Cryptosporidiosis and Cryptosporidium species in animals and humans: A thirty colour rainbow?

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Contents lists available at ScienceDirect

International Journal for Parasitology journal homepage: www.elsevier.com/locate/ijpara

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Invited Review

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Cryptosporidiosis and Cryptosporidium species in animals and humans: A thirty colour rainbow?

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Jan Šlapeta ⇑ Faculty of Veterinary Science, McMaster Building B14, University of Sydney, New South Wales 2006, Australia

a r t i c l e

i n f o

Article history: Received 25 June 2013 Received in revised form 29 July 2013 Accepted 31 July 2013 Available online xxxx Keywords: Cryptosporidium Species names Public health ssrRNA Barcode Emerging infectious disease

a b s t r a c t Parasites of the genus Cryptosporidium (Apicomplexa) cause cryptosporidiosis in humans and animals worldwide. The species names used for Cryptosporidium spp. are confusing for parasitologists and even more so for non-specialists. Here, 30 named species of the genus Cryptosporidium are reviewed and proposed as valid. Molecular and experimental evidence suggests that humans and cattle are the hosts for 14 and 13 out of 30 named species, respectively. Two, four and eight named species are considered of major, moderate and minor public health significance, respectively. There are at least nine named species that are shared between humans and cattle. The aim of this review is to outline available species information together with the most commonly used genetic markers enabling the identification of named Cryptosporidium spp. Currently, 28 of 30 named species can be identified using the complete or partial ssrRNA, serving as a retrospective ‘barcode’. Currently, the ssrRNA satisfies the implicit assumption that the reference databases used for comparison are sufficiently complete and applicable across the whole genus. However, due to unreliable annotation in public DNA repositories, the reference nucleotide entries and alignment of named Cryptosporidium spp. has been compiled. Despite its known limitations, ssrRNA remains the optimal marker for species identification. Ó 2013 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved.

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1. Introduction

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Cryptosporidium spp. cause significant diarrhoeal disease in humans and animals worldwide (Bouzid et al., 2013). The genus Cryptosporidium belongs to the phylum Apicomplexa, which includes other major pathogens of medical and veterinary importance such as Plasmodium spp. causing malaria, Toxoplasma gondii causing toxoplasmosis and Eimeria spp. causing coccidiosis. The intracellular parasites of the genus Cryptosporidium infect mammals, birds, reptiles and amphibians (Santín, 2013). Cryptosporidiosis is commonly a self-limiting disease in healthy hosts but represents a life-threatening disease in immuno-compromised and young individuals, for which there is no effective treatment (Bouzid et al., 2013). Morphologically indistinguishable isolates of Cryptosporidium spp. are rather heterogeneous in their DNA sequences, and several of those that initially acquired a ‘genotype’ status were later recognised as distinct species (Plutzer and Karanis, 2009; Xiao, 2010; Chalmers and Katzer, 2013). The nomenclature used for Cryptosporidium spp. is complex and is confusing for parasitologists and even more so for non-specialists. Once a species name is introduced and is linked to a material, it becomes the international

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standard of reference (International Commission on Zoological Nomenclature, 1999). Because it is an international standard a set of rules is in place to maintain continuity and stability of the names. On the other hand, for ‘genotypes’ there are no rules for using, renaming or introducing new examples, hence allowing constant flux. The use of stable nomenclature is needed to marry current and future research with the original published information on Cryptosporidium spp. Therefore, the aim of this review is to outline named species together with the most commonly used genetic marker enabling Cryptosporidium spp. identification. This review does not cover yet unnamed species, ‘genotypes’ and other variants and subtypes. Representative DNA entries have been compiled to enable species identification serving as a reference standard – ‘barcode’ – supplementing species descriptions.

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2. The named species are only some colours of the rainbow: How many is enough?

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In the last decade, the number of named species has grown steadily, with approximately one new named species per year, and 10 named species for 2004–2013. Currently, the total tally is 30 valid, named species (Table 1). This number appears to be a fair coverage of Cryptosporidium spp. from a medical and veterinary perspective. The species that have been named and recognised

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Table 1 Current summary of Cryptosporidium spp. Species Number

Valid species name

Public health significance

Host range

Human

Species I

C. muris Tyzzer, 1907

Minor

MB

Yes

Species II

C. parvum Tyzzer, 1912

Minor

M

(Yes)

Species III

C. meleagridis Slavin, 1955 (syn. C. tyzzeri Levine, 1961) C. wrairi Vetterling, Jervis, Merrill, Sprinz, 1971 C. agni Barker & Carbonell, 1974 (syn. C. xiaoi Fayer & Santín, 2009) C. bovis Barker & Carbonell, 1974

Moderate

MB

Yes

None None

M M

(Yes)

None

M

Yes

C. cuniculus Inman & Takeuchi, 1979 C. felis Iseki, 1979

Moderate Moderate

M M

C. serpentis Levine, 1980 C. nasoris Hoover, Hoerr, Carlton, Hinsman & Ferguson, 1981 C. baileyi Current, Upton & Haynes, 1986 C. varanii Pavlásek, Lávicˇková, Horák, Král & Král, 1995 (syn. C. saurophilum Koudela & Modry´, 1998) C. cichlidis (Paperna & Vilenkin, 1996)

None None None None

RM F B R

None

F

C. reichenbachklinkei (Paperna & Vilenkin, 1996)

None

F

C. galli Pavlásek, 1999

None

B

C. andersoni Lindsay, Upton, Owens, Morgan, Mead, & Blagburn, 2000

Minor

M

Yes

Yes

C. canis Fayer, Trout, Xiao, Morgan, Lal & Dubey, 2001

Minor

M

Yes

(Yes)

C. hominis Morgan-Ryan, Fall, Ward, Hijjawi, Sulaiman, Fayer, Thompson, Olson, Lal & Xiao, 2002 C. molnari Alvarez-Pellitero & Sitjà-Bobadilla, 2002

Major

M

Yes

Yes

Human (I) genotype

None

F

C. suis Ryan, Monis, Enemark, Sulaiman, Samarasinghe, Read, Buddle, Robertson, Zhou, Thompson & Xiao, 2004 C. scophthalmi Alvarez-Pellitero, Quiroga, Sitjà-Bobadilla, Redondo, Palenzuela, Padrós, Vázquez & Nieto, 2004 C. pestis Šlapeta, 2006

Minor

M

(Yes)

Yes

Pig genotype II

None

F

Major

M

Yes

C. fayeri Ryan, Power & Xiao, 2008

Minor

M

(Yes)

C. ryanae Fayer & Santín, Trout, 2008

None

M

C. fragile Jirku˚, Valigurová, Koudela, Krˇízˇek, Modry´ & Šlapeta, 2008

None

A

C. macropodum Power & Ryan, 2008

None

M

C. ducismarci Traversa, 2010

None

R

C. ubiquitum Fayer, Santín & Macarisin, 2010

Minor

M

Yes

C. viatorum Elwin, Hadfield, Robinson, Crouch & Chalmers, 2012

Moderate

M

Yes

C. scrofarum Kvácˇ, Kestrˇánová, Pinková, Kveˇtonˇová, Kalinová, Wagnerová, Kotková, Vítovec, Ditrich, McEvoy, Stenger & Sak, 2013

Minor

M

(Yes)

Species IV Species V Species VI Species VII Species VIII Species IX Species X Species XI Species XII Species XIII Species XIV Species XV Species XVI Species XVII Species XVIII Species XIX Species XX Species XXI Species XXII Species XXIII Species XXIV Species XXV Species XXVI Species XXVII Species XXVIII Species XXIX Species XXX

Yes Yes

Cattle

Genotype designation C. muris B genotype Mouse I genotype

(Yes)

(Yes)

C. bovis-like genotype Bovine B genotype Rabbit genotype Cat genotype

(Yes) n/a Desert monitor genotype Piscine genotype 1 Piscine genotype 2 Finch genotype C. muris A genotype Dog genotype

n/a Yes

Yes

Bovine (II) genotype Marsupial genotype I Deer-like genotype

Marsupial genotype II

Yes

Deer genotype

(Yes)

Pig genotype II

Host range: M, mammal; B, bird; R, reptile; F, fish. n/a, not applicable because the species has not been characterised using any DNA signature. (Yes), indicates extremely rare or experimental evidence.

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are the outcome of molecular surveys and experimental studies worldwide. However, in between the named species exists a great diversity of forms or genotypes that may or may not deserve specific status. The validity of any species is based on a testable species concept that is applicable across the genus (Nadler and Leòn, 2011). The 10 species of Cryptosporidium named in the last decade were proposed as distinct based on their biological and genetic characteristics (Alvarez-Pellitero et al., 2004; Ryan et al., 2004b, 2008; Fayer et al., 2008, 2010; Elwin et al., 2012b; Kvácˇ et al., 2013). The principal parasitic attribute – host specificity – is universally applied across Cryptosporidium spp., but its reliability is

ambiguous (Table 1). In fact, at least five of the 10 most recently named species have been detected in humans. Similarly, five of the 10 were shown to be experimentally infective for cattle. Not all are frequent parasites of either humans or cattle; rather the opposite. Recognition of common as opposed to rare events is only possible for these two hosts due to the enormous efforts directed towards this dataset, representing thousands of isolates analysed globally (Santín et al., 2004, 2008; Fayer et al., 2006b; Elwin et al., 2012a). The experimental evidence needs to be interpreted with caution because some experiments may represent unnatural contexts (Poulin and Keeney, 2008). Nevertheless, apparent parasite errors demonstrate the plasticity of these species and

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are recognised in an ecological concept known as ‘‘ecological fitting’’ (Agosta et al., 2010). The concept suggests that these known and experimental events or ‘fits’ are recent adaptations enabling the formation of species communities under changing environments (Agosta et al., 2010). These ‘fits’ are enabled opportunistically by the changing environment, e.g. proximity to a parasite reservoir. Otherwise specialist species with a relatively narrow host range such as Cryptosporidium scrofarum, Cryptosporidium suis, Cryptosporidium fayeri, Cryptosporidium cuniculus and Cryptosporidium parvum are found in humans, a host species with which they have no previous history of association (Xiao et al., 2002; Kvácˇ et al., 2009; Waldron et al., 2010; Rašková et al., 2013). As opportunities for these events increase, the occurrence of an emerging infectious disease increases as well. An example of such is an outbreak of human cryptosporidiosis caused by C. cuniculius (Chalmers et al., 2009b). The origin of the C. cuniculus outbreak was resolved with relative ease, because rabbits are a reservoir of C. cuniculus (Robinson et al., 2010). A recently described species, Cryptosporidium viatorum, from humans most probably represents another such event but a reservoir is yet to be documented (Elwin et al., 2012b). Cataloguing and phylogenetic analyses of the genus Cryptosporidium, together with inference from host–parasite associations are valid approaches. The use of stable nomenclature is fundamental in pinpointing the origin of an emerging infectious disease. Therefore, there is no definitive number of species names needed. The names are practical labels that enable scientific communication, e.g. C. viatorum, in the quest to identify its reservoir and epidemiology. More species will be named as the need arises and more information is gathered (i.e. chipmunk I, horse, monkey and skunk genotypes have been anecdotally reported in humans). Undeniably, DNA markers are at the forefront of Cryptosporidium spp. surveys. The ssrRNA has been the most frequently used genetic marker for Cryptosporidium. Currently, 28 of 30 named species can be identified using the complete or partial ssrRNA, serving as a retrospective ‘barcode’ that allows phylogenetic reconstruction of the entire genus Cryptosporidium (Fig. 1). However, there is no straightforward way to upload a reference dataset for Cryptosporidium spp. from primary DNA databases (GenBank: www.ncbi.nlm.nih.gov/genbank, EMBL: www.ebi.ac.uk/ena and DDBJ: www.ddbj.nig.ac.jp). In fact, the taxonomy for Cryptosporidium spp. in primary DNA databases is unreliable, implying that BLAST searches against such databases will be misleading. Therefore a reference list of nucleotide entries and the alignment of named Cryptosporidium spp. has been compiled (Supplementary data S1). On the other hand, there are known limitations to single locus typing (e.g. ssrRNA). In particular, ssrRNA is a multicopy gene that does not allow subtype identification and population analyses (Widmer and Sullivan, 2012; Chalmers and Katzer, 2013). The phylogenetic tree based on ssrRNA is poorly resolved in several branches (Fig. 1). Inclusion of actin and HSP70 gene sequences to reconstruct the phylogeny provides a strong phylogenetic signal for Cryptosporidium spp. (Morrison, 2006). Nucleotide entries to actin, HSP70 and COWP1 for each species are included in the following section to assist in species recognition.

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3. Species synopsis

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3.1. Species l. Cryptosporidium muris Tyzzer, 1907

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Cryptosporidium muris was originally described from the stomachs of laboratory mice (Tyzzer, 1907, 1910). The entire development of the parasite is confined to the epithelial cells of the stomach and the patent period is up to 30 days (Iseki et al., 1989; Chalmers et al., 1997). A prevalence of 6% was demonstrated in mice in the United Kingdom (UK) (Chalmers et al., 1997). An iso-

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late (RN66) from Japan was experimentally infective to rats, mice, guinea pigs, rabbits, dogs and cats (Iseki et al., 1989). In addition, several captive animals have been shown to be hosts of C. muris including camels (Camelus bactrianus), rock hyrax (Procavia capensis) and desert hamsters (Phodopus roborovskii). Variation in the infectivity of laboratory animals with C. muris is isolate dependent (Kvácˇ et al., 2008). Recently, isolates from captive ostriches have been characterised as C. muris, suggesting that the species is a generalist (Wang et al., 2012). The species C. muris was previously recognised as ‘‘C. muris genotype B’’ or ‘‘C. muris mouse genotype’’ based on its unique ssrRNA sequence (Morgan et al., 2000). The ssrRNA sequence is 1,725 bp long (GenBank: AB089284) and identical to the ssrRNA in the genome of RN66 (GenBank: AAZY00000000). In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382350, AF221543, AB089287). The RN66 isolate was originally obtained from a rat (Rattus norvegicus) and serves as a reference isolate for the species (Iseki et al., 1989). On several occasions, HIV-positive human subjects are documented to have shed C. muris in their stool samples (Palmer et al., 2003). This parasite species is of minor public health significance.

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3.2. Species ll. Cryptosporidium parvum Tyzzer, 1912

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The species C. parvum was originally described from laboratory mice (Tyzzer, 1912), and later found in voles and mice worldwide (Morgan et al., 1999b; Bajer et al., 2003). In the literature this species has been designated as the ‘‘mouse’’ [or ‘‘mouse I’’] genotype of the C. parvum species complex. The species readily infects neonatal and adult mice, with a patent period of 24–29 days (Tyzzer, 1912; Ren et al., 2012). The species C. parvum produces heavy infections in adult laboratory mice, but the courses of infection differ significantly between isolates (Bednarska et al., 2003). The development of C. parvum is confined to the villous epithelium of the terminal third of the small intestine (Tyzzer, 1912; Ren et al., 2012). The C. parvum species complex is divided into several species based on marker gene sequences, including ssrRNA. For comparative purposes the C. parvum unique ssrRNA sequence (GenBank: AF112571) identifies the species. The ssrRNA sequence of C. parvum is 1,750 bp long (GenBank: AF112571); the oocysts were isolated from the faeces of a mouse in the United States (USA) (Xiao et al., 1999). In addition, actin, COWP1 and HSP70 sequences are available (GenBank: AF382343, AF266268, AF221530). An unprecedented case report demonstrated the presence of DNA considered to represent C. parvum and Cryptosporidium pestis in a 25-year-old female rodent trapper/keeper, suffering from severe diarrhoea (Rašková et al., 2013). Despite thousands of identified Cryptosporidium spp. isolates worldwide, to date the above is the only record of C. parvum in a human. Therefore C. parvum is rarely, if ever, detected in human subjects. This parasite species is of minor public health significance. A recent attempt to rename the species as ‘‘Cryptosporidium tyzzeri’’ (Ren et al., 2012) is not valid, because (i) the authors worked with C. parvum in its strict sense and (ii) the species name has previously been used in the genus Cryptosporidium (Tyzzer, 1912; Šlapeta, 2012). The use of the proposed species name by Ren et al. (2012) contravenes the rules of the International Code for Zoological Nomenclature (ICZN) and creates an unwanted confusion with Cryptosporidium meleagridis (syn. C. tyzzeri).

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3.3. Species lll. Cryptosporidium meleagridis Slavin, 1955

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The species C. meleagridis was originally described from a wild turkey (Meleagris gallopavo), but was later found to be generalist with a wide host range. Cryptosporidium meleagridis is readily

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FJ262725 C. cuniculus [genotype: rabbit]

80

Cryptosporidium parvum species complex

56 60 69

Species XVIII

AF108864 C. pestis [genotype: II bovine]

Species XXII

AF112571 C. parvum [genotype: mouse I]

Species II

AF115378 C. wrairi

Species IV

AF112574 C. meleagridis (syn. C. tyzzeri) AF112570 C. fayeri [genotype: marsupial I] JN846705 C. viatorum

AF513227 C. macropodum [genotype: marsupial II]

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AF115377 C. suis [genotype: pig I] HM209366 C. ubiquitum [genotype: deer]

Public health significance:

AF108862 C. felis [genotype: cat]

minor

AF112573 C. varanii (syn. C. saurophilum) [genotype: desert monitor]

moderate

56

EF547155 C. ducismarci

major

JX424840 C. scrofarum [genotype: pig II] 98

EU410344 C. ryanae [genotype: deer-like]

94 82

FJ896050 C. agni (syn. C. xioai) [genotype: C. bovis-like] 92 AY741305 C. bovis [genotype: bovine B]

100

AF151376 C. serpentis

76 94

Species XXVIII Species VIII Species XII Species XXVII Species XXX Species XXIV Species V Species VI

Species XXV Species XV Species I Species XVI

HM243547 C. molnari

?

Species XX

AF093496 C. andersoni [genotype: A]

Species XIX

FJ769050 C. reichenbachklinkei [genotype: piscine II]

0.02

Species XXVI

AB089284 C. muris [genotype: B]

AY524773 C. cichlidis [genotype: piscine I]

ssrRNA

Species XVII

gastric

AF316624 AY168847 C. galli 79

Species XXIX

Species IX EU162751 C. fragile

74

Species XXIII

Species XI

L19068 C. baileyi

99

Species III

intestinal

AF112576 C. canis [genotype: dog]

78

Species VII

AF108865 C. hominis [genotype: I human]

Species XIII Species XIV

C. nasoris

Species X

C. scophthalmi

Species XXI

Fig. 1. Phylogeny of 28 named Cryptosporidium spp. using a reference dataset of ssrRNA sequences. The reference multiple sequence alignment consists of 1795 positions (Supplementary data S1). A Minimum Evolution tree was reconstructed with a bootstrap test (1,000 replicates). All ambiguous positions were removed for each sequence pair. The branch lengths represent evolutionary distances computed using the Maximum Composite Likelihood method in MEGA5 (Tamura et al., 2011). GenBank accession numbers accompany all species names, as well as public health significance and either intestinal or gastric development within its host. The species parasitising fish served as an outgroup (dotted line). There is no ssrRNA sequence available for Cryptosporidium nasoris (Species X) and Cryptosporidium scophthalmi (Species XXI).

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transmissible between and among mammalian and avian animal hosts (Slavin, 1955; Pedraza-Díaz et al., 2001; Akiyoshi et al., 2003; Chappell et al., 2011; Silverlås et al., 2012). The development is confined to the villous epithelium of the terminal third of the small intestine (Slavin, 1955). In experimentally infected chickens, the development is confined to the lower part of the small intestine and caeca (Akiyoshi et al., 2003). The caecal location confirms the conspecificity of C. meleagridis with the Cryptosporidium sp. reported by Tyzzer in 1929 and later in 1961 named C. tyzzeri. The name C. tyzzeri is therefore a junior synonym of C. meleagridis (Šlapeta, 2012). Experimentally infected animals with human-derived C. meleagridis (TU1867) remained asymptomatic, with the exception of gnotobiotic piglets suffering from diarrhoea (Akiyoshi et al., 2003). Immunosuppressed mice, young chicks and turkey poults, as well as colostrum-fed calves were experimentally susceptible and excreted oocysts (Akiyoshi et al., 2003). Immunocompetent, healthy human subjects were experimentally susceptible to C. meleagridis (TU1867) infection (Chappell et al., 2011). The symptoms of mild self-limiting diarrhoea in human subjects is unlikely

to lead to the need to seek medical attention (Chappell et al., 2011). The patent period in gnotobiotic pigs is 10–12 days, in healthy humans 3–4 days and in chicken 12–16 days (Sréter et al., 2000; Akiyoshi et al., 2003; Chappell et al., 2011). Cryptosporidium meleagridis is identified by its unique ssrRNA sequence (Xiao et al., 1999). The ssrRNA sequence is 1,744 bp long (GenBank: AF112574) obtained from oocysts isolated from the faeces of a turkey in the USA (Morgan et al., 2000). In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382351, AF221537, AF266266). The species C. meleagridis is the third most commonly detected species in humans (Elwin et al., 2012a). Hen–human transmission was suspected on an organic farm, with at least one symptomatic subject confirmed to be shedding C. meleagridis (Silverlås et al., 2012). During routine typing of 14,469 human cryptosporidiosis cases in the UK, 109 were C. meleagridis infections (Elwin et al., 2012a). A significant risk factor for C. meleagridis infection was travel abroad (Elwin et al., 2012a). This parasite species is of moderate public health significance, due to its zoonotic and anthroponotic spread as well as its documented high infectivity rate.

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3.4. Species lV. Cryptosporidium wrairi Vetterling, Jervis, Merrill & Sprinz, 1971

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The species C. wrairi is known only from laboratory animal facilities, where it has caused a spontaneous outbreak of cryptosporidiosis in laboratory guinea pigs (Cavia porcellus). Its identity in natural environments and in its natural host is unknown (Vetterling et al., 1971). Cryptosporidium wrairi appears to be specialised to laboratory reared guinea pigs; nevertheless some degree of infectivity to laboratory suckling mice, lambs and calves has been documented for some isolates (Vetterling et al., 1971; Angus et al., 1985; Chrisp et al., 1992). It develops in the brush border of the small intestine. The patent period is 2–3 weeks. Cryptosporidium wrairi is identified by its unique ssrRNA sequence. The ssrRNA sequence is 1,746 bp long (GenBank: AF115378), obtained from a guinea pig in the USA (Xiao et al., 1999). In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382348, AF221536, U35027). This parasite species has no zoonotic potential and therefore is of no public health significance.

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3.5. Species V. Cryptosporidium agni Barker & Carbonell, 1974

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The species C. agni infects sheep (Ovis aries), more commonly pre-weaned lambs than post-weaned lambs, worldwide. In this review it is considered conspecific with Cryptosporidium xiaoi (Fayer and Santín, 2009), therefore C. xiaoi is a junior synonym of C. agni (Barker and Carbonell, 1974). Experimental infections using oocysts from sheep led to infection in lambs that shed oocysts for 14–15 days, however an attempt to infect calves and goats was unsuccessful (Fayer and Santín, 2009). The host range is poorly understood and cross-transmission between ruminants (cattle, sheep and goats) cannot be excluded. In some publications, the species is referred as the ‘‘Cryptosporidium bovis-like genotype’’ and its genetic signature has been identified in faeces from goats and yaks (Karanis et al., 2007; Fayer and Santín, 2009). Moreover, there are reports indicating that C. bovis in sheep should be considered C. agni (Mueller-Doblies et al., 2008; Yang et al., 2009). Cryptosporidium agni has been considered to contribute to neonatal diarrhoea in kids (Díaz et al., 2010; Rieux et al., 2013). The species was redescribed using oocysts from faecal material from sheep in the USA (Fayer and Santín, 2009). The species name was originally introduced for a species in the brush border of the small intestine of a sheep in Australia (Barker and Carbonell, 1974). Due to the occurrence of a multitude of Cryptospordium spp. in pre-weaned lambs, the redescription was warranted, but a new name was unnecessary (Fayer and Santín, 2009). Cryptosporidium agni is poorly defined by ssrRNA sequence data, with a major overlap with C. bovis. In this review, a partial ssrRNA sequence 828 bp long (GenBank: FJ896050) is considered to represent C. agni, however heterogeneity has been reported (Elwin and Chalmers, 2008; Fayer and Santín, 2009). In addition, actin and HSP70 sequences are available (GenBank: FJ896042, FJ896041). There is no zoonotic potential. This parasite species is of no public health significance.

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3.6. Species Vl. Cryptosporidium bovis Barker & Carbonell, 1974

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The species C. bovis infects cattle (Bos taurus), commonly postweaned calves, worldwide. The species was previously recognised as the ‘‘Cryptosporidium genotype Bovine B’’ (Santín et al., 2004). In a study in the USA in which calves were monitored over 2 years, the cumulative prevalence of C. bovis reached 80%, and the greatest number of infected calves occurred between 12 weeks and 14 months of age (Santín et al., 2008). In a study from Sweden, C. bovis was the most common species in dairy calves, including

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74% of pre-weaned calves, 77% of 3.5–13 month old calves and 100% of cows (Silverlås et al., 2010b). The clinical significance of C. bovis infection is not documented in controlled experimental trials. Infection with this species is most likely to be asymptomatic (Silverlås et al., 2010a,b). In beef cattle, a significantly higher prevalence was observed in 6–8 month old calves (39%) than in cows over 2 years of age (4%) in a study from the USA (Feltus et al., 2008). Oocysts of C. bovis were not infective to neonatal mice and neonatal lambs, although oocysts from the same pool were infective for a calf with a patent period of 18 days (Fayer et al., 2005). The species was redescribed based on oocysts from faecal material from dairy and beef cattle in the USA (Fayer et al., 2005). The species name was originally introduced for a species found in the brush border of the intestine of a calf in Australia (Barker and Carbonell, 1974); due to a large number of Cryptosporidium spp. found in cattle, the redescription was warranted (Fayer et al., 2005). The species C. bovis is defined primarily by its unique ssrRNA sequence; a partial ssrRNA is 770 bp long (GenBank: AY741305). However, its distinction from C. agni is not unambiguous (Elwin and Chalmers, 2008; Fayer and Santín, 2009). In addition, actin and HSP70 sequences are available (GenBank: AY741307, AY741306). This species has no zoonotic potential. It is of no public health significance.

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3.7. Species Vll. Cryptosporidium cuniculus Inman & Takeuchi, 1979

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Cryptosporidium cuniculus is principally found in European rabbits (Oryctolagus cuniculus). The species came to prominence in a waterborne outbreak in the summer of 2008 in the UK (Chalmers et al., 2009b, 2011; Robinson et al., 2010). The species was first reported from apparently healthy laboratory rabbits in the USA (Inman and Takeuchi, 1979; Rehg et al., 1979). It was detected in 7% of faecal samples from asymptomatic wild feral rabbits in Australia and 2% of faecal samples from 4 to 6 month old laboratory rabbits (Nolan et al., 2010; Zhang et al., 2012). Juvenile rabbits, immunosuppressed Mongolian gerbils (Meriones unguiculatus) and immunosuppressed Porton mice are susceptible to experimental infection but neonatal mice are not (Robinson et al., 2010). None of the experimental animals showed clinical signs (Robinson et al., 2010). The development of C. cuniculus is confined to the brush border of small intestinal epithelial cells and the patent period is 7 days (Robinson et al., 2010). The species belongs to the C. parvum species complex, divided into several species based on several gene sequences, including ssrRNA. Cryptosporidium cuniculus is genetically very closely related to Cryptosporidium hominis (Robinson et al., 2010). For comparative purposes the C. cuniculus unique ssrRNA (GenBank: FJ262725) identifies the species; it was temporarily recognised as the ‘‘rabbit genotype’’ (Chalmers et al., 2009b). The partial ssrRNA sequence of C. cuniculus is 801 bp long (GenBank: FJ262725) and was obtained from oocysts isolated from the faeces of a rabbit in the UK (Robinson et al., 2010). Actin, HSP70 and COWP1 sequences are available (GenBank: GU327783, FJ262728, GU327782). In a retrospective study using material from the UK, C. cuniculus was diagnosed in 37 of 3,030 (1.2%) infections with Cryptosporidium spp. during 2007–2008, suggesting a seasonal distribution of infection peaking in late summer to early autumn (Chalmers et al., 2011). Animal-to-person transmission has not been unambiguously documented due to inadequate clinical histories for the human subjects infected with C. cuniculus. Epidemiological studies of relationships with the breeding cycle of rabbits and with the age groups of rabbits are warranted to explain the seasonal distribution of C. cuniculus human infections. Due to a tap-water out-

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break in 2008 in the UK, this parasite species is of moderate public health significance.

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3.8. Species Vlll. Cryptosporidium felis Iseki, 1979

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Natural infection with C. felis is predominantly detected in asymptomatic domestic cats (Felis catus) worldwide (Iseki, 1979; Fayer et al., 2006a; Ballweber et al., 2009). Infection with Giardia sp. was the only significant risk factor for C. felis infection in a study of domestic cats in the USA (Ballweber et al., 2009). The species appears to be restricted to cats, because no other host was experimentally susceptible to C. felis. The endogenous developmental stages were found throughout the small intestine of cats with no developmental stages in the stomach, caecum or colon (Iseki, 1979). An unprecedented case report suggested the shedding of C. felis oocysts by a cow in Poland (Bornay-Llinares et al., 1999). Cryptosporidium felis is recognised as a ‘‘cat-adapted strain’’ or ‘‘cat genotype [type]’’ based on its unique ssrRNA sequence (Pieniazek et al., 1999). The ssrRNA sequence is 1,784 bp long (GenBank: AF108862) and was obtained from oocysts isolated from the faeces of a cat in Australia (Morgan et al., 1999c). In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382347, AF221538, AF266263). Cryptosporidium felis is occasionally detected in humans. During routine typing of 14,469 human cases of cryptosporidiosis in the UK, C. felis ranked the fourth most commonly encountered species (n = 38) (Elwin et al., 2012a). In a study in the UK, significant risk factors were that the subjects were immunocompromised or had contact with cats (Elwin et al., 2012a). Both adults and children are infected with C. felis and both immunocompetent and HIV-positive subjects have been reported to shed C. felis (Pieniazek et al., 1999; Cacciò et al., 2002). Oocyst shedding was documented to last for at least 6 months in one symptomatic HIV-positive subject (Pieniazek et al., 1999). In Peru, a single HIV-negative child without diarrhoea shed C. felis on one occasion (Xiao et al., 2001). This parasite species is of moderate public health significance.

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3.9. Species lX. Cryptosporidium serpentis Levine, 1980

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Cryptosporidium serpentis colonises the stomach mucosa of snakes and lizards. In snakes, C. serpentis causes gastric mucosal hypertrophy and severe haemorrhagic and erosive gastritis. The outcome is a subclinical chronic infection or mortality (Cranfield and Graczyk, 1994; Graczyk and Cranfield, 2000). The most pronounced clinical sign in affected snakes is regurgitation of prey. In lizards it is generally asymptomatic. Infected animals are known to shed oocysts for years (Cranfield and Graczyk, 1994). A diagnosis-euthanasia control strategy is warranted under certain conditions (e.g. captive multi-reptile operation) (Xiao et al., 2004). On a single occasion, C. serpentis was isolated from calves in China and subsequently shown to be infective to mice (Chen and Qiu, 2012). This is the only reptile isolate able to cross the host barriers between a mammal and reptile. Further study is warranted to confirm the finding. Cryptosporidium serpentis was redescribed based on oocyst morphology and later characterised genetically (Upton et al., 1989; Tilley et al., 1990; Xiao et al., 2004). The ssrRNA sequence of C. serpentis is 1,743 bp long (GenBank: AF151376). It was obtained from oocysts isolated from the faeces of a wild caught corn snake (Pantherophis guttatus guttatus) from the USA (Xiao et al., 2004). In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382353, AF221541, AF266275). This parasite species has no zoonotic potential and is of no public health significance.

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3.10. Species X. Cryptosporidium nasoris Hoover et al., 1981

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Cryptosporidium nasoris was described in a case report of an anorectic captive juvenile marine tropical fish, the Lipstick Tang (Naso lituratus) from a pet shop in the USA (Hoover et al., 1981). Histological examination and electron microscopy of intestinal mucosa revealed Cryptosporidium development. However, the location of the sporulating oocysts, deep in the gut epithelium, was not reported or is not identifiable from published figures for C. nasoris (Paperna and Vilenkin, 1996; Alvarez-Pellitero and Sitjà-Bobadilla, 2002). To confirm the identity of this species a new isolate needs to be redescribed and ssrRNA amplified and sequenced to confirm its similarity to available species. This species has no zoonotic potential. It is of no public health significance.

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3.11. Species Xl. Cryptosporidium baileyi Current, Upton & Haynes, 1986

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Cryptosporidium baileyi is considered a generalist amongst birds; it infects a wide variety of wild and pet birds, as well as farmed chickens, turkeys and ducks (Goodwin, 1989; Nakamura et al., 2009; Qi et al., 2011). The prevalence is variable depending on species and locality. Infection can be manifested by clinical signs associated with parasite development in the digestive and respiratory tracts (Goodwin, 1989). In naturally and experimentally infected chickens, all stages of the parasite are found mainly in the microvillous region of epithelial cells of the intestine including the bursa of Fabricius and cloaca. This species is capable of infecting the microvillous region of the epithelial cells lining the sinuses and trachea as well as epithelial cells of the conjunctivae (Current et al., 1986). In chickens the patent period lasts from 4 to 24 days (Current et al., 1986). Cryptosporidium baileyi was described from a domestic chicken (Gallus gallus domesticus) from a commercial broiler flock in the USA (Current et al., 1986). The ssrRNA sequence is 1,733 bp long (GenBank: L19068) and was obtained from oocysts isolated from chicken faeces. In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382346, AJ310880, AF266276). This parasite species has no zoonotic potential and is of no public health significance.

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3.12. Species XII. Cryptosporidium varanii Pavlásek, Lávicˇková, Horák, Král & Král, 1995

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Cryptosporidium varanii (syn. Cryptosporidium saurophilum) has been detected in captive and wild reptiles worldwide and predominantly infects lizards, where development is confined to the intestine (Koudela and Modry´, 1998). Clinical signs, if present, have mainly been described for lizards and include emaciation and anorexia (Terrell et al., 2003). Cryptosporidium varanii was previously recognised as ‘‘Cryptosporidium desert monitor genotype’’ and C. saurophilum (Koudela and Modry´, 1998). Cryptosporidium varanii (syn. C. saurophilum) is now defined by its unique ssrRNA sequence (Xiao et al., 1999; Plutzer and Karanis, 2009; Karanis et al., 2010). The ssrRNA sequence is 1,743 bp long (GenBank: AF112573), and was obtained from oocysts isolated from faeces of a desert monitor (Varanus griseus) imported to the USA (Xiao et al., 2004). In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382349, AF221540, AF266277). This parasite species has no zoonotic potential and is of no public health significance.

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3.13. Species XIII. Cryptosporidium cichlidis (Paperna & Vilenkin, 1996) The species, C. cichlidis, was described from cultured hybrid fry and fingerling tilapia (Oreochromis aureus  Oreochromis niloticus) in fish ponds from Israel. It was reported only in 11–55 mm long tilapia fish, but not detected in fish 56–152 mm in length (Paperna and Vilenkin, 1996). The developmental stages occur in the epithelial cells of the stomach. The species is described based on its gastric location and presumed strict specificity to tilapia. For the purpose of this review a parasite found in the gastric mucosa of a guppy (Poecilia reticulata) in a breeding facility that lost approximately 40 guppies over a period of 3–4 days is considered to be conspecific with C. cichlidis (Ryan et al., 2004a). The parasite development the in gastric mucosa was identified histologically. Therefore, a partial ssrRNA sequence 483 bp long (GenBank: AY524773) defines the species (Ryan et al., 2004a). It is also recognised as the ‘‘piscine genotype 1’’ (Zanguee et al., 2010). This parasite species has no zoonotic potential and is of no public health significance. 3.14. Species XlV. Cryptosporidium reichenbachklinkei (Paperna & Vilenkin, 1996)

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The species C. reichenbachklinkei was described from reared gouramis (Trichogaster leeri) from a commercial fish farm in Israel. The fish originated from an unknown source in southeastern Asia (Paperna and Vilenkin, 1996). The developmental stages occur in the epithelial cells of the stomach. The species was described based on its gastric location and its presumed strict specificity to gouramis. For the purpose of this review a parasite reported to cause clinical gastric cryptosporidiosis in a freshwater angelfish (Pterophyllum scalare) hatchery with variable levels of emaciation, poor growth rates, anorexia and increased mortality is considered conspecific with C. reichenbachklinkei (Murphy et al., 2009). Therefore, a partial ssrRNA sequence 695 bp long (GenBank: FJ769050) defines the species (Murphy et al., 2009). It is also recognised as the ‘‘piscine genotype 2’’ (Zanguee et al., 2010). This parasite species has no zoonotic potential. It is of no public health significance.

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3.15. Species XV. Cryptosporidium galli Pavlásek, 1999

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Cryptosporidium galli is described from a wide range of birds including the domestic chicken (G. g. domesticus) where it infects the epithelial cells of the proventriculus (Ryan et al., 2003b). In some aviaries, this parasite can be quite prevalent especially in song birds (da Silva et al., 2010). The majority of infected birds present no clinical signs. Nevertheless C. galli has been suggested to cause chronic gastritis in canaries, lesser seed-finches and a cockatiel, leading to mortalities (Antunes et al., 2008). Cryptosporidium galli is identified by its unique ssrRNA sequence (Ryan et al., 2003b). For a comparative purpose two partial ssrRNA sequences representing C. galli have been selected (GenBank: AF316624, AY168847), 671 and 537 bp long, respectively. In addition, actin and HSP70 sequences are available (GenBank: AY163901, AY168849). This parasite species has no zoonotic potential. It is of no public health significance.

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3.16. Species XVl. Cryptosporidium andersoni Lindsay, Upton, Owens, Morgan, Mead & Blagburn, 2000 Cryptosporidium andersoni has been detected primarily in domestic cattle (B. taurus), where it infects the epithelial cells of

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the abomasum (Lindsay et al., 2000). The prevalence is variable; one study from the USA demonstrated an overall prevalence in 1–2 year old dairy heifers on 14 farms in seven states of 5% (Trout et al., 2006). In addition, C. andersoni has been detected in captive camels (C. bactrianus) and sporadically in sheep and goats. The species is infective under experimental conditions to gerbils (M. unguiculatus) (Lindsay et al., 2000). Some strains of C. andersoni are infective to mice (Masuno et al., 2013). Natural infection in cattle is asymptomatic (Kvácˇ et al., 2008). However, in one study, cattle infected with C. andersoni had reduced weight gains compared with uninfected cattle (Ralston et al., 2010). A patent period of greater than 2.5 years was suggested in naturally infected cattle (Lindsay et al., 2000). Cryptosporidium andersoni was previously recognised as ‘‘ C. muris genotype A’’ or ‘‘C. muris calf genotype’’ based on its unique ssrRNA sequence (Morgan et al., 2000). The ssrRNA sequence is 1,743 bp long (GenBank: AF093496); it was obtained from oocysts isolated from the faeces of a cow in the USA. In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382352, AF221542, AF266262). Molecular characterisation of Cryptosporidum isolates from HIVpositive human subjects in France have revealed the presence of partial ssrRNA that very likely represents C. andersoni in a single male patient with a very low CD4 cell count (Guyot et al., 2001). Therefore, C. andersoni is rarely, if ever, detected in human subjects. This parasite species is of minor public health significance.

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3.17. Species XVll. Cryptosporidium canis Fayer, Trout, Xiao, Morgan, Lal & Dubey, 2001

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Cryptosporidium canis infects domestic dogs (Canis familiaris) worldwide (Fayer et al., 2001). Very little is known about this species and the disease it causes in dogs. Endogenous stages and aspects of its development are not known. Attempts to infect laboratory mice were not successful, but it is experimentally infective to cattle (Fayer et al., 2001). Cryptosporidium canis was previously recognised as ‘‘Cryptosporidium sp. (canine genotype)’’ (Pieniazek et al., 1999) or ‘‘dog genotype [type]’’ based on its unique ssrRNA sequence. The ssrRNA sequence is 1,741 bp long (GenBank: AF112576), and it was obtained from oocysts isolated from faeces of a dog in the USA. In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382340, AF221529, AF221529). Based on very similar ssrRNA sequences, C. canis has been detected in coyotes (Canis latrans) (Trout et al., 2006) and red foxes (Vulpes vulpes) (Nolan et al., 2013). Cryptosporidium canis is occasionally detected in humans. A dog-to-human transmission was suspected in a household with two symptomatic children and an asymptomatic dog shedding C. canis oocysts (Xiao et al., 2007a). However, due to the limited number of human infections, the risk factor for C. canis cannot be established. During routine typing of 14,469 human cryptosporidiosis cases in the UK, only a single adult male subject with a recent history of travel to the Indian subcontinent was infected with C. canis (Elwin et al., 2012a). In Peru, two non-symptomatic children were infected with C. canis including a child less than 1 year old (Xiao et al., 2001). In addition, an HIV-infected patient was demonstrated to shed C. canis for 3 months (Pieniazek et al., 1999). This parasite species is therefore of minor public health significance.

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3.18. Species XVlll. Cryptosporidium hominis Morgan-Ryan, Fall, Ward, Hijjawi, Sulaiman, Fayer, Thompson, Olson, Lal & Xiao, 2002

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Cryptosporidium hominis causes diarrhoea in humans (Homo sapiens) and has been associated with community outbreaks worldwide. The species was formerly known as the human geno-

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type, H, 1 (‘‘type 1’’) or anthroponotic genotype of the C. parvum species complex. Analysis of the genetic heterogeneity of the complex led to the identification of two independent human transmission cycles associated with two distinct genetic types, anthroponotic and zoonotic (Morgan et al., 1995; Carraway et al., 1996; Peng et al., 1997). Currently, a number of DNA markers and techniques are available to recognise C. hominis, including identification of a unique ssrRNA sequence. Infection with C. hominis appears to be symptomatic in human subjects with no asymptomatic shedders, based on the experimental infection of healthy volunteers (Chappell et al., 2006). An experimental challenge of healthy adult volunteers with C. hominis isolate TU502 led to clinical illness in 40% of subjects receiving 10 oocysts and a serum IgG response in subjects who received more than 30 oocysts (Chappell et al., 2006). Surveys of storm water employing ssrRNA amplification and sequencing have demonstrated the occasional presence of C. hominis (Jiang et al., 2005; Chalmers et al., 2010; Ruecker et al., 2012). Experimentally, a neonatal piglet model has been used in the laboratory to study the pathogenesis of cryptosporidiosis caused by C. hominis (Tzipori et al., 1994; Moore et al., 1995; Pereira et al., 2002). Cryptosporidium hominis is poorly infective for calves or lambs, but it can be maintained experimentally with relatively high doses of oocysts (Tzipori et al., 1994; Moore et al., 1995; Giles et al., 2001; Tanriverdi et al., 2003; Smith et al., 2005). Natural infections in cattle, goats and sheep occur but are rare or below the detection limit of the usual assays (Tanriverdi et al., 2003; Ryan et al., 2005; Giles et al., 2009). The host range and involvement in monkeys is yet to be fully understood (Li et al., 2011). The parasites infect the microvillous border of intestinal epithelial cells in immunocompetent as well as immunocompromised hosts (MorganRyan et al., 2002). In at least one case, C. hominis was identified as the species infecting the respiratory tract of a patient with HIV (Mercado et al., 2007). The species belongs to the C. parvum species complex divided into several species based on gene sequences, including ssrRNA. The human-specific species, C. hominis, is defined by isolates from Australia (Morgan-Ryan et al., 2002). For comparative purposes the unique ssrRNA of C. hominis ssrRNA (GenBank: AF108865) identifies the species. The ssrRNA sequence of C. hominis is 1,753 bp long (GenBank: AF108865) and is identical to the ssrRNA in the publically available genome of the TU502 isolate (GenBank: AAEL00000000) (Xu et al., 2004). Actin, COWP1 and HSP70 sequences are available (GenBank: AF382337, AF266265, AF221535). The TU502 isolate was obtained from a child with cryptosporidiosis and later propagated in gnotobiotic piglets; the TU502 isolate is not the type isolate (Akiyoshi et al., 2002; Morgan-Ryan et al., 2002). Nevertheless it is considered to serve this purpose. The main risk factor for C. hominis infection is travel, contact with another person with diarrhoea and changing young children’s nappies (Hunter et al., 2004). This parasite species is of major public health significance. 3.19. Species XIX. Cryptosporidium molnari Alvarez-Pellitero & SitjàBobadilla, 2002 Cryptosporidium molnari was described from naturally infected gilt-head sea bream (Sparus aurata) in the Mediterranean and Cantabric and Atlantic Spanish waters (Alvarez-Pellitero and SitjàBobadilla, 2002). Host specificity is not fully understood. However, C. molnari from gilt-head sea bream has been transmitted experimentally to the European sea bass (Dicentrarchus labrax) (SitjàBobadilla and Alvarez-Pellitero, 2003). The developmental stages occur preferentially in epithelial cells of the gastric mucosa, with sporogonial stages observed deep in the epithelium (Alvarez-Pelli-

tero and Sitjà-Bobadilla, 2002). Molecular surveys suggest a possible worldwide distribution and the ability of C. molnari to infect a range of fish, including ornamental fish in pet shops and the Murray cod (Maccullochella peelii peelii) in Australia (Zanguee et al., 2010; Baragahare et al., 2011). A mild to moderate gastritis has been reported in the Murray cod (Baragahare et al., 2011). A significant relationship between the presence of C. molnari and both fish weight and season was demonstrated in a long-term epidemiological study (Sitjà-Bobadilla et al., 2005). Cryptosporidium molnari is defined by its unique ssrRNA sequence and localisation of the developmental stages in the stomachs of fish (Alvarez-Pellitero and Sitjà-Bobadilla, 2002; Palenzuela et al., 2010). The partial ssrRNA sequence is 823 bp long (GenBank: HM243547), and was obtained from oocysts in the gastric mucosa from gilt-head sea bream (Palenzuela et al., 2010). In addition, an actin sequence is available (GenBank: HM365219). This species has no zoonotic potential and is of no public health significance.

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3.20. Species XX. Cryptosporidium suis Ryan, Monis, Enemark, Sulaiman, Samarasinghe, Read, Buddle, Robertson, Zhou, Thompson & Xiao, 2004

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Cryptosporidium suis is the dominant species infecting pigs (Sus scrofa) younger than 5 weeks of age worldwide (Ryan et al., 2004b). Cryptosporidium suis infects all age categories of pigs, although prevalence is lower in older pigs compared with C. scrofarum (Jeníková et al., 2011). Infection with C. suis is asymptomatic and the presence of diarrhoea in pigs is not associated with C. suis (Jeníková et al., 2011). However, concomitant infection of C. suis with rotavirus has led to fatalities in piglets (Enemark et al., 2003). Development is confined to the pig’s small and large intestine with a patent period of 9–15 days (Enemark et al., 2003). The species has been experimentally transmitted to cattle, with very low numbers of detectable oocysts in faeces (Enemark et al., 2003), and is not infective to nude mice (Morgan et al., 1999a). Cryptosporidium suis was previously recognised as ‘‘Cryptosporidium pig genotype I’’ or ‘‘C. parvum porcine genotype’’ based on its unique ssrRNA sequence (Morgan et al., 1999a; Enemark et al., 2003). The ssrRNA sequence is 1,749 bp long (GenBank: AF115377) obtained from oocysts isolated from a pig. In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382344, AF221533, AF266270). A single published case demonstrated the ssrRNA signature of C. suis in one HIV-positive 24-year old human who presented without symptoms of cryptosporidiosis (Xiao et al., 2002). Despite the above report, this species has low zoonotic potential. This parasite species is of minor public health significance.

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3.21. Species XXl. Cryptosporidium scophthalmi Alvarez-Pellitero, Quiroga, Sitjà-Bobadilla, Redondo, Palenzuela, Padrós, Vázquez & Nieto, 2004

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Cryptosporidium scophthalmi was described from cultured turbots (Scophthalmus maximus) from fish farms at different sites in Spain (Atlantic Ocean), with maximum prevalence observed in young fish (4–100 g) (Alvarez-Pellitero et al., 2004). The developmental stages occur in the distal intestine (mainly in the rectum) and rarely in epithelial cells of the stomach (Alvarez-Pellitero et al., 2004). The species is described based on its intestinal location and presumed strict specificity to turbot (Alvarez-Pellitero et al., 2004). In an epidemiological study, the poor condition of young fish during spring or summer was significantly associated with the presence of C. scophthalmi (Alvarez-Pellitero et al., 2009). Cryptosporidium scophthalmi is not defined by ssrRNA sequence data or any other genetic marker.

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This parasite species has no zoonotic potential. It is of no public health significance.

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3.22. Species XXll. Cryptosporidium pestis Šlapeta, 2006

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Cryptosporidium pestis is a generalist species, infecting mammals. It is the principal zoonotic species worldwide and the cause of cryptosporidiosis in calves and humans. In the literature, this species has been designated as genotype C, 2 (‘‘type 2’’) or bovine type of the C. parvum species complex. As mentioned in Section 3.18, analysis of the genetic heterogeneity of the C. parvum species complex led to the identification of two independent human transmission cycles associated with two distinct genetic types of Cryptosporidium (Morgan et al., 1995; Carraway et al., 1996; Peng et al., 1997). Currently, a number of DNA markers and techniques are available to recognise C. pestis, including identification of a unique ssrRNA sequence. A major reservoir for C. pestis under farm conditions is young, preweaned calves under 2 weeks of age (Santín et al., 2004). On the other hand, cattle over 2 months of age are less likely to shed C. pestis (Atwill et al., 1999). The development is confined to the small intestine, however, in mice it extends to the pyloric region of the stomach (Griffiths, 1998). Surveys of faecal samples employing ssrRNA sequencing of domestic and wild animals as well as pristine water and storm water samples have demonstrated a global distribution of C. pestis. For example, during a long-term study of Cryptosporidium in the agriculturally-intensive South Nation River watershed in Ontario, Canada, the human pathogenic species, C. pestis and C. hominis, represented only 1.6% of detected samples (Ruecker et al., 2012). This highly successful species, C. pestis, also known as the bovine, C or 2 genotype, is for comparative purposes identified by its unique ssrRNA sequence. The ssrRNA sequence of C. pestis is 1,749 bp long (GenBank: AF108864), and is identical to the ssrRNA sequence in the genome of the Iowa isolate (GenBank: AAEE00000000) (Abrahamsen et al., 2004). Actin, HSP70 and COWP1 sequences are available (GenBank: M86241, U11761, BX538351). The Iowa isolate, originally from a calf ( B. taurus), serves as the type for the species (Šlapeta, 2006). There is significant variation in the degree of infectivity of C. pestis for humans. The infectivity of C. pestis isolates (‘Iowa’ from a calf and ‘TAMU’ from a horse) for healthy adults demonstrated a trend toward a longer duration of diarrhoea of 94.5 h for TAMU compared to 64.2 h for Iowa (Okhuysen et al., 1999). The median infective dose (ID50) is as low as nine oocysts of TAMU for a healthy adult (Okhuysen et al., 1999). The patent period lasts from 3.3 to 8.4 days (Okhuysen et al., 1999). In contrast to C. hominis, asymptomatic shedding was seen in volunteers infected with C. parvum. The site of infection is the villous surface throughout the small intestine. In some heavy infections and in immunocompromised individuals infection can extend to adjacent organs and ducts (i.e. colon, biliary system). In an experimental infection with the Iowa isolate, healthy human subjects mounted a serum IgG response to reinfection after they failed to mount a serum IgG response to a primary challenge (Okhuysen et al., 1998). The main risk factor for C. pestis infection is contact with cattle (Hunter et al., 2004). However, C. pestis can be transmitted from person-to-person, therefore the origin can be either bovine or human. This parasite species is of major public health significance.

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3.23. Species XXlll. Cryptosporidium fayeri Ryan, Power & Xiao, 2008

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Cryptosporidium fayeri infects marsupials in Australia, including kangaroos, wallabies and koalas (Ryan et al., 2008). The infection is asymptomatic in its hosts. It was formally described from the fae-

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ces of naturally infected red kangaroos (Macropus rufus) (Power and Ryan, 2008) and based on the ssrRNA sequence found in the faeces of koalas (Phascolarctos cinereus) and eastern grey kangaroos (Macropus giganteus) (Power et al., 2004; Ryan et al., 2008). Attempts to infect laboratory animals were not successful (Ryan et al., 2008). No developmental stages have been described and the patent period is unknown. Cryptosporidium fayeri was previously recognised as ‘‘Cryptosporidium ‘marsupial’ genotype I’’ (Power et al., 2004) based on its unique ssrRNA sequence. The ssrRNA sequence is 1,750 bp long (GenBank: AF112570) and was obtained from oocysts isolated from the faeces of a red kangaroo. In addition, actin, HSP70 and COWP1 sequences are available (GenBank: AF382345, AF221531, AF266269). An unprecedented case report demonstrated the ssrRNA signature of C. fayeri in one 29-year old human suffering from prolonged gastrointestinal illness (Waldron et al., 2010). The affected woman reported frequent contact with partially domesticated marsupials (Waldron et al., 2010). To date, this is the only record of C. fayeri in a human; together with the inability to establish experimental infections, this species has minor zoonotic potential.

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3.24. Species XXlV. Cryptosporidium ryanae Fayer, Santín & Trout, 2008

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Cryptosporidium ryanae infects cattle ( B. taurus), commonly post-weaned calves, worldwide. The species was previously recognised as the ‘‘Cryptosporidium deer-like genotype’’ (Santín et al., 2004). In a study in the USA in which calves were monitored over 2 years, a cumulative prevalence attained 60%, the greatest number of infected calves being 18–20 weeks of age (Santín et al., 2008). In a study in Sweden, 6% of pre-weaned and 17% of post-weaned calves were infected with C. ryanae (Silverlås et al., 2010b). The clinical significance of C. ryanae infection is not documented in controlled experimental trials. The infection with this species is most likely asymptomatic (Silverlås et al., 2010a,b). Oocysts of C. ryanae from the faeces of an 18 week old dairy calf were used for the formal description and the same oocysts were used in experimental infections (Fayer et al., 2008). The endogenous stages are not known. Experimental infection was successful in 3 week old dairy calves, but was unsuccessful in lambs and two 5 day old BALB/c mice (Fayer et al., 2008). The patent period was 15–17 days in calves (Fayer et al., 2008). The host range of C. ryanae appears to be restricted to cattle. The species C. ryanae is defined primarily on its unique ssrRNA sequence; a partial ssrRNA is 771 bp long (GenBank: EU410344). In addition, actin and HSP70 sequences are available (GenBank: EU410345, EU410346). This parasite species has no zoonotic potential and is of no public health significance.

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˚, Valigurová, Koudela, 3.25. Species XXV. Cryptosporidium fragile Jirku ´ & Šlapeta, 2008 Krˇízˇek, Modry

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The species C. fragile is the only species infecting amphibians. It infects a single frog species, the black-spined toad (Duttaphrynus melanostictus) (Jirku˚ et al., 2008; Valigurová et al., 2008). The parasite infects the stomach. The infected toads showed signs of weight loss, weakness and/or apathy that subsided together with eventual spontaneous disappearance of infection. Histopathological examination of the affected gastric tissues showed mild mucosal thickening. The frogs were originally collected in the Malay Peninsula (no specific locality is known). Therefore, the description is based on hosts imported for the pet trade. The patent period is 22–46 days but is intermittent.

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Cryptosporidium fragile is recognised by its unique ssrRNA sequence. The ssrRNA sequence is 1,745 bp long (GenBank: EU162751). No additional sequence material is available. This parasite species has no zoonotic potential and is of no public health significance.

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3.26. Species XXVl. Cryptosporidium macropodum Power, Ryan, 2008

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Cryptosporidium macropodum infects marsupials in Australia, specifically kangaroos and wallabies, and has never been recorded from humans (Power and Ryan, 2008). The infection is asymptomatic despite excretion of very high numbers of oocysts (Power et al., 2005; Power and Ryan, 2008). It was formally described from the faeces of wild eastern grey kangaroos (M. giganteus) (Power and Ryan, 2008). Based on ssrRNA sequence, C. macropodum is found naturally in the swamp wallaby (Wallabia bicolor), the western grey kangaroo (Macropus fuliginosus) and the red kangaroo (M. rufus). No experimental transmission has been undertaken. No developmental stages are described and the patent period is unknown. Cryptosporidium macropodum was previously recognised as ‘‘Cryptosporidium ‘marsupial’ genotype II (EGK3)’’ (Power et al., 2004) based on its unique ssrRNA sequence. The ssrRNA sequence is 1,803 bp long (GenBank: AF513227). It was obtained from oocysts isolated from the faeces of an eastern grey kangaroo. In addition, actin, HSP70 and COWP1 sequences are available (GenBank: EU124664, AY237634, AY237635). This emerging parasite species has no zoonotic potential and is of no public health significance.

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3.27. Species XXVll. Cryptosporidium ducismarci Traversa, 2010

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Cryptosporidium ducismarci infects tortoises, including the marginated tortoise (Testudo marginata) (Traversa et al., 2008). Confirmed records are restricted to a handful of reports (PedrazaDíaz et al., 2009; Griffin et al., 2010; Traversa, 2010). In two cases the presence of intestinal development of C. ducismarci was associated with intestinal pathology in histological sections from dead tortoises (Griffin et al., 2010). The unique ssrRNA sequence is reported only from captive pet tortoises. The natural host is unknown and the association of C. ducismarci with pet tortoises is not understood. Cryptosporidium ducismarci is characterised by its unique ssrRNA sequence (Traversa, 2010). The partial ssrRNA sequence is 531 bp long (GenBank: EF547155). In addition, actin and COWP1 sequences are available (GenBank: EF519704). This species has no zoonotic potential and is of no public health significance.

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3.28. Species XXVlll. Cryptosporidium ubiquitum Fayer, Santín & Macarisin, 2010 Cryptosporidium ubiquitum infects mammals including humans worldwide and is considered a generalist among mammals. Cryptosporidium ubiquitum was previously recognised as the ‘‘Cryptosporidium cervine genotype’’ (or ‘‘deer’’ or ‘‘genotype 3’’). The zoonotic potential of C. ubiquitum was first recognised in a study from Canada, by the demonstration of the unique ssrRNA of C. ubiquitum in faecal samples from paediatric patients (less than 10 years of age) with clinical symptoms consistent with cryptosporidiosis (Ong et al., 2002). The unique ssrRNA was known from wildlife and storm water in New York, USA (Xiao et al., 2000; Perz and Le Blancq, 2001). An association with either pre-weaned or post-weaned lambs/sheep has been suggested in several surveys (Ryan et al., 2005; Santín and Fayer, 2007; Shen et al., 2011), including a claim that sheep serve as the reservoir for C. ubiquitum (Santín and Fayer, 2007). However, the distribution and host range

are not fully understood (Fayer et al., 2010). It appears that C. ubiquitum is commonly identified in water samples from both pristine as well as waste water (McCarthy et al., 2008; Chalmers et al., 2010). Recently, C. ubiquitum was formally defined using a series of infection trials demonstrating its capacity to infect a wide range of laboratory animals. The original isolate came from captive Brazilian prehensile-tailed porcupines (Coendou prehensilis) from a zoo in the USA. At that time, the porcupines had diarrhoea but only one infant was in poor health; 60 days later no C. ubiquitum was detected. The isolate was infective sequentially to 3-month-old goats and calves, as well as to 69-day-old Mongolian gerbils and 5-day-old suckling BALB/c mice, all shedding C. ubiquitum with the same unique ssrRNA signature (GenBank: HM209366). In a 5-days-old male Holstein-Friesian calf (B. taurus), the patent period was 12 days. Endogenous stages and development are not known. All animals experimentally infected with C. ubiquitum remained asymptomatic. Cryptosporidium ubiquitum is characterised by its unique ssrRNA sequence (Fayer et al., 2010). The partial ssrRNA sequence is 725 bp long (GenBank: HM209366). In addition, actin and COWP1 sequences are available (GenBank: HM209377, HM209388). Person-to-person transmission has been suggested (Fayer et al., 2010), but the epidemiology and seasonal risk associated with transmission of C. ubiquitum have not been evaluated due to the rarity of cases in humans (Chalmers et al., 2009a). The high prevalance of C. ubiquitum in sheep and reports from pristine water are not reflected in zoonotic outbreak(s) (Ryan et al., 2005). This emerging parasite species is of minor public health significance.

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3.29. Species XXlX. Cryptosporidium viatorum Elwin, Hadfield, Robinson, Crouch & Chalmers, 2012

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This recently identified species infects humans (Elwin et al., 2012b). The parasite has unique ssrRNA signatures (GenBank: JN846705, JN846708) and was discovered in travellers returning to the UK from the Indian subcontinent, suffering clinical cryptosporidiosis. In a case series (n = 10), clinical signs lasting 13– 30 days included diarrhoea, abdominal pain, fever, nausea and in one case vomiting. None of the patients was immunocompromised, two had Campylobacter sp. infections and one was concurrently shedding C. meleagridis and Giardia duodenalis (assemblage not reported) (Elwin et al., 2012b). Currently, only a few cases in persons returning to the UK from the Indian subcontinent have been reported, being diagnosed predominantly between January to March, compared with July to September for C. hominis and C. pestis. Cryptosporidium viatorum appears to be distributed globally, because it has been reported in two patients who travelled to Kenya and Guatemala respectively (Insulander et al., 2013). It has been suggested that the 20-29 year age group is more likely to be infected with C. viatorum compared with C. hominis or C. pestis (Hunter et al., 2004; Elwin et al., 2012b). The site of C. viatorum infection within the gastrointestinal tract is not known. Cryptosporidium viatorum is characterised by its unique ssrRNA sequence (GenBank: JN846705, JN846708) (Elwin et al., 2012b). The partial type ssrRNA sequence (766 bp long; GenBank: JN846705) of C. viatorum differs by one nucleotide (G/A) from that obtained in the original study from a different patient (GenBank: JN846708). No identical sequences have been recovered in other hosts. In addition, actin and HSP70 sequences are available (GenBank: JN846706, JN846706). Person-to-person transmission has not been unambiguously documented, and the epidemiology and seasonal risk associated with transmission of C. viatorum require further elucidation (Elwin et al., 2012b). Due to the limited knowledge of this emerging par-

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asite species, it is considered of moderate public health significance. 3.30. Species XXX. Cryptosporidium scrofarum Kvácˇ, Kestrˇánová, Pinková, Kveˇtonˇová, Kalinová, Wagnerová, Kotková, Vítovec, Ditrich, McEvoy, Stenger & Sak, 2013

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Cryptosporidium scrofarum infects domestic pigs and Eurasian wild boars ( S. scrofa) (García-Presedo et al., 2013; Kvácˇ et al., 2013). The species appears to be cosmopolitan in domestic and wild pigs, usually with a low prevalence, <5%, and in pigs over 6 weeks old (Kvácˇ et al., 2013). Pigs infected with C. scrofarum are asymptomatic and the parasite multiplies in the duodenum, jejunum and ileum based on histological investigation of experimentally infected pigs (Kvácˇ et al., 2013). Attempts to infect laboratory animals were not successful (Kvácˇ et al., 2013). Cryptosporidium scrofarum has previously been recognised as the ‘‘Cryptosporidium pig genotype II’’, a name coined for its unique ssrRNA sequence (Ryan et al., 2003a). The partial ssrRNA sequence of C. scrofarum is 768 bp long (GenBank: JX424840). In addition, actin and HSP70 sequences are available (GenBank: JX424841, JX424842). A singular record demonstrated the ssrRNA signature of C. scrofarum and G. duodenalis (assemblage A) in a 29-year old human suffering from diarrhoea (Kvácˇ et al., 2009). The study looked at 457 stool samples from 203 immunocompetent patients (less than 69 years of age) suspected to suffer from cryptosporidiosis (Kvácˇ et al., 2009). Thus far this is the only record of C. scrofarum in humans. Together with the inability to establish experimental infection in hosts other than pigs, it is likely that this species has minor zoonotic potential.

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4. Future prospects

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Cryptosporidium spp. are recognised as major waterborne parasites worldwide (Baldursson and Karanis, 2011). Of the 30 species recognised above, 14 have been documented to infect humans. Two, four and eight named species are considered of major, moderate and minor public health significance, respectively. There are at least nine named species that are shared between humans and cattle (Table 1). This synopsis of 30 species was based on a re-evaluation of data in the literature relevant to the validity of Cryptosporidium spp. All 30 species have a valid name conforming to the rules of the ICZN. To suppress any of the names in favour of another name requires the preparation of a formal case, which must be referred to the ICZN Commission for a ruling (Šlapeta, 2011). Recently, two such circumstances related to C. parvum and C. tyzzeri have been suggested, but no action – in terms of a formal case – has as yet been taken. Consequently, the system presented above remains valid (Šlapeta, 2007, 2012; Xiao et al., 2007b, 2012). The nomenclatural discussion apart, a Roman numeral system has been introduced (Table 1, Fig. 1). The Roman numeral system for each valid species and the synopsis with reference sequences identifies all valid Cryptosporidium spp. to date. Humans are infected most frequently with C. hominis (Species XVIII) and C. pestis (Species XXII), belonging to the Cryptosporidium parvum species complex (Fig. 1). Isolates of both species have been the subject of population genetics studies demonstrating the presence of complex clonal to panmictic populations within the species complex (Jex and Gasser, 2010; Widmer and Sullivan, 2012). The addition of all members of the species complex including C. parvum (Species II) and C. cuniculus (Species VII) will enable the discovery and understanding of the genetic basis of host–parasite relationships leading to emerging infectious diseases. Whole genome amplifica-

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tion, partial genome sequencing and the establishment of farm animal experimental models will dramatically expand in the near future (Bouzid et al., 2010; Widmer et al., 2012; Widmer and Sullivan, 2012; Grinberg et al., 2013). Consequently, a universally acceptable naming scheme for the species of Cryptosporidium is not just an interesting academic exercise, but reflects an urgent need for integrated research on apicomplexan parasites and their role in public health (Šlapeta, 2009).

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Appendix A. Supplementary data

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Abrahamsen, M.S., Templeton, T.J., Enomoto, S., Abrahante, J.E., Zhu, G., Lancto, C.A., Deng, M., Liu, C., Widmer, G., Tzipori, S., Buck, G.A., Xu, P., Bankier, A.T., Dear, P.H., Konfortov, B.A., Spriggs, H.F., Iyer, L., Anantharaman, V., Aravind, L., Kapur, V., 2004. Complete genome sequence of the apicomplexan, Cryptosporidium parvum. Science 304, 441–445. Agosta, S.J., Janz, N., Brooks, D.R., 2010. How specialists can be generalists: resolving the ‘‘parasite paradox’’ and implications for emerging infectious disease. Zoologia 27, 151–162. Akiyoshi, D.E., Feng, X., Buckholt, M.A., Widmer, G., Tzipori, S., 2002. Genetic analysis of a Cryptosporidium parvum human genotype 1 isolate passaged through different host species. Infect. Immun. 70, 5670–5675. Akiyoshi, D.E., Dilo, J., Pearson, C., Chapman, S., Tumwine, J., Tzipori, S., 2003. Characterization of Cryptosporidium meleagridis of human origin passaged through different host species. Infect. Immun. 71, 1828–1832. Alvarez-Pellitero, P., Sitjà-Bobadilla, A., 2002. Cryptosporidium molnari n. sp. (Apicomplexa: Cryptosporidiidae) infecting two marine fish species, Sparus aurata L. and Dicentrarchus labrax L. Int. J. Parasitol. 32, 1007–1021. Alvarez-Pellitero, P., Quiroga, M.I., Sitjà-Bobadilla, A., Redondo, M.J., Palenzuela, O., Padrós, F., Vázquez, S., Nieto, J.M., 2004. Cryptosporidium scophthalmi n. sp. (Apicomplexa: Cryptosporidiidae) from cultured turbot Scophthalmus maximus. Light and electron microscope description and histopathological study. Dis. Aquat. Organ. 62, 133–145. Alvarez-Pellitero, P., Perez, A., Quiroga, M.I., Redondo, M.J., Vázquez, S., Riaza, A., Palenzuela, O., Sitjà-Bobadilla, A., Nieto, J.M., 2009. Host and environmental risk factors associated with Cryptosporidium scophthalmi (Apicomplexa) infection in cultured turbot, Psetta maxima (L.) (Pisces, Teleostei). Vet. Parasitol. 165, 207– 215. Angus, K.W., Hutchison, G., Munro, H.M., 1985. Infectivity of a strain of Cryptosporidium found in the guinea-pig (Cavia porcellus) for guinea-pigs, mice and lambs. J. Comp. Pathol. 95, 151–165. Antunes, R.G., Simoes, D.C., Nakamura, A.A., Meireles, M.V., 2008. Natural infection with Cryptosporidium galli in canaries (Serinus canaria), in a cockatiel (Nymphicus hollandicus), and in lesser seed-finches (Oryzoborus angolensis) from Brazil. Avian Dis. 52, 702–705. Atwill, E.R., Johnson, E.M., Pereira, M.G., 1999. Association of herd composition, stocking rate, and duration of calving season with fecal shedding of Cryptosporidium parvum oocysts in beef herds. J. Am. Vet. Med. Assoc. 215, 1833–1838. Bajer, A., Cacciò, S., Bednarska, M., Behnke, J.M., Pieniazek, N.J., Sinski, E., 2003. Preliminary molecular characterization of Cryptosporidium parvum isolates of wildlife rodents from Poland. J. Parasitol. 89, 1053–1055. Baldursson, S., Karanis, P., 2011. Waterborne transmission of protozoan parasites: review of worldwide outbreaks – an update 2004–2010. Water Res. 45, 6603– 6614. Ballweber, L.R., Panuska, C., Huston, C.L., Vasilopulos, R., Pharr, G.T., Mackin, A., 2009. Prevalence of and risk factors associated with shedding of Cryptosporidium felis in domestic cats of Mississippi and Alabama. Vet. Parasitol. 160, 306–310. Baragahare, R., Becker, J.A., Landos, M., Šlapeta, J., Dennis, M.M., 2011. Gastric cryptosporidiosis in farmed Australian Murray cod, Maccullochella peelii peelii. Aquaculture 314, 1–6. Barker, I.K., Carbonell, P.L., 1974. Cryptosporidium agni sp.n. from lambs, and Cryptosporidium bovis sp.n. from a calf, with observations on the oocyst. Z. Parasitenkd. 44, 289–298. Bednarska, M., Bajer, A., Kulis, K., Sinski, E., 2003. Biological characterisation of Cryptosporidium parvum isolates of wildlife rodents in Poland. Ann. Agric. Environ. Med. 10, 163–169. Bornay-Llinares, F.J., da Silva, A.J., Moura, I.N., Myjak, P., Pietkiewicz, H., KruminisLozowska, W., Graczyk, T.K., Pieniazek, N.J., 1999. Identification of Cryptosporidium felis in a cow by morphologic and molecular methods. Appl. Environ. Microbiol. 65, 1455–1458. Bouzid, M., Heavens, D., Elwin, K., Chalmers, R.M., Hadfield, S.J., Hunter, P.R., Tyler, K.M., 2010. Whole genome amplification (WGA) for archiving and genotyping of clinical isolates of Cryptosporidium species. Parasitology 137, 27–36.

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Please cite this article in press as: Šlapeta, J. Cryptosporidiosis and Cryptosporidium species in animals and humans: A thirty colour rainbow? Int. J. Parasitol. (2013), http://dx.doi.org/10.1016/j.ijpara.2013.07.005

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