CC

MJa

b

c

d

e

ARRA

KCTMMTC

1

ltCiM

tC

0h

Veterinary Parasitology 201 (2014) 9–17

Contents lists available at ScienceDirect

Veterinary Parasitology

jo u r nal homep age: www.elsev ier .com/ locate /vetpar

ryptosporidium erinacei n. sp. (Apicomplexa:ryptosporidiidae) in hedgehogs

artin Kváca,b,∗, Lada Hofmannovác,d, Lenka Hláskováa, Dana Kvetonováa,irí Vítovecb, John McEvoye, Bohumil Saka

Institute of Parasitology, Biology Centre of the Academy of Sciences of the Czech Republic, v.v.i., Ceské Budejovice, Czech RepublicFaculty of Agriculture, University of South Bohemia in Ceské Budejovice, Czech RepublicDepartment of Pathology and Parasitology, University of Veterinary and Pharmaceutical Sciences, Brno, Czech RepublicCEITEC – VFU, University of Veterinary and Pharmaceutical Sciences, Brno, Czech RepublicVeterinary and Microbiological Sciences Department, North Dakota State University, Fargo, USA

a r t i c l e i n f o

rticle history:eceived 6 September 2013eceived in revised form 11 January 2014ccepted 19 January 2014

eywords:ryptosporidium erinaceiaxonomyorphology

a b s t r a c t

The morphological, biological, and molecular characteristics of Cryptosporidium hedgehoggenotype are described, and the species name Cryptosporidium erinacei n. sp. is proposed toreflect its specificity for hedgehogs under natural and experimental conditions. Oocysts ofC. erinacei are morphologically indistinguishable from Cryptosporidium parvum, measuring4.5–5.8 �m (mean = 4.9 �m) × 4.0–4.8 �m (mean = 4.4 �m) with a length to width ratio of1.13 (1.02–1.35) (n = 100). Oocysts of C. erinacei obtained from a naturally infected Europeanhedgehog (Erinaceus europaeus) were infectious for naïve 8-week-old four-toed hedgehogs(Atelerix albiventris); the prepatent period was 4–5 days post infection (DPI) and the patentperiod was longer than 20 days. C. erinacei was not infectious for 8-week-old SCID and

olecular analyses

ransmission studiesryptosporidium hedgehog genotype

BALB/c mice (Mus musculus), Mongolian gerbils (Meriones unguiculatus), or golden ham-sters (Mesocricetus auratus). Phylogenetic analyses based on small subunit rRNA, 60 kDaglycoprotein, actin, Cryptosporidium oocyst wall protein, thrombospondin-related adhe-sive protein of Cryptosporidium-1, and heat shock protein 70 gene sequences revealed thatC. erinacei is genetically distinct from previously described Cryptosporidium species.

© 2014 Elsevier B.V. All rights reserved.

. Introduction

Protozoans of the genus Cryptosporidium are epicellu-ar parasites infecting a gastrointestinal and/or respiratoryract of a broad range of vertebrates. Greater than 24 validryptosporidium species have been described in amphib-

ans, reptiles, birds, and mammals (Ryan and Power, 2012).uch of Cryptosporidium diversity is found associated with

∗ Corresponding author at: Institute of Parasitology, Biology Centre ofhe Academy of Sciences of the Czech Republic, v.v.i., Ceské Budejovice,zech Republic. Tel.: +42 0387775419.

E-mail address: kvac@paru.cas.cz (M. Kvác).

304-4017/$ – see front matter © 2014 Elsevier B.V. All rights reserved.ttp://dx.doi.org/10.1016/j.vetpar.2014.01.014

wild mammals, and only a fraction of this diversity has beendescribed to the species level.

Hedgehog cryptosporidiosis was first detected in 1998in a juvenile captive four-toed hedgehog (Ateletrix albiven-tris) from the Baltimore Zoo (Maryland, USA) (Graczyket al., 1998). Cryptosporidium oocysts have since beenidentified in the feces of juvenile and adult Europeanhedgehogs (Erinaceus europaeus) in the United Kingdom(Meredith and Milne, 2009; Sturdee et al., 1999), Germany(Pantchev and Möller, 2007), and Denmark (Enemark et al.,2002). An isolate from a European hedgehog originating

from Denmark was genotyped by partial amplificationand sequencing of the Cryptosporidium oocyst wall pro-tein (COWP) gene, the 18S ribosomal RNA gene, and amicrosatellite locus, and was found to be distinct from C.

y Parasi

10 M. Kvác et al. / Veterinar

parvum and other Cryptosporidium species (Enemark et al.,2002).

In the first large-scale molecular study of Cryptosporid-ium in hedgehogs (Dyachenko et al., 2010), 30.0% (56/188)of European hedgehogs were found to be shedding oocystsof C. parvum or a phylogenetically distinct genotype, whichwas named Cryptosporidium hedgehog genotype.

Cryptosporidium hedgehog genotype appears to be rel-atively specific for hedgehogs, and was detected onlyrarely in horses in Algeria (Laatamna et al., 2013). How-ever, the pathogenicity of this genotype for hedgehogsremains unclear. Although there have been three reports ofdiarrhea in Cryptosporidium positive hedgehogs (Graczyket al., 1998; Meredith and Milne, 2009; Pantchev andMöller, 2007), the Cryptosporidium species/genotypes caus-ing the diarrhea were not identified. In addition, the courseof infection, localization of endogenous life-cycle stages,and morphometry of oocysts causing cryptosporidiosis inhedgehogs have not yet been described.

No other names have been used for Cryptosporid-ium hedgehog genotype in publications or GenBank.Dyachenko et al. (2010) named subtypes of the 60 kDa gly-coprotein gene (gp60) as family VII. However, in a parallelpublication by Lv et al. (2009), gp60 sequences from Cryp-tosporidium wrairi also were named subtype family VII.Laatamna et al. (2013) proposed to continue using the sub-type family designation VII for C. wrairi (valid species) andto introduce a novel family XIII for the Cryptosporidiumhedgehog genotype. A search of GenBank entries on June12, 2013, revealed 8, 8, 4 and 3 partial sequences of SSU(small subunit rRNA), gp60 (60 kDa glycoprotein), actin,and HSP70 (heat shock protein 70) genes, respectively.

We undertook this study to determine the experimentaltransmission, oocyst morphology, and molecular charac-teristics of Cryptosporidium hedgehog genotype. Based onthe collective data from this and other studies, which showthat Cryptosporidium hedgehog genotype is adapted tohedgehogs, and is genetically and biologically distinct fromknown Cryptosporidium species, we propose the speciesname Cryptosporidium erinacei n. sp.

2. Materials and methods

2.1. Source of oocysts for transmission studies

Oocysts of C. erinacei originated from a natu-rally infected juvenile European hedgehog (Erinaceuseuropaeus), which was born in August/September 2012and was rescued for its low weight in November 2012.Hedgehog feces was screened for endoparasites usingstandard parasitological methods (data not shown)including microscopic examination of fecal smears stainedwith aniline–carbol–methyl violet (ACMV) for presenceCryptosporidium oocysts (Milácek and Vítovec, 1985).Fecal specimens were collected daily and stored in a 2.5%potassium dichromate solution at 8 ◦C. Cryptosporidiumoocysts were purified for morphometry, phylogeny, and

infectivity analyses using sucrose gradient (Arrowoodand Sterling, 1987) and cesium chloride gradient cen-trifugation (Kilani and Sekla, 1987). Purified oocystswere stored for up to 4-weeks in PBS with antimycotics

tology 201 (2014) 9–17

and antibiotics at 4 ◦C in darkness. The parasite wasconfirmed to be the Cryptosporidium hedgehog genotypeby sequence analysis of the SSU gene, using the methoddescribed below. The number of oocysts administeredto animals was determined by hemocytometer counting.The viability of oocysts was examined using propidiumiodide (PI) staining by a modified assay of Sauch et al.(1991). Examined oocysts were washed in distilled water(DW; 100,000 oocysts in 100 �l) and mixed with 10 �lof PI (1% solution, SIGMA). After 30 min of incubation atroom temperature in the dark, the oocysts were washedtwice with DW. Oocyst viability was examined usingfluorescence microscopy (filter 420 nm, Olympus IX70).Oocysts with red fluorescence were considered to be dead,and those without fluorescence were considered viable. Atotal of 5 × 100 oocysts were counted.

2.2. Oocyst morphology

Oocysts were examined using differential interferencecontrast (DIC) microscopy following ACMV staining, andfluorescence microscopy following labeling with genus-specific FITC-conjugated antibodies (Cryptosporidium IFTest, Crypto cel, Medac) (Olympus IX70 microscope; Olym-pus CZ, Czech Republic). Morphology and morphometrywere determined using digital analysis of images (M.I.C.Quick Photo Pro v.1.2 soft-ware; Optical Service, CzechRepublic) collected using an Olympus Camedia C 5060WIDEZOOM 5.1 megapixel digital camera (Optical Ser-vice). A 20-�l aliquot containing 10,000 purified oocystswas examined for each measurement. Length and widthof oocysts (n = 100) were measured under DIC at 1000×magnification, and these were used to calculate the shapeindex and length-to-width ratio of each oocyst. As a control,the morphometry of C. parvum (n = 100) from a natu-rally infected 25-day-old Holstein calf was measured bythe same person using the same microscope. Photomicro-graphs of C. erinacei oocysts observed by DIC, ACMV andIFA were deposited as a phototype at the Institute of Para-sitology, Biology Centre of the Academy of Sciences of theCzech Republic.

2.3. DNA extraction and molecular analyses

Total DNA was extracted from 200 mg of feces, 50 �lof purified oocysts, or 200 mg of tissue by bead dis-ruption for 60 s at 5.5 m/s using 0.5 mm glass beadsin a FastPrep®24 Instrument (MP Biomedicals, CA, USA)followed by isolation/purification using a commerciallyavailable kit in accordance with the manufacturer’s instruc-tions (QIAamp® DNA Stool Mini Kit or DNeasy® Blood &Tissue Kit, Qiagen, Hilden, Germany). Purified DNA wasstored at −20 ◦C prior to being used for PCR. A nestedPCR approach was used to amplify a region of the smallribosomal subunit (SSU; ∼830 bp; Jiang et al., 2005; Xiaoet al., 1999), actin (∼1066 bp; Sulaiman et al., 2002), Cryp-tosporidium Oocyst Wall Protein (∼550 bp; Pedraza-Diaz

et al., 2001; Spano et al., 1997), Thrombospondin-related adhesive protein of Cryptosporidium-1 (TRAP-C1;∼780 bp; Spano et al., 1998), Heat shock protein (HSP70;∼1950 bp; Sulaiman et al., 2000), and 60 kDa glycoprotein

y Parasi

(PtwcoS(sEsasci

2

t1eBm7siwby5MShb

2

2

mohBRhfis

2

li(tdwbsw

M. Kvác et al. / Veterinar

∼830 bp; Alves et al., 2003). Both primary and secondaryCR reactions were carried out in a volume of 50 �l;he primary reaction contained 2 �l of genomic DNA (orater as a negative control) and the secondary reaction

ontained 2 �l of the primary reaction as template. DNAf Cryptosporidium hominis was used as positive control.econdary PCR products were detected by agarose gel2.0%) electrophoresis, visualized by ethidium bromidetaining (0.2 �g/ml) and extracted using QIAquick® Gelxtraction Kit (Qiagen). Purified secondary products wereequenced in both directions with an ABI 3130 geneticnalyser (Applied Biosystems, Foster City, CA) using theecondary PCR primers and the BigDye1 Terminator V3.1ycle sequencing kit (Applied Biosystems, Foster City, Cal-fornia) in 10 �l reactions.

.4. Phylogenetic analyses

The nucleotide sequences of each gene obtained inhis study were edited using the program ChromasPro.7.5 (Technelysium, Pty, Ltd.), and were aligned withach other and with reference sequences from Gen-ank using ClustalX 2.0.6. Alignment adjustments wereade manually to remove artificial gaps using BioEdit

.0.5.3. Phylogenetic analyses were performed using theoftware MEGA5 (Tamura et al., 2011). Neighbor join-ng (NJ) trees were constructed. All ambiguous positions

ere removed for each sequence pair. The reliability ofranches in trees was assessed using the bootstrap anal-sis with 1000 pseudoreplicates. Bootstrap values above0% were reported. Phylograms were drawn using theEGA5 and were manually adjusted using CorelDrawX5.

SU, actin, COWP, TRAP-C1, HSP70 and gp60 sequencesave been deposited in GenBank under the accession num-ers KF612324–KF612329.

.5. Transmission studies

.5.1. AnimalsFive 8-week-old SCID mice (strain C.B-17) and BALB/c

ice (Charles River, Germany), Mongolian gerbils (Meri-nes unguiculatus) (Charles River, Germany), and goldenamsters (Mesocricetus auratus) (Institute of Parasitology,iology Centre of the Academy of Sciences of the Czechepublic, Czech Republic); one 3-months-old Europeanedgehog (E. europaeus) (rescued); and three 8-week-old

our-toed hedgehogs (A. albiventris) (private; Cryptosporid-um free breed) were used for experimental infectiontudies.

.5.2. Experimental designTo prevent environmental contamination with oocysts,

aboratory rodents were housed in plastic cages with ster-lized wood-chip bedding situated in flexible film isolatorsBEM, Znojmo, Czech Republic) with high-efficiency par-iculate air filters. The mice were supplied with a sterilizediet (TOP-VELAZ, Prague, Czech Republic) and sterilized

ater ad libitum. Hedgehogs were individually kept in the

oxes with sterilized wood-chip bedding and hay, and wereupplied with a diet of sterilized cat food and sterilizedater ad libitum. For one week prior to infection, fecal

tology 201 (2014) 9–17 11

samples from all animals were screened daily for thepresence of Cryptosporidium spp. using parasitological andmolecular tools, as described in previous sections. Rodents(five animals per group) were each inoculated orallyby stomach tube with 1,000,000 purified viable oocystssuspended in 200 �l of distilled water. Three four-toedhedgehogs received the same infection dose in a 1 mlsuspension via syringe. Fecal samples from all experi-mental animals were collected daily and all experimentswere terminated 20 days post infection (DPI), with theexception of one hedgehog (no. 3), which was sacri-ficed on day 7 post infection. Samples were stained withaniline–carbol–methyl violet (Milácek and Vítovec, 1985)and the presence of Cryptosporidium specific DNA was con-firmed using nested PCR targeting the SSU gene. Courseof infection indicators, including fecal consistency, fecalcolor and infection intensity, were examined daily. Infec-tion intensity was reported as the number of oocysts pergram (OPG) of feces as previously described (Kvác et al.,2007).

2.5.3. Clinical and histopathological examinationsThe complete examination of all gastrointestinal organs

was conducted at necropsy. Tissue specimens from thestomach, small and large intestine (the entire tract wasdivided into 1 cm sections) were sampled and processedfor histology according to Kvác and Vítovec (2003) andfor PCR analyses (see Section 2.3). Histology sections werestained with hematoxylin and eosin (HE), Wolbach’s mod-ified Giemsa stain, and genus-specific FITC conjugatedmonoclonal antibodies targeting Cryptosporidium oocystwall antigens (Cryptosporidium IF Test, Crypto cel, Medac).

2.5.4. Animal careAnimal caretakers wore new disposable coveralls, shoe

covers, and gloves every time they entered the buildings.All wood-chip bedding, feces, and disposable protectiveclothing were sealed in plastic bags, removed from thebuildings and incinerated. All housing, feeding, and exper-imental procedures involving hedgehogs, laboratory miceof different strains, Mongolian gerbils, and golden hamsterswere conducted under protocols approved by the Instituteof Parasitology, Biology Centre of the Academy of Sciencesof the Czech Republic and Institute and National Commit-tees (Protocol No. 071/2010 and 114/2013).

3. Results

3.1. Oocyst morphology

Oocysts of C. erinacei measuring 4.5–5.8(mean = 4.9 �m) × 4.0–4.8 �m (mean = 4.4) with a length towidth ratio of 1.13 (1.02–1.35) (n = 100) were similar in sizeto C. parvum, measuring 5.1–5.4 (mean = 5.3 �m) × 4.6–5.1(mean = 4.7 �m) with length to width ratio of 1.12 (1.05–1.33) (n = 100) (Fig. 1A). Oocysts of C. erinacei recovered fromexperimentally infected hedgehogs were morphologically

similar to those used for infection. Oocysts in fecal smearsshowed typical Cryptosporidium ACMV staining character-istics (Fig. 1B). Fixed C. erinacei oocysts labeled with FITCconjugated anti-Cryptosporidium oocyst wall antibody

12 M. Kvác et al. / Veterinary Parasitology 201 (2014) 9–17

alized iium FITC

Fig. 1. Cryptosporidium erinacei and Cryptosporidium parvum oocysts visuand stained by (B) aniline–carbol–methyl violet and (C) anti-Cryptosporid

and examined by epifluorescence microscopy displayedtypical apple green, halo-like fluorescence (Fig. 1C).

3.2. Molecular characterization

Isolates obtained from naturally infected Europeanhedgehogs shared 100% sequence identity at the SSU,actin, COWP, HSP70, TRAP-C1, and gp60 loci with iso-lates recovered from experimentally infected four-toedhedgehogs. SSU sequences were identical to the Gen-Bank sequence of a hedgehog genotype obtained from ahorse in Algeria (KC305650) and were closely related tosequences of hedgehog genotypes (GQ259141, GQ214082,and GQ214078) originally isolated from hedgehogs inGermany (Fig. 2A). The actin sequence was identical toa sequence from a hedgehog in Germany (GQ214079;Fig. 2B). Analyses of the gp60 gene locus revealed iden-tity with sequences belonging to family XIIIa (Dyachenkoet al., 2010; Laatamna et al., 2013; Lv et al., 2009) (Fig. 2C).The gp60 subtype of the identified hedgehog genotype had21 serine-coding TCA repeats immediately followed by 11repeats of the sequence ACATCA, and was therefore namedXIIIa A21R11, based on the established gp60 nomencla-ture (Sulaiman et al., 2005). Previously reported C. erinaceisubtypes include A21R10 (GQ214085), A22R9 (KC305644),A19R12 (GQ214081), and A22R11 (GQ259140). Outsidethe hypervariable microsatellite region, C. erinacei gp60sequences are identical, with the exception of GQ214081,which has a single nucleotide polymorphism (G–A) result-ing in an amino acid change from methionine to valine.The original (field) isolate and all experimentally recov-ered isolates produced amplicons that were identicalto C. parvum sequences at the COWP gene (GU904398,Fig. 3A). The HSP70 sequence was 99.2 and 99.6% simi-lar to sequences GQ214080 and GQ214084, respectively,

which were obtained from hedgehogs in Germany (Fig. 3B).The TRAP-C1 gene sequence from C. erinacei, which wasclosely related to C. parvum, is the first TRAP-C1 sequencereported from this species (Fig. 3C). Neighbor-joining trees

n various preparations: (A) differential interference contrast microscopy-conjugated antibody. Bar = 5 �m.

constructed from SSU, actin, COWP, HSP70 and TRAP-C1sequences support the position of C. erinacei as a separatephylogenetic group that is most similar to C. parvum, C.hominis, and Cryptosporidium tyzzeri (Figs. 2 and 3).

3.3. Experimental transmission studies

Oocyst used for experimental infections had >93% via-bility determined by PI staining. Experimentally inoculatedSCID and BALB/c mice, Mongolian gerbils and golden ham-sters did not produce detectable C. erinacei infection bymicroscopy, histology, or PCR during 20 DPI. No clinicalsigns of cryptosporidiosis were detected in any labora-tory rodents. Histological and molecular examination ofgastrointestinal tract tissue from these rodents did notreveal the presence of Cryptosporidium developmentalstages.

C. erinacei was fully infectious for all three 8-week-old four-toed hedgehogs. The first oocyst shedding wasdetected at 4 DPI and continued until 20 DPI of the study.Infection intensity ranged from 2000 to 670,000 oocyst pergram (OPG) with maximum shedding at 7 DPI. From 12DPI the infection was intermittent based on microscopydetection (OPG under detection limit), although specificDNA was present in feces during the patent period (Fig. 4).Infected hedgehogs showed no symptoms of the diseaseand hedgehogs necropsied at 7 and 20 DPI showed nomacroscopic signs of intestinal cryptosporidiosis. SpecificC. erinacei DNA was detected in tissue samples of all partsof small and large intestine. Histological examination of thegastrointestinal tract of hedgehogs infected with C. erinaceirevealed the presence of developmental stages attached tothe microvillar border along the first half of small intes-tine (Fig. 5). The presence of developmental stages wassporadic; the ratio of infected to non-infected glands was

approximately 1:100. Developmental stages were absent inthe second half of the small and large intestine. No macro-scopical changes were observed and the surface epitheliawere not destroyed. The lamina propria was slightly

M. Kvác et al. / Veterinary Parasitology 201 (2014) 9–17 13

Fig. 2. Phylogenetic relationships between Cryptosporidium erinacei (highlighted) and other Cryptosporidium spp. as inferred by a neighbor-joining analysisof (A) the SSU (750 base positions in the final dataset), (B) actin (725 base positions in the final dataset), (C) GP60 (637 base positions in the final dataset)g ustered

r Scale ba

es

3

S(tau

p

1h

t

osC

enes. The percentage of replicate trees in which the associated taxa clepresent bootstrap values for the nodes gaining more than 50% support.

dematous and the occasional dilatation of lymphatic ves-els was observed.

.4. Taxonomic summary

C. erinacei n. sp.Diagnosis: Oocysts are shed fully sporulated.

porulated oocysts (n = 100) measure 4.5–5.8mean = 4.9 �m) × 4.0–4.8 �m (mean = 4.4) with a lengtho width ratio of 1.13 (1.02–1.35). Four sporozoitesre present in each oocyst. Endogenous stages arenknown.

Type host: Erinaceus europaeus Linnaeus 1758; Euro-ean hedgehog (Dyachenko et al., 2010).

Other natural hosts: Equus ferus caballus Linnaeus,758; domestic horse and Homo sapiens Linnaeus, 1758;uman (Kvác et al., 2014; Laatamna et al., 2013).

Experimental host: A. albiventris (Wagner, 1841); four-oed hedgehog.

Prepatent period in hedgehog: 4–6 days.Patent period in hedgehog: at least 16 days.Type locality: Brno, Czech RepublicOther localities: Germany, AlgeriaSite of infection: small intestine

Material deposited: A phototype, description of

ocysts, and DNA are deposited at the Institute of Para-itology, Biology Centre of the Academy of Sciences of thezech Republic.

together in the bootstrap test (1000 replicates). Numbers at the nodesr included in each tree.

DNA sequences: Partial sequences of SSU, actin, COWP,gp60, HSP70 and TRAP-C1 genes were submitted to Gen-Bank under the accession numbers KF612324-KF612329.

Etymology: The species name erinacei is derived fromthe Latin noun “erinaceus” (meaning a hedgehog) accord-ing to ICZN Article 11.9.1–3 as a singular in the genitivecase, as it appears to be adapted to hedgehogs.

4. Discussion

The specificity of C. erinacei for hedgehogs under naturaland experimental conditions distinguishes it from phy-logentically closely related species, including C. parvum(relatively broad specificity), C. tyzzeri (house mouse spe-cific), C. hominis (human specific), and Cryptosporidiumcuniculus (rabbit specific). With the exception of a singlereport in a horse, natural C. erinacei infections have beenidentified only in hedgehogs. The narrow host specificityof C. erinacei also is evident from experimental transmis-sion studies, which show that mice (immunocompetentand immunocompromised), gerbils, and hamsters are notsusceptible to infection. In contrast, under experimentalconditions, gerbils are susceptible to C. parvum, C. hominisand C. cuniculus, and adult mice are susceptible to C. tyzzeri,

C. hominis and C. cuniculus (Kvác et al., 2013b; Robinsonet al., 2010).

The prepatent period of C. erinacei in hedgehogs is sim-ilar to that of C. parvum in calves (2–7 days), C. tyzzeri in

14 M. Kvác et al. / Veterinary Parasitology 201 (2014) 9–17

Fig. 3. Phylogenetic relationships between Cryptosporidium erinacei (highlighted) and other Cryptosporidium spp. as inferred by a neighbor-joining analysisof (A) COWP (384 base positions in the final dataset), (B) HSP70 (277 base positions in the final dataset) and (C) TRAP-C1 (523 base positions in the finaldataset). The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates). Numbers at the nodesrepresent bootstrap values for the nodes gaining more than 50% support. Scale bar included in each tree. Interrupted branches have been shortened ten-fold.

M. Kvác et al. / Veterinary Parasitology 201 (2014) 9–17 15

F d four-te

aleCpdipR

F(

ig. 4. Course of infection of Cryptosporidium erinacei in three 8-week-olxamination of feces.

dult immunocompetent mice (4–7 days), and C. cunicu-us in rabbits (4–7 days) (Kvác et al., 2013b; Robinsont al., 2010; Tzipori, 1983). Similar to other host-adaptedryptosporidium spp., such as C. suis and C. scrofarum inigs, C. tyzzeri in mice, and C. ryanae in cattle, C. erinacei

oes not appear to cause clinical disease in hedgehogs, and

nfections are characterized by low oocyst shedding for arolonged period (Fayer et al., 2008; Kvác et al., 2013b;en et al., 2012). Horses infected by C. erinacei also had

ig. 5. Cryptosporidium life cycle stages (arrows) in mucosal glandular epitheliumAtelerix albiventris) with dose 1,000,000 oocysts of Cryptosporidium erinacei. Bar

oed hedgehogs (Atelerix albiventris) based on molecular and coprological

no clinical symptoms (Laatamna et al., 2013). Dyachenkoet al. (2010) reported that ∼30% of European hedgehogsco-infected with C. erinacei and C. parvum had diarrhea,suggesting that C. erinacei may, under certain conditions,be pathogenic for hedgehogs. It is equally plausible, how-

ever, that the diarrheal outcome was caused by C. parvumalone or that the co-infection was necessary for the diseaseoutcome. In other reports of clinical cryptosporidiosis inhedgehogs, including fatal infections in juvenile hedgehog

from the small intestine of experimentally infected four-toed hedgehogs25 �m.

y Parasi

16 M. Kvác et al. / Veterinar

(A. albiventris) housed at the Baltimore Zoo (Graczyk et al.,1998) and hemorrhagic diarrhea in a European hedgehog(Meredith and Milne, 2009), the species/genotype causingthe disease was not identified.

C. erinacei oocysts were morphometrically indistin-guishable from those of C. parvum (5.3 �m × 4.7 �m),C. scrofarum (5.16 �m × 4.83 �m), and C. ubiquitum(5.04 �m × 4.66 �m) (this study; Fayer et al., 2010; Kvácet al., 2013a). Although the mean shape index of C. erina-cei oocysts was similar to that of C. parvum, some C. erinaceioocysts appeared more oval shaped (middle panel of Fig. 1Band first panel of Fig. 1C). Oocysts of C. erinacei were smallerthan C. suis (6.2 �m × 5.5 �m) and larger than C. ryanae(3.16 �m × 3.73 �m) and C. xiaoi (3.94 �m × 3.44 �m) (Fallet al., 2003; Fayer and Santín, 2009; Fayer et al., 2008, 2010;Kvác et al., 2013a; Vítovec et al., 2006).

Phylogenetic analyses based on SSU, actin, COWP, TRAP-C1 and HSP70 gene sequences showed that C. erinacei isgenetically distinct from known species and is most closelyrelated to C. parvum, C. hominis, C. cuniculus and C. tyzzeri.At the SSU locus, C. erinacei is 99.5%, 99.1%, 98.6% and99.0%, similar to C. parvum, C. hominis, C. cuniculus and C.tyzzeri, respectively. This is comparable to the similaritiesbetween Cryptosporidium andersoni and Cryptosporidiummuris (99.1%), C. hominis and C. cuniculus (98.9%), and C.bovis and C. xiaoi (99.5%). At the actin locus, C. erinaceiwas 99.5%, 98.6%, 98.7% and 99.1% similar to C. parvum,C. hominis, C. cuniculus and C. tyzzeri, respectively. Thesesimilarities are higher than the 96.5% similarity between C.andersoni and C. muris and are comparable to the similaritybetween C. cuniculus and C. hominis (99.8%). At the gp60locus, C. erinacei forms a group that is closely related to C.cuniculus family Vb. Cryptosporidium erinacei and C. parvumare identical at the COWP gene locus examined. This issimilar to the 100% identity shared by C. cuniculus and C.hominis at this locus (Robinson et al., 2010). At the HSP70locus examined, C. erinacei was 99.2% similar to C. parvum,which is less than the 99.6% similarity shared between C.cuniculus and C. hominis, and greater than the 97.8% sim-ilarity between C. andersoni and C. muris. At the TRAP-C1locus, C. erinacei was 99.8%, 98.4%, and 98.2% similar to C.parvum, C. hominis, and C. tyzzeri, respectively.

In conclusion, morphological, genetic, and biologicaldata support the establishment of Cryptosporidium hedge-hog genotype as a new species. According to ICZN andcriteria for naming Cryptosporidium species (Xiao et al.,2004) we propose the name C. erinacei.

Acknowledgements

The authors would like to thank Cathy Giddings andMichaela Kotková for expert technical assistance. Thisstudy was funded by projects from the Ministry of Educa-tion, Youth and Sports of the Czech Republic (LH11061), theGrant Agency of University of South Bohemia (011/2013/Z).

References

Alves, M., Xiao, L.H., Sulaiman, I., Lal, A.A., Matos, O., Antunes, F., 2003.Subgenotype analysis of Cryptosporidium isolates from humans, cattle,and zoo ruminants in Portugal. J. Clin. Microbiol. 41, 2744–2747.

tology 201 (2014) 9–17

Arrowood, M.J., Sterling, C.R., 1987. Isolation of Cryptosporidium oocystsand sporozoites using discontinuous sucrose and isopycnic Percollgradients. J. Parasitol. 73, 314–319.

Dyachenko, V., Kuhnert, Y., Schmaeschke, R., Etzold, M., Pantchev,N., Daugschies, A., 2010. Occurrence and molecular character-ization of Cryptosporidium spp. genotypes in European hedge-hogs (Erinaceus europaeus L.) in Germany. Parasitology 137,205–216.

Enemark, H.L., Ahrens, P., Juel, C.D., Petersen, E., Petersen, R.F.,Andersen, J.S., Lind, P., Thamsborg, S.M., 2002. Molecular character-ization of Danish Cryptosporidium parvum isolates. Parasitology 125,331–341.

Fall, A., Thompson, R.C., Hobbs, R.P., Morgan-Ryan, U., 2003. Morphologyis not a reliable tool for delineating species within Cryptosporidium. J.Parasitol. 89, 399–402.

Fayer, R., Santín, M., 2009. Cryptosporidium xiaoi n. sp. (Apicomplexa: Cryp-tosporidiidae) in sheep (Ovis aries). Vet. Parasitol. 164, 192–200.

Fayer, R., Santín, M., Macarisin, D., 2010. Cryptosporidium ubiquitum n. sp.in animals and humans. Vet. Parasitol. 172, 23–32.

Fayer, R., Santin, M., Trout, J.M., 2008. Cryptosporidium ryanae n. sp. (Api-complexa: Cryptosporidiidae) in cattle (Bos taurus). Vet. Parasitol.156, 191–198.

Graczyk, T.K., Cranfield, M.R., Dunning, C., Strandberg, J.D., 1998. Fatalcryptosporidiosis in a juvenile captive African Hedgehog (Ateletrixalbiventris). J. Parasitol. 84, 178–180.

Jiang, J., Alderisio, K.A., Xiao, L., 2005. Distribution of Cryptosporidiumgenotypes in storm event water samples from three watersheds inNew York. Appl. Environ. Microbiol. 71, 4446–4454.

Kilani, R.T., Sekla, L., 1987. Purification of Cryptosporidium oocysts andsporozoites by cesium chloride and Percoll gradients. Am. J. Trop. Med.Hyg. 36, 505–508.

Kvác, M., Kestránová, M., Pinková, M., Kvetonová, D., Kalinová, J., Wag-nerová, P., Kotková, M., Vítovec, J., Ditrich, O., McEvoy, J., Stenger,B., Sak, B., 2013a. Cryptosporidium scrofarum n. sp. (Apicomplexa:Cryptosporidiidae) in domestic pigs (Sus scrofa). Vet. Parasitol. 191,218–227.

Kvác, M., McEvoy, J., Loudová, M., Stenger, B., Sak, B., Kvetonová, D., Ditrich,O., Rasková, V., Moriarty, E., Rost, M., Macholán, M., Piálek, J., 2013b.Coevolution of Cryptosporidium tyzzeri and the house mouse (Musmusculus). Int. J. Parasitol. 43, 805–817.

Kvác, M., Ondrácková, Z., Kvetonová, D., Sak, B., Vítovec, J., 2007. Infectiv-ity and pathogenicity of Cryptosporidium andersoni to a novel host,southern multimammate mouse (Mastomys coucha). Vet. Parasitol.143, 229–233.

Kvác, M., Saková, K., Kvetonová, D., Kicia, M., Wesolowska, M., McEvoy, J.,Sak, B., 2014. Gastroenteritis caused by the Cryptosporidium hedge-hog genotype in an immunocompetent man. J. Clin. Microbiol. 52,347–349.

Kvác, M., Vítovec, J., 2003. Prevalence and pathogenicity of Cryptosporid-ium andersoni in one herd of beef cattle. J. Vet. Med. B Infect. Dis. Vet.Public Health 50, 451–457.

Laatamna, A.E., Wagnerová, P., Sak, B., Kvetonová, D., Aissi, M., Rost,M., Kvác, M., 2013. Equine cryptosporidial infection associatedwith Cryptosporidium hedgehog genotype in Algeria. Vet. Parasitol.197, 1–6.

Lv, C., Zhang, L., Wang, R., Jian, F., Zhang, S., Ning, C., Wang, H., Feng,C., Wang, X., Ren, X., Qi, M., Xiao, L., 2009. Cryptosporidium spp. inwild, laboratory, and pet rodents in China: prevalence and molecularcharacterization. Appl. Environ. Microbiol. 75, 7692–7699.

Meredith, A.L., Milne, E.M., 2009. Cryptosporidial infection in a cap-tive European hedgehog (Erinaceus europaeus). J. Zoo Wildl. Med. 40,809–811.

Milácek, P., Vítovec, J., 1985. Differential staining of cryptosporidia byaniline-carbol-methyl violet and tartrazine in smears from feces andscrapings of intestinal mucosa. Folia Parasitol. (Praha) 32, 50.

Pantchev, N., Möller, C., 2007. Successful treatment of cryptosporidiosisin a European hedgehog (Erinaceus europaeus) with paromomycinsulfate (Humatin (R)) – a case report and review of the literature.Kleintierpraxis 52, 368–373.

Pedraza-Diaz, S., Amar, C., Nichols, G.L., McLauchlin, J., 2001. Nested poly-merase chain reaction for amplification of the Cryptosporidium oocystwall protein gene. Emerg. Infect. Dis. 7, 49–56.

Ren, X., Zhao, J., Zhang, L., Ning, C., Jian, F., Wang, R., Lv, C., Wang, Q.,Arrowood, M.J., Xiao, L., 2012. Cryptosporidium tyzzeri n. sp. (Api-

complexa: Cryptosporidiidae) in domestic mice (Mus musculus). Exp.Parasitol. 130, 274–281.

Robinson, G., Wright, S., Elwin, K., Hadfield, S.J., Katzer, F., Bartley,P.M., Hunter, P.R., Nath, M., Innes, E.A., Chalmers, R.M., 2010. Re-description of Cryptosporidium cuniculus Inman and Takeuchi, 1979

y Parasi

R

S

S

S

S

S

M. Kvác et al. / Veterinar

(Apicomplexa: Cryptosporidiidae): morphology, biology and phy-logeny. Int. J. Parasitol. 40, 1539–1548.

yan, U., Power, M., 2012. Cryptosporidium species in Australian wildlifeand domestic animals. Parasitology 139, 1673–1688.

auch, J.F., Flanigan, D., Galvin, M.L., Berman, D., Jakubowski, W., 1991. Pro-pidium iodide as an indicator of Giardia cyst viability. Appl. Environ.Microbiol. 57, 3243–3247.

pano, F., Putignani, L., McLauchlin, J., Casemore, D.P., Crisanti, A., 1997.PCR-RFLP analysis of the Cryptosporidium oocyst wall protein (COWP)gene discriminates between C. wrairi and C. parvum, and between C.parvum isolates of human and animal origin. FEMS Microbiol. Lett.150, 209–217.

pano, F., Putignani, L., Naitza, S., Puri, C., Wright, S., Crisanti, A., 1998.Molecular cloning and expression analysis of a Cryptosporidiumparvum gene encoding a new member of the thrombospondin family.Mol. Biochem. Parasitol. 92, 147–162.

turdee, A.P., Chalmers, R.M., Bull, S.A., 1999. Detection of Cryptosporid-ium oocysts in wild mammals of mainland Britain. Vet. Parasitol. 80,

273–280.

ulaiman, I.M., Hira, P.R., Zhou, L., Al-Ali, F.M., Al-Shelahi, F.A., Shweiki,H.M., Iqbal, J., Khalid, N., Xiao, L., 2005. Unique endemicity ofcryptosporidiosis in children in Kuwait. J. Clin. Microbiol. 43,2805–2809.

tology 201 (2014) 9–17 17

Sulaiman, I.M., Lal, A.A., Xiao, L., 2002. Molecular phylogeny and evolu-tionary relationships of Cryptosporidium parasites at the actin locus. J.Parasitol. 88, 388–394.

Sulaiman, I.M., Morgan, U.M., Thompson, R.C., Lal, A.A., Xiao, L., 2000.Phylogenetic relationships of Cryptosporidium parasites based on the70-kilodalton heat shock protein (HSP70) gene. Appl. Environ. Micro-biol. 66, 2385–2391.

Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S., 2011.MEGA5: molecular evolutionary genetics analysis using maximumlikelihood, evolutionary distance, and maximum parsimony methods.Mol. Biol. Evol. 28, 2731–2739.

Tzipori, S., 1983. Cryptosporidiosis in animals and humans. Microbiol. Rev.47, 84–96.

Vítovec, J., Hamadejová, K., Landová, L., Kvác, M., Kvetonová, D., Sak, B.,2006. Prevalence and pathogenicity of Cryptosporidium suis in pre-and post-weaned pigs. J. Vet. Med. B 53, 239–243.

Xiao, L., Fayer, R., Ryan, U., Upton, S.J., 2004. Cryptosporidium taxonomy:recent advances and implications for public health. Clin. Microbiol.

Rev. 17, 72–97.

Xiao, L., Morgan, U.M., Limor, J., Escalante, A., Arrowood, M., Shulaw, W.,Thompson, R.C., Fayer, R., Lal, A.A., 1999. Genetic diversity withinCryptosporidium parvum and related Cryptosporidium species. Appl.Environ. Microbiol. 65, 3386–3391.