tailed field mice (Apodemus sylvaticus) and bank voles (Myodes glareolus) in the Czech Republic, Slovakia and Poland. Species

Country

Locality

No. of examined/positive animals (%)

Positive samples Faeces

Apodemus agrarius

No.

Genotyping

17 × Ec II 1 × Ec II 2 × Ec II 7 × Ec II 4 × Ec III 1 × Ec II 2 × Ec II

0 0 0 5

– – – 1 × Ec II 4 × Ec III 1 × Ec II 2 × Ec II

72/17 (23.6) 22/1 (4.5) 33/2 (6) 61/11 (18)

17 1 2 11

Slovakia

SK1 SK2

7/1 (14.3) 8/2 (25) 203/34 (16.7)

1 2 34

CZ1 CZ2 PL1 PL2 PL3 PL4 SK1 SK2

15/3 (20) 23/5 (21.7) 33/7 (21.2) 28/2 (7.1) 9/0 (0.0) 9/4 (44.4) 19/4 (21) 13/2 (15.3) 149/27 (18.1)

2 5 6 2 0 2 4 2 23

2 × Ec II 5 × Ec II 6 × Ec II 2 × Ec II – 2 × Ec II 4 × Ec II 2 × Ec II

3 3 1 0 0 2 3 2 14

3 × Ec II 3 × Ec II 1 × Ec II – – 2 × Ec II 3 × Ec II 2 × Ec II

CZ1 CZ2 SK1 SK2

15/2 (13.3) 9/1 (11.1) 13/3 (23.1) 8/0 (0.0) 45/6 (13.3)

2 1 3 0 6

2 × Ec II 1 × Ec II 3 × Ec II –

1 1 2 0 4

1 × Ec II 1 × Ec II 2 × Ec II –

CZ2 PL1 PL2 PL3 PL4 SK1 SK2

12/1 (8.3) 4/0 (0.0) 20/0 (0.0) 4/0 (0.0) 7/0 (0.0) 15/2 (13.3) 6/0 (0.0) 68/3(4.4) 465/70 (15.1)

1 0 0 0 0 1 0 2 65

1 × Ec II – – – – 1 × Ec I –

1 0 0 0 0 1 0 2 28

1 × Ec II – – – – 1 × Ec II –

The Czech Republic

Slovakia Subtotal The Czech Republic Slovakia Subtotal Myodes glareolus

Genotyping

PL1 PL2 PL3 PL4

Poland

Apodemus sylvaticus

No. Poland

Subtotal Apodemus flavicollis

Spleen

The Czech Republic Poland

Slovakia Subtotal Total

1 2 8

Bold values emphasize and summarize the particular hosts (as subtotals) and the obtained results (as total).

can be considered as an important source of Encephalitozoon spp., which is infectious to humans and animals. The faecal samples and spleen tissues obtained from 465 rodent individuals were analysed for the presence of Encephalitozoon spp. by the nested PCR method (Table 1). Molecular analysis showed that the prevalence of Encephalitozoon spp. (positive for faeces and/or spleen) was 15.1% over all sampled rodents and sites. The highest infection rate was recorded for A. flavicollis (18.1%), and the lowest for M. glareolus (4.4%); the difference, with regard to host species, was significant (χ2 = 7.91; df = 3; p = .04). A higher infection rate was observed in the faecal samples (14%) than in the spleen samples (6%) of infected rodents and the occurrence of Encephalitozoon spp. in faeces was found to be signif-

icant with regard to the rodent species (χ2 = 8.522; df = 3; p = 0.04). In 32.9% of infected rodents, Encephalitozoon spp. was recorded for both faeces and spleen while, in the remaining group of rodents, the microsporidium was detected in either faecal or spleen samples. Differences in infection rates between study sites were not significant (p > 0.05). Sequence analyses showed that E. cuniculi was the only species recorded in this study. The alignment of the obtained ITS sequences with those previously deposited in GenBank showed 100% homology with E. cuniculi genotype II (Acc. No. GU213880) or E. cuniculi genotype III (Acc. No. KX925860). The most frequently identified E. cuniculi genotype II was detected in three Apodemus species, whereas E. cuniculi genotype III was recorded only in A. agrarius. Mixed

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infection, with genotype II and genotype III identified in faeces and spleen tissue, was recorded in three individuals of the A. agrarius species. Additionally, one isolate obtained from M. glareolus shared 100% identity with E. cuniculi genotype I (Acc. No. KJ941140). The genotypes of E. cuniculi initially appeared to exhibit geographical and host preferences but increasing numbers of descriptions of the genotypes in different hosts and from different geographic areas made this trend less clear (Fayer and Santin-Duran 2014). Encephalitozoon cuniculi genotypes I, II, and III have been identified in both humans and animals, supporting the potential for zoonotic transmission (Didier et al. 2000; Mathis et al. 2005; Sokolova et al. 2011). Tsukada et al. (2013) showed 24.1% prevalence of Encephalitozoon spp. in Japanese wild rodents, with the highest infection rate observed for the wood mouse Apodemus speciosus (32.5%), followed by the Japanese grass vole Microtus montebelli (18.9%), and the small Japanese field mouse Apodemus argenteus (12.5%). Three species of this pathogen were detected: E. hellem and E. cuniculi genotypes I and III were the most frequent among Japanese rodents, along with E. intestinalis; E. cuniculi genotype II was not detected as it has only been confirmed in Europe to date. Sak et al. (2011) showed Encephalitozoon spp. infection rate, similar to those recorded in Japan, but in the house mouse Mus musculus (24.6%), and identified E. hellem and E. cuniculi genotypes I and II. Lower infection rates of E. cuniculi were recorded in Microtus arvalis (6%) and Arvicola terrestris (7%) in Austria (Fuehrer et al. 2010). Surveys in Slovakia detected Encephalitozoon spp. (E. cuniculi genotype I and E. intestinalis) in only 0.7% of the faecal samples of the house mouse (Daniˇsová et al. 2015). In the present study, no other Encephalitozoon species besides E. cuniculi genotypes I, II, and III were identified. Encephalitozoon cuniculi genotype III seems to be rarely identified in rodents (Tsukada et al. (2013); this study). Surprisingly, we did not detect E. hellem in any of the samples from the ornithological reserve (PL1 locality), even though the presence of waterfowl and other birds would be expected to increase the risk of E. hellem infections in rodents. The recorded prevalence of E. cuniculi in that study area was quite high for A. flavicollis (21.2%) and A. agrarius (23.6%). Our previous survey (Perec-Matysiak et al. 2015), conducted at the same locality and concerning the occurrence of E. bieneusi in the same species of rodents, also showed high infection rates of E. bieneusi for A. flavicollis and A. agrarius, at the levels of 32% and 27%, respectively. In the present survey, the highest infection rate (44%) was recorded in A. flavicollis obtained from the rural site (PL4) which is used as a recreational area for humans. Our findings demonstrate that wild living rodents of the Murinae and Arvicolinae subfamilies contribute to the spread of spores of microsporidian species which are hazardous for the health of humans and animals; the survey also presents data on the occurrence of Encephalitozoon in new host species of rodents obtained from Central European countries.

Statement on collection of the animals All of the procedures were conducted in accordance with the laws of the Czech Republic, Poland and Slovakia on the use of animals, safety and the use of pathogenic agents. The research was conducted under ethical protocols approved by Local Ethics Committees (protocol nos. 46/2008, 071/2010, 48/2012, 114/2013 and 52/2014).

Author contributions A.PM, M.K. and B.S. conceptualised the project. A.PM., ˇ C. ˇ carried out the research. K.BG. perK.L., M.K. and S. ˇ C. ˇ and formed statistical analysis. A.PM., K.L., K.BG., S. J.H. trapped the rodents and collected samples. A.PM. wrote the manuscript. A.PM, M.K. and B.S. revised the manuscript. All authors read and approved the final manuscript.

Acknowledgments This study was co-financed by the European Union under the European Social Fund and the Grant Agency of the University of South Bohemia (project No. 002/2016/Z) and the Czech Science Foundation (17-12871S).

References Åkerstedt, J., Nordstoga, K., Mathis, A., Smeds, E., Deplazes, P., 2002. Fox encephalitozoonosis: isolation of the agent from an outbreak in farmed blue foxes (Alopex lagopus) in Finland and some hitherto unreported pathologic lesions. J. Vet. Med. B 49, 400–405, http://dx.doi.org/10.1046/j.1439-0450.2002.00588.x. Bornay-Llinares, F.J., da Silva, A.J., Moura, H., Schwartz, D.A., Visvesvara, G.S., Pieniazek, N.J., Cruz-López, A., Hernández-Jaúregui, P., Guerrero, J., Javier Enriquez, F.J., 1998. Immunologic, microscopic, and molecular evidence of Encephalitozoon intestinalis (Septata intestinalis) infection in mammals other than humans. J. Infect. Dis. 178, 820–826, http://dx.doi.org/10.1086/515356. Childs-Sanford, S.E., Garner, M.M., Raymond, J.T., Didier, E.S., Kollias, G.V., 2006. Disseminated microsporidiosis due to Encephalitozoon hellem in an Egyptian fruit bat (Rousettus aegyptiacus). J. Comp. Pathol. 134, 370–373, http://dx.doi.org/10.1016/j.jcpa.2006.01.004. Daniˇsová, O., Valenˇcáková, A., Stanko, M., Luptáková, L., Hasajová, A., 2015. First report of Enterocytozoon bieneusi and Encephalitozoon intestinalis infection of wild mice in Slovakia. Ann. Agr. Env. Med. 22, 251–252, http://dx.doi.org/10.5604/12321966.1152075. Didier, E.S., Weiss, L.M., 2011. Microsporidiosis: not just in AIDS patients. Curr. Opin. Infect. Dis. 24, 490–495, http://dx.doi.org/10.1097/QCO.0b013e32834aa152. Didier, E.S., Vossbrinck, C.R., Baker, M.D., Rogers, L.B., Bertucci, D.C., Shadduck, J.A., 1995. Identification and characterization of three Encephalitozoon cuniculi strains. Parasitology 111, 411–421.

A. Perec-Matysiak, K. Le´snia´nska, K. Bu´nkowska-Gawlik, et al. / European Journal of Protistology 69 (2019) 14–19

Didier, E.S., Visvesvara, G.S., Baker, M.D., Rogers, L.B., Bertucci, D.C., De Groote, M.A., Vossbrinck, C.R., 1996. A microsporidian isolated from an AIDS patient corresponds to Encephalitozoon cuniculi III, originally isolated from domestic dogs. J. Clin. Microbiol. 34, 2835–2837. Didier, E.S., Didier, P.J., Snowden, K.F., Shadduck, J.A., 2000. Microsporidiosis in mammals. Microbes Infect. 2, 709–720, http://dx.doi.org/10.1016/S1286-4579(00)00354-3. Didier, E.S., Stovall, M.E., Green, L.C., Brindley, P.J., Sestak, K., Didier, P.J., 2004. Epidemiology of microsporidiosis: sources and modes of transmission. Vet. Parasitol. 126, 145–166, http://dx.doi.org/10.1016/j.vetpar.2004.09.006. Didier, E.S., 2005. Microsporidiosis: an emerging and opportunistic infection in humans and animals. Acta Trop. 94, 61–76, http://dx.doi.org/10.1016/j.actatropica.2005.01.010. Fayer, R., Santin-Duran, M., 2014. Epidemiology of microsporidia in human infections. In: Weiss, L.M., Becnel, J.J. (Eds.), Microsporidia. Pathogens of opportunity. Wiley Blackwell, pp. 135–164. Fuehrer, H.P., Blöschl, I., Siehs, C., Hassl, A., 2010. Detection of Toxoplasma gondii, Neospora caninum, and Encephalitozoon cuniculi in the brains of common voles (Microtus arvalis) and water voles (Arvicola terrestris) by gene amplification techniques in western Austria (Vorarlberg). Parasitol. Res. 107, 469–473, http://dx.doi.org/10.1007/s00436-010-1905-z. Gajda, E., Hildebrand, J., Sprong, H., Bu´nkowska-Gawlik, K., Perec-Matysiak, A., Coipan, E.C., 2017. Spotted fever rickettsiae in wild-living rodents from south-western Poland. Parasit. Vectors 10, 413, http://dx.doi.org/10.1186/s13071-017-2356-5. Graczyk, T.K., Bosco-Nizeyi, J., da Silva, A.J., Moura, I.N., Pieniazek, N.J., Cranfield, M.R., Lindquist, H.D., 2002. A single genotype of Encephalitozoon intestinalis infects free-ranging gorillas and people sharing their habitats in Uganda. Parasitol. Res. 88, 926–931, http://dx.doi.org/10.1007/s00436-002-0693-5. Hersteinsson, P., Gunnarsson, E., Hjartardottir, S., Skirnisson, K., 1993. Prevalence of Encephalitozoon cuniculi antibodies in terrestrial mammals in Iceland, 1986 to 1989. J. Wildl. Dis. 29, 341–344, http://dx.doi.org/10.7589/0090-3558-29.2.341. Hinney, B., Sak, B., Joachim, A., Kváˇc, M., 2016. More than a rabbit´s tale — Encephalitozoon spp. in wild mammals and birds. Int. J. Parasitol.: Parasites Wildl. 5, 76–87, http://dx.doi.org/10.1016/j.ijppaw.2016.01.001. Juan-Salles, C., Garner, M.M., Didier, E.S., Serrato, S., Acevedo, L.D., Ramos-Vara, J.A., Nordhausen, R.W., Bowers, L.C., Parás, A., 2006. Disseminated encephalitozoonosis in captive, juvenile, cotton-top (Saguinus oedipus) and neonatal emperor (Saguinus imperator) tamarins in North America. Vet. Pathol. 43, 438–446, http://dx.doi.org/10.1354/vp.43-4-438. Katzwinkel-Wladarsch, S., Lieb, M., Helse, W., Loscher, T., Rinder, H., 1996. Direct amplification and species determination of microsporidian DNA from stool specimens. Trop. Med. Int. Health. 1, 373–378, http://dx.doi.org/10.1046/j.1365-3156.1996. d01-51.x. Li, X., Palmer, R., Trout, J.M., Fayer, R., 2003. Infectivity of microsporidia spores stored in water at environmen-

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tal temperatures. J. Parasitol. 89, 185–188, http://dx.doi.org/ 10.1645/0022-3395(2003)089[0185:IOMSSI2.0.CO;2]. Müller-Doblies, U.U., Herzog, K., Tanner, I., Mathis, A., Deplazes, P., 2002. First isolation and characterisation of Encephalitozoon cuniculi from a free-ranging rat (Rattus norvegicus). Vet. Parasitol. 107, 279–285, http://dx.doi.org/10.1016/s0304-4017(02)00163-2. Mathis, A., Åkerstedt, J., Tharaldsen, J., Odegaard, O., Deplazes, P., 1996. Isolates of Encephalitozoon cuniculi from farmed blue foxes (Alopex lagopus) from Norway differ from isolates from Swiss domestic rabbits (Oryctolagus cuniculus). Parasitol. Res. 82, 27–30, http://dx.doi.org/10.1007/s004360050192. Mathis, A., Weber, R., Deplazes, P., 2005. Zoonotic potential of the microsporidia. Clin. Microbiol. Rev. 18, 423–445, http://dx.doi.org/10.1128/CMR.18.3.423-445.2005. Meredith, A.L., Cleaveland, S.C., Brown, J., Mahajan, A., Shaw, D.J., 2015. Seroprevalence of Encephalitozoon cuniculi in wild rodents, foxes and domestic cats in three sites in the United Kingdom. Transbound. Emerg. Dis. 62, 148–156, http://dx.doi.org/10.1111/tbed.12091. Murphy, T.M., Walochnik, J., Hassl, A., Moriartyc, J., Mooneya, J., Tooland, D., Sanchez-Miguele, C., O’Loughlinf, A., McAuliffea, A., 2007. Study on the prevalence of Toxoplasma gondii and Neospora caninum and molecular evidence of Encephalitozoon cuniculi and Encephalitozoon (Septata) intestinalis infections in red foxes (Vulpes vulpes) in rural Ireland. Vet. Parasitol. 146, 227–234, http://dx.doi.org/10.1016/j.vetpar.2007.02.017. Perec-Matysiak, A., Bu´nkowska-Gawlik, K., Kváˇc, M., Sak, B., Hildebrand, J., Le´snia´nska, K., 2015. Diversity of Enterocytozoon bieneusi genotypes among small rodents in southwestern Poland. Vet. Parasitol. 214, 242–246, http://dx.doi.org/10.1016/j.vetpar.2015.10.018. Sak, B., Kváˇc, M., Kvˇetoˇnová, D., Albrecht, T., Piálek, J., 2011. The first report on natural Enterocytozoon bieneusi and Encephalitozoon spp. infections in wild East-European House Mice (Mus musculus musculus) and West-European House Mice (M. m. domesticus) in a hybrid zone across the Czech Republic-Germany border. Vet. Parasitol. 178, 246–250, http://dx.doi.org/10.1016/j.vetpar.2010.12.044. Si´nski, E., 2003. Environmental contamination with protozoan parasite infective stages: biology and risk assessment. Acta Microbiol. Pol. 52, 97–107. Sokolova, O.I., Demyanov, A.V., Bowers, L.C., Didier, E.S., Yakovlev, A.V., Skarlato, S.O., Sokolova, Y.Y., 2011. Emerging microsporidian infections in Russian HIVinfected patients. J. Clin. Microbiol. 49, 2102–2108, http://dx.doi.org/10.1128/JCM.02624-10. Talabani, H., Sarfati, C., Pillebout, E., van Gool, T., Derouin, F., Menotti, J., 2010. Disseminated infection with a new genovar of Encephalitozoon cuniculi in a renal transplant recipient. J. Clin. Microbiol. 48, 2651–2653, http://dx.doi.org/10.1128/JCM.02539-09. Tsukada, R., Tsuchiyama, A., Sasaki, M., Park, C.H., Fujii, Y., Takesue, M., Hatai, H., Kudo, N., Ikadai, H., 2013. Encephalitozoon infections in Rodentia and Soricomorpha in Japan. Vet. Parasitol. 198, 193–196, http://dx.doi.org/10.1016/j.vetpar.2013.08.018.