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Ryan, U. (2010) Cryptosporidium in birds, fish and amphibians. Experimental Parasitology, 124 (1). pp. 113-120.

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Cryptosporidium in birds, fish and amphibians

Una Ryan

PII: S0014-4894(09)00034-4 DOI: 10.1016/j.exppara.2009.02.002 Reference: YEXPR 5701

To appear in: Experimental Parasitology

Received Date: 17 November 2008 Revised Date: 3 February 2009 Accepted Date: 5 February 2009

Please cite this article as: Ryan, U., Cryptosporidium in birds, fish and amphibians, Experimental Parasitology (2009), doi: 10.1016/j.exppara.2009.02.002

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Cryptosporidium in birds, fish and amphibians

A/Prof. Una Ryan

Division of Veterinary and Biomedical Sciences, Murdoch University, Murdoch, Western Australia,

6150.

______

Corresponding author: Una Ryan

Phone: 08 9360 2482

Fax: 08 9310 4144

E-mail: [email protected]

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Abstract

Whilst considerable information is available for avian cryptosporidiosis, scant information is available for Cryptosporidium infections in fish and amphibians. The present review details recent studies in avian cryptosporidiosis and our current knowledge of piscine and amphibian infections.

Keywords: Cryptosporidium, birds, fish, amphibians, avian genotypes.

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1.0 Introduction

Cryptosporidiosis is one of the most prevalent parasitic infections in domesticated, caged and wild birds (O'Donoghue, 1995, Sreter and Varga, 2000), and the parasite has been reported in more than 30 avian species worldwide (Lindsay and Blagburn, 1990, O'Donoghue, 1995, Sreter and

Varga, 2000, Ng, et al., 2006; Ryan and Xiao, 2008). There have been extensive studies conducted on avian cryptosporidiosis, however, our knowledge of piscine and amphibian cryptosporidiosis is much less complete. Cryptosporidium has been described in fresh and marine water fish, as well as frogs and toads but little is known about the epidemiology, pathogenicity and zoonotic transmission of piscine and amphibian species of Cryptosporidium.

1.1 Cryptosporidium in Birds

Currently only three avian Cryptosporidium spp. are recognised; Cryptosporidium meleagridis, Cryptosporidium baileyi and Cryptosporidium galli. Two other species of

Cryptosporidium have been named from birds: Cryptosporidium anserinum from a domestic goose

(Anser domesticus) (Proctor and Kemp, 1974) and Cryptosporidium tyzzeri from chickens (Gallus gallus domesticus) (Levine, 1961). Neither of these reports gave adequate description of oocysts or provided other useful information and are therefore not considered valid species (Lindsay and

Blagburn, 1990). A number of genetically distinct avian genotypes have however recently been described, which may be re-described as species in the future once more biological information becomes available.

Naturally occurring cryptosporidiosis in birds manifests itself in three clinical forms; respiratory disease, enteritis and renal disease. Usually only one form of the disease is present in an outbreak (Lindsay and Blagburn, 1990). At present no effective chemotherapy is available for the treatment of avian cryptosporidiosis (Lindsay and Blagburn, 1990, Ryan and Xiao, 2008). Current control methods against avian cryptosporidiosis generally rely on limiting or preventing infection.

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1.2 Cryptosporidium meleagridis Slavin, 1955.

Tyzzer first described avian cryptosporidiosis in 1929 (Tyzzer, 1929) but it was not until

35 years later in 1955, that Slavin found a structurally similar parasite in the ileum of turkey poults and named the parasite C. meleagridis (Slavin, 1955). Oval oocysts measuring 4.5 x 4.0 µm appeared indistinguishable from those of C. parvum (Slavin, 1955). Lindsay et al., (1989a) gave measurements of 5.2 x 4.6 (4.5 - 6.0 x 4.2 - 5.3) µm for viable oocysts from turkey feces (Table 1).

Little is known of the life cycle of C. meleagridis. Both asexual and sexual stages were described by Slavin in 1955. More recently, the life cycle of what is presumed to be C. meleagridis has been described from naturally infected 30 day old commercial turkeys (Tacconi, et al., 2001).

Endogenous developmental stages of a C. meleagridis isolate of human origin have also been described in a mouse and a chicken experimentally infected with this isolate (Akiyoshi, et al.,

2003).

Cryptosporidium meleagridis is found in the small and large intestine and bursa and infection has been associated with enteritis and mortality (Slavin, 1955, Gharagozlou, et al., 2006; Pagès-

Manté et al., 2007). Like C. parvum, C. meleagridis appears to have a large host range and has been described from a wide range of avian species including turkeys, parrots, chickens, cockatiels and a red-legged partridge (O'Donoghue, 1995, Morgan, et al., 2000b, Sreter and Varga, 2000,

Darabus and Olariu, 2003, Abe and Iseki, 2004; Huber et al., 2007; Pagès-Manté et al., 2007;

Soltane et al., 2007). Experimental transmission studies have shown that C. meleagridis can infect broiler chickens, ducks, turkeys, calves, pigs, rabbits, rats, mice (O'Donoghue, 1995, Akiyoshi, et al., 2003, Darabus and Olariu, 2003, Huang, et al., 2003). Overall, the rate of infectivity and virulence of C. meleagridis for the mammalian species was similar to that of C. parvum (Akiyoshi, et al., 2003). Recently a natural C. meleagridis infection was reported in a dog from the Czech

Republic (Hajdusek, et al., 2004).

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Cryptosporidium meleagridis is also an emerging human pathogen and is the third most common Cryptosporidium parasite in humans (Xiao and Fayer, 2008). It has previously been suggested that C. meleagridis may be C. parvum (Champliaud, et al., 1998, Sreter and Varga,

2000), however molecular analysis of US, Australian and European isolates at the small subunit ribosomal RNA (SSU rRNA), heat shock protein 70 (HSP70), Cryptosporidium oocyst wall protein (COWP), actin and glycoprotein 60 (GP60) loci as well as the Cryptosporidium double stranded RNA virus have demonstrated the genetic and biological uniqueness of C. meleagridis

(Xiao, et al., 1999, Morgan, et al., 2000, Sreter, et al., 2000, Sulaiman, et al., 2000, Xiao, et al.,

2000, Morgan, et al., 2001, Sulaiman, et al., 2002, Xiao, et al., 2002).

Phylogenetic analyses suggests that C. meleagridis was possibly originally a mammalian

Cryptosporidium parasite which subsequently became established in birds via ingestion of infected prey (Xiao, et al., 2002). This is supported by the recent findings on the ability of C. meleagridis to infect a wide range of mammals, which is certainly not the case with C. baileyi.

1.3 Cryptosporidium baileyi Current, Upton and Haynes, 1986.

A second species of avian Cryptosporidium, originally isolated from commercial broiler chickens, was named C. baileyi based on its life cycle and morphologic features (Current, et al.,

1986). The life cycle of C. baileyi has been described in detail (Current, et al., 1986; Cheadle, et al.,

1999).

Natural C. baileyi infections have been reported in many anatomic sites in avian hosts including the conjunctiva, nasopharynx, trachea, bronchi, air sacs, small intestine, large intestine, ceca, cloaca, bursa of fabricius, kidneys and urinary tract (Lindsay and Blagburn, 1990). Following oral inoculation, the primary sites of development was the BF and cloaca (Lindsay, et al., 1986).

Intra-tracheal (IT) inoculation of oocysts into chickens resulted in extensive parasitization of the respiratory tract (Lindsay, et al., 1986, Blagburn, et al., 1987, Lindsay, et al., 1987a). Conjunctival infections occurred in some birds when oocysts were placed directly on the conjunctival sac

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(Lindsay, et al., 1987a). However, birds were more likely to develop infections in the BF and cloaca

(Lindsay, et al., 1987a).

Cryptosporidium baileyi is more frequently associated with respiratory cryptosporidiosis and high morbidity and mortality has been reported with C. baileyi respiratory infections of birds, especially broiler chickens (Lindsay and Blagburn, 1990; Sreter, et al., 1995, Sreter, et al., 2000).

Renal cryptosporidiosis also occurs in birds (Gardiner and Imes, 1984; Randall, 1986; Nakamura and Abe, 1988; Lindsay and Blagburn, 1990) and has been experimentally induced in specific- pathogen-free (SPF) chickens coinfected with C. baileyi and Marek's disease virus (MDV)

(Abbassi, et al., 1999).

Cryptosporidium baileyi is probably the most common avian Cryptosporidium sp., and has been reported in a wide range of avian hosts including black-headed gulls, chickens, cormorants, cranes, a channel-billed toucan, an eastern golden-backed weaver, turkeys, ducks, geese, cockatiels, a brown quail, a grey-bellied bulbul, an ostrich, a red-rumped cacique, a crested oropendola, a red crowned amazon, a rose-ringed parakeet, a grey partridge and mixed-bred falcons (Lindsay and

Blagburn, 1990, Pavlásek, 1993, Ryan, et al., 2003a, Abe and Iseki, 2004, Jellison, et al., 2004,

Kimura et al., 2004, Chvala, et al., 2006; Huber et al., 2007; Van Zeeland et al., 2008).

Experimental cross-transmission of C. baileyi to other birds including Japanese quail, domestic ducks, geese, pheasants, a chukar partridge and turkeys were successful with the exception of bobwhite quail (Current, et al., 1986, Lindsay, et al., 1987a, Lindsay, et al., 1989b,

Lindsay and Blagburn, 1990, Cardozo, et al., 2005). Limited life cycle stages were observed in some turkey poults and heavy infections developed only in the BF in 1 day old ducks and 2 day old geese (Current, et al., 1986). Mice and goats inoculated with C. baileyi oocysts, however, did not become infected (Current, et al., 1986).

Cryptosporidium baileyi has been analysed at a number of loci including the SSU rRNA,

HSP70, COWP and actin loci and has been shown to be genetically distinct (Xiao, et al., 1999,

Sulaiman, et al., 2000, Xiao, et al., 2000, Morgan, et al., 2001, Egyed, et al., 2002, Sulaiman, et al.,

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2002). Cryptosporidium baileyi is most closely related to avian genotype I and II (see 1.5.1). The relationship between C. baileyi and the rest of the intestinal and gastric parasites is unclear as phylogenetic analysis at the SSU rRNA, actin and COWP loci group C. baileyi with the intestinal parasites but analysis at the HSP70 locus placed C. baileyi in a cluster that contained all gastric

Cryptosporidium instead of the intestinal parasite cluster (Xiao, et al., 2002).

1.4 Cryptosporidium galli Pavlasek, 1999.

A third species of avian Cryptosporidium was first described by Pavlásek (Pavlásek, 1999,

Pavlásek, 2001) in hens on the basis of biological differences. More recently, the parasite was re- described on the basis of both molecular and biological differences (Ryan, et al., 2003b).

Cryptosporidium galli oocysts are statistically different in size (p < 0.05) from C. baileyi (6.3 x 5.2

µm) and C. meleagridis oocysts (5.2 x 4.6 µm) and measure 8.25 x 6.3 (8.0 - 8.5 x 6.2 - 6.4) µm, with a length to width ratio of 1.3 (Pavlásek, 1999, Pavlásek, 2001) (Table 1). Unlike other avian species, life cycle stages of C. galli developed in epithelial cells of the proventriculus and not the respiratory tract or small and large intestines (Pavlásek, 1999, Pavlásek, 2001).

Blagburn et al., (1990) may also have detected C. galli in birds when they used light and electron microscopy to characterize Cryptosporidium sp. in the proventriculus of an Australian diamond firetail finch that died of acute diarrhea. A subsequent publication also identified a species of Cryptosporidium infecting the proventriculus in finches and inadvertently proposed the name

Cryptosporidium blagburni in Table 1 of the paper (Morgan, et al., 2000). However, Pavlásek

(Pavlásek, 1999, Pavlásek, 2001) had provided a detailed description of what appeared to be the same parasite and named it C. galli. More recent molecular analyses have revealed C. galli and C. blagburni to be the same species (Ryan, et al., 2003a).

Little is known of the life cycle of C. galli. Endogenous developmental stages appear to be localized to glandular epithelial cells of the proventriculus. Life cycle stages ranging from oocysts to trophozoites and macrogamonts have been observed.

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Cryptosporidium galli appears to be associated with clinical disease and high mortality

(Blagburn, et al., 1990, Pavlásek, 1999, Morgan, et al., 2001, Pavlásek, 2001). Histopathology of infected finches demonstrated necrosis and hyperplasia of proventricular glandular epithelial cells associated with large numbers of Cryptosporidium oocysts attached to the surface of glandular epithelial cells (Morgan, et al., 2001).

Natural C. galli infections have been reported in finches, chickens, capercaille, pine grosbeak, turquoise parrots, greater flamingos, red cowled cardinals and rhinoceros hornbills

(Pavlásek, 1999, Pavlásek, 2001, Ryan, et al., 2003a, Ng, et al., 2006). In addition, morphologically similar oocysts have been observed in a variety of exotic and wild birds including Phasianidae,

Passeriformes and Icteridae (Ryan, et al., 2003b). A species which may have been C. galli has also been documented in the proventriculus of canaries (Tsai, et al., 1983). Future studies are required to determine the extent of the host range for C. galli. Cross-transmission studies have shown that C. galli oocysts were infectious for 9 day old but not 40 day old chickens (Pavlásek, 2001).

Distance, parsimony and maximum likelihood analysis of 3 loci (SSU rRNA, HSP70 and actin) identified C. galli as a distinct species (Ryan, et al., 2003b). The gastric location of C. galli in the host and its large size suggest that it is most closely related to the other gastric Cryptosporidium species and this has been supported by molecular analysis. At the SSU rRNA locus, C. galli is most closely related to avian genotype IV and the gastric parasites Cryptosporidium muris,

Cryptosporidium andersoni and Cryptosporidium serpentis.

1.5. Avian genotypes

In addition to the three recognized avian species, a total of 10 novel avian genotypes have been identified in birds. Cryptosporidium hominis and C. parvum have also been identified in birds

(see 1.5.4).

1.5.1 Avian Genotypes I-IV

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Avian genotypes I-IV have been identified from various avian hosts (Meireles, et al., 2006,

Ng, et al., 2006) (Tables 1 and 2). Sequence and distance analysis at the SSU rRNA gene locus indicated that although avian genotype I and avian genotype II were most closely related to C. baileyi, they were genetically distinct (~99.4 % and 97.6 % similarity respectively to C. baileyi) and exhibited ~98.2 % similarity to each other. At the actin gene locus, avian genotype I and avian genotype II exhibited only 95.7 % and 88.3 % similarity respectively to C. baileyi.

Attempted transmission of avian genotype II oocysts to two groups of 2 day old chickens did not result in infection as determined by histology, mucosal smears and fecal screening until 4 weeks PI (Meireles, et al., 2006). Avian genotype II has been reported to parasitize the cloacal epithelium and to a lesser extent the epithelium of the rectum and BF of ostriches (Santos, et al.,

2005). An earlier study reported a Cryptosporidium sp. in the feces of 14 of 165 (8.5 %) ostriches imported into Canada (Gajadhar, 1994). The mean size of 40 oocysts measured was 4.6 x 4.0 µm

(range 3.9 - 6.1 x 3.3 - 5.0 µm) with a shape index (length/width ratio) of 1.15 (range 1.00 - 1.38).

In cross-transmission experiments, this Cryptosporidium sp. failed to infect suckling mice, chickens, turkeys, or quail. Molecular data is not available for these ostrich isolates and therefore it is not possible to determine if the species described is the same as avian genotype II.

Avian genotype III isolates are genetically most closely related to the Eurasian woodcock genotype (98.8 % and 98.5 % genetic similarity at the SSU and actin loci respectively). Oocysts from avian genotype III measured 7.5 x 6.0 µm, which is larger than the mean oocyst size of C. serpentis (5.94 x 5.11 µm) but smaller than the oocyst size of the Eurasian woodcock-derived oocysts (8.5 x 6.4 µm). No clinical signs, such as diarrhea, dyspnea, coughing or sneezing, were detected in the avian hosts for avian genotypes I-III.

Avian genotype IV was identified in a Japanese whiteeye from the Czech Republic and exhibited ~97.9 % similarity to its closest relative C. galli at the SSU rRNA locus. The Japanese whiteeye exhibited diarrhea and anorexia. Microscopic analysis detected only Cryptosporidium

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oocysts; no pathogenic bacteria were observed. Oocysts from avian genotype IV were similar in size to C. galli and measured 8.25 x 6.3 µm.

1.5.2 Eurasian Woodcock Genotype

The Eurasian woodcock genotype groups most closely with avian genotype III and the gastric parasites (C. serpentis, C. muris, C. andersoni) (Ryan, et al., 2003a; Ng et al., 2006). All endogenous developmental stages including oocysts were detected in the proventriculus only.

1.5.3 Duck Genotype

A novel genotype was identified in a black duck (Morgan, et al., 2001), which appears to be most closely related to goose genotypes I and II at the SSU rRNA locus (96.9 - 97.5 % similarity respectively). Microscopic analysis of hematoxylin and eosin (H & E) stained sections from the black duck indicated an enteric infection with numerous oocysts attached to the apical surface of the enterocytes of the small intestine but clinical information on this bird was not available. The duck genotype has subsequently been identified in Canada geese (Branta canadensis) (Jellison, et al.,

2004, Zhou, et al., 2004).

1.5.4 Goose Genotypes I-IV

A recent study identified 5 Cryptosporidium sp. and genotypes from Canada geese collected from 13 sites in Ohio and Illinois; goose genotype I, goose genotype II, the duck genotype, C. parvum and C. hominis (Zhou, et al., 2004). Phylogenetic and sequence analysis of the SSU rRNA gene has revealed that goose genotypes I and II and the duck genotype are closely related. Another study identified five Cryptosporidium genotypes in Canada geese in the United States (Jellison, et al., 2004). Three of the Cryptosporidium genotypes belonged to goose genotypes I (geese 1, 2, 3a, 6 and 8) and II (goose 9) and the duck genotype (goose 5) whereas the remaining two genotypes

(geese 3b and 7) represented new Cryptosporidium genotypes; goose genotype III and IV

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respectively. Recently, C. parvum was confirmed by molecular analysis in an outbreak of intestinal cryptosporidiosis in Stone curlews kept in a mixed-species rearing unit in Dubai (Zylan et al., 2008), confirming the ability of C. parvum to infect avian hosts.

2.0 Cryptosporidium in fish

Cryptosporidium has been described in both fresh and marine water piscine species with parasitic stages located either on the stomach or intestinal surface, or at both sites (Table 3). The first account of Cryptosporidium in a piscine host was Cryptosporidium nasorum, identified in a

Naso tang, a tropical fish species (Hoover et al., 1981). Hoover and colleagues also noted a similar infection in an unnamed species of marine fish (Hoover et al., 1981). Levine (1984) suggested that

C. nasorum should be given the status of species. However, lack of viable oocyst measurements and taxonomically valid diagnostic features and the fact that only developmental stages on the intestinal microvillous surface were described, has resulted in C. nasorum being considered a nomen nudem

(ie. a name that is invalid because it was published without sufficient description) (Ryan et al.,

2004; Xiao et al., 2004).

Cryptosporidium developmental stages have been identified along the intestinal villi of barramundi (Glazebrook & Campbell, 1987) and carp (Pavlasek, 1983). Intestinal contents confirmed Cryptosporidium infections in brown trout, although this study did not include histological or electron microscopy descriptions (Rush et al., 1987). By contrast, stomach mucosa infections only were seen in fry and fingerling cichlids (predominantly oocysts in necrotic epithelial cells) (Landsberg and Paperna, 1986), red drum (Camus and Lopez, 1996), pearl gourami and cichlids (Paperna and Vilenkin, 1996). Muench and White (1997) described parasite developmental stages (mainly trophozoites, macrogametes and sporozoites) on both the luminal surface of the stomach and the proximal small intestines of cat fish. More recently Cryptosporidium has been detected in two herring sp. in New York state (Ziegler et al., 2007).

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Experimental transmission of C. parvum into rainbow trout revealed no life-cycle stages of

Cryptosporidium in any part of the apical border of the digestive tract upon histological examination of hematoxylin-eosin-stained gastrointestinal sections (Freire-Santos et al., 1998).

However, sections of the stomach and pyloric region from exposed fish displayed large numbers of

5-7-µm spherical structures located deep within the epithelial tissue. Under conditions of host stress, the number of these structures increased remarkably. An immunofluorescence antibody test with IgG and IgM anti-cryptosporidial antibodies revealed fluorescence reactivity in these structures. Simultaneously, wild trout were analyzed in order to detect natural cryptosporidial infections; Cryptosporidium oocyst-like bodies were found in the intestinal content of 10% of the specimens (Freire-Santos et al., 1998).

With fish parasites, transmission and dispersion by water are facilitated by the aquatic habitat of the host and the frequent releasing of fully sporulated oocysts with mucus casts or feces. Recent studies have implicated the brine shrimp Artemia in the transmission of Cryptosporidium in cultured fish as it is the most common live food used in fish and shellfish larviculture (Martinez-Hermida et al., 2007). The viability of Cryptosporidium oocysts ingested by Artemia franciscana metanauplii was evaluated using fluorogenic vital dyes. There was no significant difference (p = 0.09) between the viability of oocysts maintained in saline (control) and those recovered from the digestive tract of the microcrustacean 24 h after ingestion (95 vs 90% viable oocysts) (Méndez-Hermida et al., 2007). The ingestion of other infective stages besides oocysts, favored by frequent cannibalism among fish, could also contribute to the dispersion of the disease. Recirculation systems may also contribute to oocyst concentration and dispersion in aquaculture facilities.

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A recent study examined the risk of exposure to Cryptosporidium based on fish samples and hand wash samples taken from Baltimore urban anglers (Roberts et al., 2007). A total of 46 fish and hand wash samples were collected from several popular angling locations in the Baltimore

Metropolitan area. FISH (fluorescence in situ hybridization) in conjunction with immunofluorescent antibodies (IFA) assay were used for identification and viability assessment of C. parvum oocysts for both the fish and hand wash samples. 10 of 18 (56%) hand samples and 7 of 28 (25%) fish samples were positive for Cryptosporidium. No genotyping was conducted to determine what species or genotypes of Cryptoporidium were present but the authors concluded that the mean probability of infection with Cryptosporidium was relatively high (Roberts et al., 2007).

A new genus for Cryptosporidium-like species infecting a number of piscine hosts, designated Piscicryptosporidium was proposed by Paperna and Vilenkin (1996). The genus included two species, P. reichenbachklinkei and P. cichlidaris (Paperna and Vilenkin, 1996).

Oocysts in the genus Piscicryptosporidium were found in the basal part of the epithelium of the stomach and that endogenous stages were significantly smaller than those found in other host species (Paperna and Vilenkin, 1996). In addition, the author’s reported that the surface of the parasitophorous sac was covered by rudimentary microvilli (Paperna and Vilenkin, 1996).

However, these apparently differential features have also been described in some mammalian

Cryptosporidium spp. For example, C. parvum has been found within some cells occasionally

(Beyer et al., 2000; Marcial and Madara 1986) and microvilli are usually retained in different mammalian species (Alvarez-Pellitero and Sitja-Bobadilla, 2002). Recently, P. reichenbachklinkei and P. cichlidaris were published as Cryptosporidium reichenbachklinkei and C. cichlidaris (Jirku et al., 2008), with the comment that the genus was considered a synonym of Cryptosporidium.

However, no molecular data has been provided and until further data is generated, the validity of the genus remains to be determined.

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2.1 Cryptosporidium molnari, Alvarez-Pellitero and Sitja-Bobadilla, 2002

A new species, Cryptosporiduim molnari, has been described in two marine fish, gilthead sea bream (Sparus aurata L.) and European sea bass (Dicentrarchus labrax L.), mainly in the stomach and rarely in the intestines (Alvarez-Pellitero and Sitja-Bobadilla, 2002) (Table 3).

Histological investigation revealed mature (fully sporulated) oocysts within the stomach epithelium, lumen and faeces. All developmental stages were located in host cell-associated parasitophorous vacuoles bordered by a distinct double host microvillar membrane. The near spherical oocysts were small (3.23 – 5.45 x 3.02 – 5.04 µm), and contained four sporozoites.

Only gilthead sea bream with a high intensity of infection exhibited clinical signs, consisting of whitish faeces, abdominal swelling and ascites. Histopathological examination revealed that while meronts and gamonts in extracytoplasmic position apparently produced no harm to the fish tissue, zygotes and oocysts produced a massive necrosis of epithelial cells (Alvarez-Pellitero and

Sitja-Bobadilla, 2002). Mortalities associated with C. molnari were also reported in some fingerling stocks (Alvarez-Pellitero and Sitja-Bobadilla, 2002). Cryptosporidium molnari oogonial and sporogonial stages were located deep within the epithelium of the infected fish, a characteristic differing from all previously described species of Cryptosporidium (Alvarez-Pellitero and Sitja-

Bobadilla, 2002).

Stomach mucosal scrapings obtained from donor gilthead sea bream infected with C. molnari were inoculated per os to gilthead sea bream and sea bass, resulting in successful infections

(Alvarez-Pellitero and Sitja-Bobadilla, 2002). Sea bass also became infected by cohabitation with infected gilthead sea bream (Alvarez-Pellitero and Sitja-Bobadilla, 2002). No genetic data has been made available for C. molnari, although the authors reported that monoclonal antibodies against C. parvum did not cross-react with C. molnari (Alvarez-Pellitero and Sitja-Bobadilla, 2002).

More recently, a long-term epidemiological study of C. molnari in aquacultured European sea bass and gilthead sea bream was performed in different types of facilities on the Atlantic,

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Cantabric, and Mediterranean coasts (Sitja-Bobadilla et al., 2005). Cryptosporidium molnari was more prevalent in gilthead sea bream than in European sea bass. A seasonal distribution of C. molnari was found in gilthead sea bream, with maximum prevalence and intensity occurring in spring, followed by summer (Sitja-Bobadilla et al., 2005). The highest infection levels were observed in pre-ongrowing and early ongrowing fish, with a tendency to decrease with fish weight (peak in 30- to 100-g fish).

There was a significant relationship between the presence of the parasite and the production step; i.e. the prevalence of infection was higher in fish recently introduced in sea cages and in preongrowing systems and fish were almost never infected before entering the postlarval and nursery facilities. This implies that postlarval or nursery stage is where the fish become infected and suggests that the fish became infected through the water in production steps with less stringent water treatment (Sitja-Bobadilla et al., 2005).

2.2 Cryptosporidium scophthalmi, Alvarez-Pellitero et al., 2004

A second picine species, C. scophthalmi was described in the farmed turbot (Scophthalmus maximus), sampled from different farms on the coast of NW Spain (Alvarez-Pellitero et al., 2004).

In contrast to C. molnari, the parasite was found mainly in the intestinal epithelium and very seldom in the stomach. Oocysts were almost spherical and measured 3.7-5.03 x 3.03-4.69 µm

(mean 4.44 x 3.91) (shape index 1.05-1.34, mean 1.14) (Alvarez-Pellitero et al., 2004). As with C. molnari, sporogonial stages were observed deeply embedded within the epithelium. Infection prevalence was very variable, and juvenile fish were most frequently and intensively parasitised.

Epidemiological data obtained for C. scophthalmi over a period of several years have

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shown a high prevalence and intensity of infection in small fish, with a sharp decrease in

larger animals (Alvarez-Pellitero et al., unpublished). Parasitized fish did not exhibit external clinical signs, although some fish showed intestinal distension at necropsy. Severe infection resulted in marked histopathological damage including distension of epithelial cells by large vacuoles, containing clusters of oocysts, which can lead to sloughing of epithelial cell remnants and oocysts or even detachment of intestinal mucosa. An inflammatory reaction involving leucocyte infiltration was sometimes observed) (Alvarez-Pellitero et al., 2004).

As with C. molnari, no genetic data has been made available for C. scolphthalmi and the provision of samples for molecular analysis by other laboratories has not been forthcoming. This makes it impossible to determine the phylogenetic relationships between C. molnari and other

Cryptosporidium species.

2.3 The guppy genotype

At present, the only genetic data that is available for picine Cryptosporidium species is from a guppy. Histological and morphological characteristics of the guppy isolate were similar to C. molnari. These common characteristics included histological damage (necrotic epithelium), sloughing epithelial cells detaching from the lumen and plentiful cell debris with minimal inflammation response, parasitic stages dominant in the stomach, lack of epithelial infection in the intestinal mucosa, similar oocyst measurements and most importantly the proposed feature distinct to C. molnari, oogonial and sporogonial stages deep within the epithelium.

Phylogenetic analysis of the actin and small subunit (SSU) rRNA loci of the guppy isolate showed that it was genetically distinct and grouped separately from gastric and intestinal species of

Cryptosporidium. Therefore it is plausible that piscine species of Cryptosporidium may represent the most primitive Cryptosporidium species, part of the genus dating back prior to the divide into gastric and intestinal specific forms (Ryan et al., 2004).

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3.0 Cryptosporidium in amphibians.

The first report of Cryptosporidium in amphibians was the description of Cryptosporidium oocysts in the faeces of a captive mice-fed ornate horned frog, Ceratophrys ornata (Crawshaw and

Mehren, 1987). The identity of the species is unknown and as mice can be infected with several species and genotypes of Cryptosporidium, it remains unknown if the frog was naturally infected or passing oocysts from infected mice. Both natural and experimental cryptosporidiosis in giant toads, Rhinella marina (previously Bufo marinus) have also been described (Arcay and Bruzal

1993, Arcay et al., 1995). However, as with the previous publication, the identity of

Cryptosporidium isolate used in the experiments is unclear and the cryptosporidiosis in R. marina thus needs further investigation and re-evaluation. Attempts to transmit C. parvum and C. serpentis to amphibians were not successful (Graczyk et al., 1996, 1998). Cryptosporidium has also been described from the gastric mucosa in a single laboratory reared African clawed frog (Xenopus laevis), causing proliferative gastritis (Green et al., 2003). The 2-year-old female frog was euthanized because of chronic weight loss. At necropsy, there was no evidence of bacterial, fungal or viral disease; however, the histopathologic findings indicated a proliferative gastritis and the presence of numerous cryptosporidial stages throughout the intestinal tract. Developmental stages were confirmed using TEM and oocysts were also recovered from the sediment in the aquarium.

The affected frog had been laboratory-reared by a commercial producer and kept in a laboratory facility in the USA with ~2,000 individuals, but no other frog in the facility was affected. No genetic characterization was conducted and the identity of the species remains unknown.

3.1 Cryptosporidium fragile, Jirku et al., 2008

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Recently a new species, Cryptosporidium fragile was described from the stomach of naturally infected black-spined toads (Duttaphrynus melanostictus) from the Malay peninsula (Jirku et al., 2008). The parasitized animals were directly imported from Malaysia and harboured C. fragile at the time of arrival. A total of 18 animals were examined and 15 were infected with C. fragile (83% prevalence). Oocysts were subspherical to elliptical with irregular contour in optical section, measuring 6.2 (5.5-7.0) x 5.5 (5.0-6.5) µm. The oocysts of C. fragile were comparable in size with C. serpentis but were variable in shape. The oocyst wall was smooth and colourless in light microscopy. Oocysts were extremely sensitive to hypertonic conditions, crumpling immediately when exposed to flotation solutions and spontaneously disintegrated after 4 weeks of storage in water at 4°C, 10°C and 20°C. The species was named C. fragile to reflect the fragile nature of the oocysts (Jirku et al., 2008).

The endogenous development of C. fragile in the stomach of black-spined toad was analysed in detail using light and electron microscopy. Developmental stages were confined to the surface of gastric epithelial cells. No developmental stages were found in other examined tissues.

In transmission experiments, C. fragile was not infective for one fish species (Poecilia reticulate), four amphibian species (Bufo bufo, Rana temporaria, Litoria caerulea, X. laevis) one species of reptile (Pantherophis guttatus) and SCID mice. Phylogenetic analysis of the full length

18S rRNA revealed C. fragile to be genetically distinct and grouped with the gastric species. It appeared to be most closely related to an avian genotype from a Eurasian woodcock (AY273769),

C. serpentis and reptile isolates from a star tortoise (AY120914) and leopard gecko (AY120915).

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Table 1. Morphometric comparisons of the oocysts from Cryptosporidium meleagridis,

Cryptosporidium baileyi, Cryptosporidium galli and avian Cryptosporidium genotypes for which measurements are available.

Oocyst Species/genotype Oocyst width Length/width length Reference µm ratio µm C. meleagridis 4.5 - 6.0 4.2 - 5.3 1.00 - 1.33 Lindsay et al., 1989 C. baileyi 6.0 - 7.5 4.8 - 5.7 1.05 - 1.79 Lindsay et al., 1989 C. galli 8.0 - 8.5 6.2 - 6.4 1.3 Ryan et al., 2003 Meireles et al., 2006; Avian genotype IIa 6.0 - 6.5 4.8 - 6.6 1 - 1.25 Ng et al., 2006 Avian genotype III 7.5 6.0 1.25 Ng et al., 2006 Avian genotype IV 8.25 6.3 1.30 Ng et al., 2006 Eurasian woodcock genotype 8.5 6.4 1.32 Ryan et al., 2003

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Table 2 Avian Cryptosporidium genotypes and their GenBank accession numbers for various loci.

SSU Geographic Actin HSP-70 Genotype Host species rRNA Reference origin Locus locus Locus Avian Red Factor Canary DQ650 Ng et al., Genotype WA DQ650339 (Serinus canaria) 346 2006 I Eclectus (Eclectus roratus), Galah (Eolophus DQ650340 DQ650 roseicapilla), , 347, Meireles et Avian Cockatiel DQ650341 DQ650 al., 2006, Genotype (Nymphicus WA, Brazil DQ002929 , 348, Ng et al., II hollandicus), Major DQ002931 DQ002 2006 Mitchell Cockatoo 930 (Cacatua leadbeateri, Ostrich (Struthio camelus) Galah (Eolophus roseicapilla), DQ650 Avian Cockatiel DQ650342 349, Ng et al., Genotype (Nymphicus WA , NA DQ650 2006 III hollandicus), Sun DQ650343 350 Conure (Aratinga solstitialis) Avian Japaneese whiteeye Czech Ng et al., Genotype (Zosterops DQ650344 NA NA Republic 2006 IV japonica) Eurasian woodcock Ryan et al., Eurasian Czech DQ650 (Scolopax AY273769 AY273773 2003, Ng et woodcock Republic 345 rusticola) al., 2006 Morgan et black duck (Anas AF316630, al., 2001, Duck rubripes), Canada Australia, AY504514 Zhou et al., NA NA genotype Geese (Branta USA , 2004, canadensis) AY324639 Jellison et al., 2004 AY120912 , Xiao et al., Canada Geese AY504513 2002, Zhou Goose AY120 (Branta USA , NA et al., 2004, genotype I 929 canadensis) AY504516 Jellison et , al., 2004 AY324642 AY504512 Zhou et al., Goose Canada Geese , 2004, genotype (Branta USA AY504515 NA NA Jellison et II canadensis) , al., 2004 AY324643 Canada Geese Goose (Branta Jellison et genotype USA AY324638 NA NA canadensis) - al., 2004 III isolate KLJ-3b Canada Geese Goose (Branta Jellison et genotype USA AY324641 NA NA canadensis)- isolate al., 2004 IV KLJ-7

NA = Not available

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Table 3. Cryptosporidium sp. reported in piscine hosts

Species Host Site of Size (µm) Reference

Infection L x W

C. nasorum Naso literatus Intestine 3.6 x - (Hoover et al.,

1981)

Cryptoporidium sp. Cyprinus carpio Intestine - Pavlasek, 1983 Cryptoporidium sp. Oreochromis sp. Stomach - Landsberg and Paperna, 1986 Cryptoporidium sp. Salmo trutta Intestine - Rush et al., 1987 Cryptoporidium sp. Lates calcarifer Intestine - Glazebrook and Campbell, 1987 Cryptoporidium sp. Oncorhynchus Stomach 5-7 Freire-Santos et mykiss al., 1998 Cryptoporidium sp. Sciaenops Stomach 7 x 4 Camus and Lopez, ocellatus 1996 Cryptoporidium sp. Plecostomus sp. Intestine and - Muench and Stomach White, 1997 Piscicryptosporidium Trichogaster leeri Stomach 2.4-3.18 x 2.4- (Paperna and reinchenbachklinkei 3.0 Vilenkin, 1996) Piscicyrptosporidium Oreochromis sp. Stomach 4.0-4.7 x 2.5- (Paperna and cichlidis (previously 3.5 Vilenkin, 1996) Cryptocystidium villithecum) Piscicyrptosporidium Sparus auratus Stomach 3.1 x - (Paperna and sp. (previously Vilenkin, 1996) Chloromyxum-like) C. molnari Sparus aurata, Stomach 3.23-5.45 x Alvarez-Pellitero Dicentrarchus 3.02-5.04 and Sitja- labrax Bobadilla, 2002 C. scophthalmi Scophthalmus Intestine 3.7-5.03 x Alvarez-Pellitero maximus 3.03-4.69 et al., 2004 C. molnari-like sp. Poecilia reticulate Stomach 4.6 by 4.4 Ryan et al., 2004 Cryptosporidium sp. Alosa - - Ziegler et al., 2007 pseudoharengus

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