Description of Cryptosporidium ornithophilus n. sp. (Apicomplexa: Cryptosporidiidae) in farmed ostriches

Background Avian cryptosporidiosis is a common parasitic disease that is caused by five species, which are well characterised at the molecular and biological level, and more than 18 genotypes for which we have limited information. In this study, we determined the occurrence and molecular characteristics of Cryptosporidium spp. in farmed ostriches in the Czech Republic. Methods The occurrence and genetic identity of Cryptosporidium spp. were analysed by microscopy and PCR/sequencing of the small subunit rRNA, actin, HSP70 and gp60 genes. Cryptosporidium avian genotype II was examined from naturally and experimentally infected hosts and measured using differential interference contrast. The localisation of the life-cycle stages was studied by electron microscopy and histologically. Infectivity of Cryptosporidium avian genotype II for cockatiels (Nymphicus hollandicus (Kerr)), chickens (Gallus gallus f. domestica (L.)), geese (Anser anser f. domestica (L.)), SCID and BALB/c mice (Mus musculus L.) was verified. Results A total of 204 individual faecal samples were examined for Cryptosporidium spp. using differential staining and PCR/sequencing. Phylogenetic analysis of small subunit rRNA, actin, HSP70 and gp60 gene sequences showed the presence of Cryptosporidium avian genotype II (n = 7) and C. ubiquitum Fayer, Santín & Macarisin, 2010 IXa (n = 5). Only ostriches infected with Cryptosporidium avian genotype II shed oocysts that were detectable by microscopy. Oocysts were purified from a pooled sample of four birds, characterised morphometrically and used in experimental infections to determine biological characteristics. Oocysts of Cryptosporidium avian genotype II measure on average 6.13 × 5.15 μm, and are indistinguishable by size from C. baileyi Current, Upton & Haynes, 1986 and C. avium Holubová, Sak, Horčičková, Hlásková, Květoňová, Menchaca, McEvoy & Kváč, 2016. Cryptosporidium avian genotype II was experimentally infectious for geese, chickens and cockatiels, with a prepatent period of four, seven and eight days post-infection, respectively. The infection intensity ranged from 1000 to 16,000 oocysts per gram. None of the naturally or experimentally infected birds developed clinical signs in the present study. Conclusions The molecular and biological characteristics of Cryptosporidium avian genotype II, described here, support the establishment of a new species, Cryptosporidium ornithophilus n. sp.

Unlike C. baileyi, which infects a broad range of birds from different orders, many recently described Cryptosporidium species and genotypes appear to have a relatively narrow host range. For example, Cryptosporidium avian genotype VI appears to be restricted to North American red-winged blackbirds [8], and Cryptosporidium goose and duck genotypes have been found only in anseriform birds [11,15]. Similarly, C. avium and Cryptosporidium avian genotype I are almost exclusively found in psittacines and passerines, respectively [5][6][7]19]. Cryptosporidium avian genotype II has been found predominantly in ostriches but also in other species within the order Struthioniformes as well as orders Galliformes and Psittaciformes (Table 1).
In the present study, we report on the occurrence of Cryptosporidium spp. in farmed ostriches. For the most prevalent genotype in ostriches, Cryptosporidium avian genotype II, we further describe oocyst morphometry, experimental host specificity, developmental stage localization and molecular characteristics. Based on the collective data from this and previous studies, we conclude that Cryptosporidium avian genotype II is genetically and biologically distinct from the species of Cryptosporidium considered valid, and propose the name Cryptosporidium ornithophilus n. sp. for this genotype.

Specimens studied
Faecal samples were collected from ostriches on four farms in the Czech Republic. Faecal samples from juvenile (aged 9-12 months) and adult (older than three years) ostriches were individually collected into sterile plastic vials and stored at 4-8 °C until subsequent processing. Faecal smears were prepared from each sample, stained with aniline-carbol-methyl violet (ACMV), and examined for the presence of Cryptosporidium spp. oocysts [37]. Faecal samples were also screened for the presence of Cryptosporidium-specific DNA by PCR/ sequencing (described below). Oocysts of C. ornithophilus n. sp. were purified from pooled faecal samples from a naturally infected juvenile common ostrich (no. 43588, Struthio camelus L.) kept on the farm number 4 using caesium chloride gradient centrifugation [38]. Purified oocysts were used for morphometry and preparation of the inoculum. The propidium iodide (PI) staining was used for test of oocysts viability [39]. Cryptosporidium ornithophilus n. sp. oocysts from a common ostrich were pooled and used to infect a single one-day-old chickens (chicken 0; Gallus gallus f. domestica). Oocysts recovered from the faeces of chicken 0 were used to infect other experimental animals. The purity of C. ornithophilus n. sp. isolate before performing the experimental infection and taking the measurements, and during the experiments was verified by the following procedure. The sequence of the original isolate (ostrich) was compared to the sequence obtained from chicken 0 and from tissue specimens and faecal samples of experimentally inoculated animals (below). The oocyst size of the original isolate was compared with isolates obtained from susceptible hosts.

Oocyst morphometry
Oocysts of C. ornithophilus n. sp. from naturally and experimentally infected hosts (50 oocysts from each isolate) were examined and length and width measurements were taken using differential interference contrast (DIC) at 1000× magnification. All measurements are in micrometres and are given as the range followed by the mean ± standard deviation (SD) in parentheses. These measurements were used to calculate the lengthto-width ratio. Sample containing purified C. parvum Tyzzer, 1912 oocysts from a naturally infected Holstein calf was used as a size control (n = 50). Size of oocysts was measured using the same microscope and by the same person. Each slide was screened a meandering path to prevent repeated measurement of an oocyst. Additionally, different staining methods were used for visualisation of oocysts. Faecal smears with C. ornithophilus n. sp. and C. parvum (data not shown) oocysts were stained by ACMV, modified Ziehl-Neelsen [ZN; 40], phenol staining [AP; 41] and labelled with genus-specific FITC-conjugated antibodies (IFA; Cryptosporidium IF Test, Crypto cel, Cellabs Pty Ltd., Brookvale, Australia). Morphometry was determined using digital analysis of images (Olympus cellSens Entry 2.1 software and Olympus Digital Colour camera DP73, Olympus Corporation, Shinjuku, Tokyo, Japan). Photomicrographs of C. ornithophilus n. sp. oocysts observed by DIC, ACMV, ZN, AP and IFA were stored at the Institute of Parasitology, Biology Centre of the Czech Academy of Sciences, Czech Republic.

Molecular analyses
Total genomic DNA was extracted from 20,000 purified oocysts, 200 mg of faeces, or 200 mg of tissue by bead disruption for 60 s at 5.5 m/s using 0.5 mm glass beads in a FastPrep ® 24 Instrument (MP Biomedicals, CA, USA) followed by isolation/purification using Exgene TM Stool DNA mini (GeneAll Biotechnology Co. Ltd, Seoul, Korea) or DNeasy Blood & Tissue Kit (Qiagen, Hilden, Germany) in accordance with the manufacturer's instructions. Purified DNA was stored at − 20 °C. A nested PCR approach was used to amplify a partial region of the small subunit (SSU) rRNA [42,43], actin [44], 70 kilodalton heat-shock protein (HSP70) [45] and gp60 [46][47][48] genes. The PCR conditions were slightly modified, for more details see [6]. Molecular grade water and DNA of C. parvum were used as negative and positive controls, respectively. Secondary PCR products were detected in 1.5% agarose gel stained with ethidium bromide. PCR products were cut out from gel, purified using Gen Elute Gel Extraction Kit (Sigma, St. Louis, MO, USA) and sequenced in both directions with an ABI 3130 genetic analyser (Applied Biosystems, Foster City, CA) using the secondary PCR primers in commercial laboratory (SEQme, Dobříš, Czech Republic).

Phylogenetic analyses
The nucleotide sequences obtained in this study were edited using the ChromasPro 2.4.1 software (Technelysium, Pty, Ltd., South Brisbane, Australia) and aligned with reference sequences downloaded from GenBank using MAFFT version 7 online server (http://mafft .cbrc. jp/align ment/softw are/). The most appropriate evolutionary models for phylogeny analyses and values of all parameters for each model were selected using the MEGAX software [49,50]. The evolutionary history was inferred by using the Maximum Likelihood (ML) method based on the Tamura 3-parameter model [51] selected for SSU and HSP70 alignments and the general time reversible model [52] was selected for actin alignment.

Animals for transmission studies
Five adult cockatiels (Nymphicus hollandicus (Kerr)), five one-day-old chickens, five one-day-old geese (Anser anser f. domestica L.), five seven-day-and eight-weekold SCID mice (Mus musculus; strain C.B-17) and five seven-day and eight-week-old BALB/c mice were used for transmission studies. Three adult cockatiels, chickens, geese and seven-day and eight-week-old SCID and BALB/c mice used as a negative control. As a control, the infectivity of C. parvum from a naturally infected Holstein calf for three adult cockatiels, chickens, geese and seven-day and eight-week-old SCID and BALB/c mice was verified. All animals, except chickens, geese and seven-day-old mice, which were hatched under laboratory conditions, were screened every other day for the presence of oocysts of Cryptosporidium spp. and specific DNA two weeks prior to transmission studies. Cockatiels originated from breeders located in the Czech Republic and laboratory mice were obtained from Charles River (Germany).

Animal care
Rodents were individually housed in ventilated cages (Tecniplast, Buguggiate, Italy). Chickens and geese were housed in boxes and cockatiels were kept in separate aviaries. The size of boxes and aviaries were according to regulated by Czech legislation (Act No 246/1992 Coll., on protection of animals against cruelty). An external source of heat was used in the first five days for chickens and geese. Sterilized diet and water were available for all animals ad libitum. Animal caretakers wore sterile shoe covers and disposable coveralls and disposable gloves always they entered the experimental room. Wood-chip bedding and disposable protective clothing were removed from the experimental room and incinerated.

Experimental design
A total 20,000 purified oocysts of C. ornithophilus n. sp., suspended in 10 µl of distilled water, were dropped into the mouth/beak of each animal. Animals serving as negative controls were inoculated orally with 10 µl of distilled water. Faecal samples from all animals were screened daily for the presence of Cryptosporidium oocysts using ACMV staining and the presence of Cryptosporidium-specific DNA was confirmed using nested PCR/ sequencing targeting the SSU gene. All experiments were terminated 30 days post-infection (dpi). Infection intensity was reported as the number of oocysts per gram (opg) of faeces, as previously described by Kváč et al. [53].
In addition, faecal consistency and colour and general health status were examined daily. The sequence identity of the Cryptosporidium DNA recovered from infected hosts to inoculum and original isolate at SSU, actin and HSP70 was verified in each experimentally infected animal.

Histopathological and scanning electron microscopy (SEM) examinations
Two animals from each group (at 10 and 20 dpi) were examined at necropsy. Tissue samples from oesophagus; stomach in rodents and proventriculus and ventriculus in birds; duodenum; jejunum (proximal, central and distal); ileum; caecum and colon were collected for histology and SEM followed by processing according [6]. Slides for histology were examined at 100-400× magnification and documented using Olympus cell Sens Entry 2.1 (Olympus Corporation, Shinjuku, Tokyo, Japan) equipped with a digital camera (Olympus DP73). Samples for SEM were examined using a JEOL JSM-7401F-FE SEM and documented using ETD Detector A PRED (Thermo Fisher Scientific, Waltham, MA, USA). Additionally, DNA from tissue samples was isolated and the sequence identity to inoculum and original isolate at SSU, actin and HSP70 was verified.

Staining of mucosal smears
Wright staining procedures were used to visualize Cryptosporidium spp. developmental stages in the gastrointestinal tract of chickens [54]. Tissue samples of the large intestine (selected on the basis of histological examination) were washed with cold PBS with subsequent exposure to serum from Cryptosporidium-negative chickens for five min. The mucous membrane was gently scrapped with a scalpel and smeared on a glass slide. Wet mucosal smears were fixed with osmium vapour for 15 min followed by Wright staining for 6 min. Slides were viewed at 1000× magnification and documented using Olympus cell Sens Entry 2.1 (Olympus Corporation, Shinjuku, Tokyo, Japan) equipped with a digital camera (Olympus DP73).

Statistical analysis
Differences in Cryptosporidium spp. oocysts size were tested using Hotellingʼs multivariate version of the 2 sample t-test, package ICSNP: Tools for Multivariate Nonparametrics in R 4.0.0. [55]. The hypothesis tested was that two-dimensional mean vectors of measurement are the same in the two populations being compared.

Results
A total of 164 juvenile and 40 adult ostriches were screened for the presence of Cryptosporidium infection. Cryptosporidium spp. was detected on three out of four ostrich farms. Out of 204 faecal samples, five (2.5%) were microscopically positive for the presence of Cryptosporidium oocysts and 12 (5.9%) contained specific DNA of Cryptosporidium spp. (Table 2). All microscopically positive samples were also positive for Cryptosporidium DNA. Only juvenile ostriches (n = 12) were infected with Cryptosporidium spp. Screened animals had good health and faecal consistency appropriate to the age of birds and feeding.
All birds positive for Cryptosporidium-specific DNA were successfully genotyped by sequence analysis of SSU and actin genes ( Table 2). ML trees constructed from SSU and actin sequences in this study showed the presence of C. ubiquitum Fayer, Santín & Macarisin, 2010 (n = 5) and C. ornithophilus n. sp. (n = 7; Table 2, Figs. 1, 2). HSP70 gene sequences were successfully amplified only from samples positive for C. ornithophilus n. sp. (Fig. 3). The C. ubiquitum gp60 gene was amplified and sequenced from five positive DNA samples from farm no. 1 ( Table 2, Fig. 4). Sequences were identical to each other and clustered with subtype family XIIa (Fig. 4). Out of seven ostriches positive for C. ornithophilus n. sp., five shed microscopically detectable oocysts (6000-18,000 opg, Table 2). Birds positive for C. ubiquitum DNA did not shed oocysts detectable by microscopy.
Cryptosporidium ornithophilus n. sp. oocysts did not infect 7-day-old and 8-week-old BALB/c or SCID mice, whereas 7-day-old BALB/c and both age categories of SCID mice were infected with C. parvum (control group, data not shown). All chickens, geese and cockatiels inoculated with oocysts of C. ornithophilus n. sp. developed infections. Oocysts or specific DNA were first detected at 4 dpi, 7 dpi and 8 dpi in geese, chickens and cockatiels, respectively (Fig. 5). The infection intensity ranged from 2000 to 16,000 opg in chickens and cockatiels and from 1000 to 8000 opg in geese (Fig. 5).
Molecular, histological and SEM analyses and examination of stained mucosal smears of gastrointestinal tract tissue in birds with C. ornithophilus n. sp. showed  the presence of developmental stages only in the caecum and colon of chickens and geese sacrificed 10 and 20 dpi (Figs. 6, 7). Few developmental stages were detected on each villus (Figs. 6, 7). Developmental stages were not detected in cockatiels, but specific DNA was detected exclusively in the caudal part of the ileum.
The morphometry of the developmental stages of C. ornithophilus n. sp. was examined in preparations with Wright's stain ( Table 3). Most of the detected developmental stages were enveloped by a parasitophorous sac, which appeared as an unstained halo (Fig. 8). A large number of oocysts was detected, and most were unstained with sporozoites not visible (Fig. 8). We were not able to differentiate between thin-and thick-walled oocysts. Free sporozoites were not detected, but a photomicrograph of sporozoites following oocyst excystation is included in Fig. 8. Mononuclear trophozoites were the most frequently observed developmental stage which also showed a high variability in size ( Fig. 8; Table 3). Type I meronts, containing 8 merozoites, were observed frequently (Fig. 8), while Type II meronts, with 4 merozoites, were found rarely (Fig. 8). Free merozoites were found rarely (Fig. 8). Microgamonts were found rarely (Fig. 8), but macrogamonts, typified by a number of amylopectin granules in their cytoplasm and a foam-like appearance, were frequently observed (Fig. 8). Zygotes were lightly stained compared to the unstained oocysts (Fig. 8).
SSU, actin and HSP70 sequences obtained from the original isolate of C. ornithophilus n. sp. (ostrich) were identical to isolates recovered from faeces of chicken 0 and all other birds infected during the whole experiment. Additionally, sequences obtained from the tissue specimens of caecum and colon of chickens and geese and in the ileum of cockatiels were also identical to the inoculum. The gene encoding gp60 was not successfully amplified in any animal experimentally infected with C. ornithophilus n. sp., indicating the absence of C. ubiquitum or other species and genotypes of Cryptosporidium spp. (e.g. C. parvum) that could be part of the inoculum.
The above data tend to justify the distinct status of Cryptosporidium ornitophilus n. sp., which is described below.

Discussion
Birds are naturally parasitized with several Cryptosporidium species and genotypes [16,18]. Here, we reported the occurrence of Cryptosporidium spp. in ostriches farmed commercially and described Cryptosporidium avian genotype II as a new species. Previous studies have shown that ostriches are frequently infected with C. baileyi [32][33][34] and C. ornithophilus n. sp. [19,36,59]; however, we detected C. ornithophilus n. sp. and C. ubiquitum. The absence of C. baileyi could be explained by the age of the birds screened in the present study. Previous studies  reported C. baileyi in ostriches younger than 3 months with older birds being infected rarely or not at all [32,34]. In this study, the occurrence of C. ornithophilus n. sp. in birds aged 9-14 months was 4.3% (7/164), which is similar to that reported in Vietnamese ostriches older than 12 months (5.8%; [36]). The absence of C. ornithophilus n. sp. in birds older than three years in this study could be due to age-related resistance or immunity, as described for C. baileyi, C. avium, C. parvum, C. muris and C. andersoni Lindsay, Upton, Owens, Morgan, Mead & Blagburn, 2000 in various hosts [60][61][62], but this needs to be examined experimentally. Cryptosporidium ubiquitum is not typically found in birds so our finding of five ostriches on a single farm  [63] also detected C. ubiquitum in birds (common hill mynas, Gracula religiosa L.) at commercial markets in China. It is possible that the detected DNA was due to mechanical passage, not an active infection. The cohabitation of livestock, companion and wild animals can result in Cryptosporidium oocyst passage through non-susceptible animals without establishing infection [64][65][66]. We cannot exclude that some wild animals may be the source of C. ubiquitum. Our failure to detect oocysts also suggests that any infection was likely to be of low intensity.
Phylogenetic analyses based on SSU, actin and HSP70 gene sequences showed that C. ornithophilus n. sp. is genetically distinct from known species and is most closely related to C. baileyi and C. avium. At the SSU locus, C. ornithophilus n. sp. shares 92.8% and 93.5% similarity with C. baileyi and C. avium, respectively. This is comparable to the similarity between C. andersoni and C. ryanae (91.1%) or C. muris and C. suis (93.3%). At the actin locus, similarities with C. baileyi and C. avium are 88.7% and 98.1%, respectively. In comparison, C. bovis and C. ryanae share 88.1% similarity and C. parvum and C. erinacei share 98.3% similarity at the actin locus. At the HSP70 locus, C. ornithophilus n. sp. shares 91.3% and 95.6% similarity with C. baileyi and C. avium, respectively. In comparison, C. parvum and C. erinacei share 99.2% similarity at the HSP70 locus.