Parasitologists United Journal

: 2014  |  Volume : 7  |  Issue : 1  |  Page : 47--55

Comparison between human and fish species of Cryptosporidium and Cyclospora

Mona M El Temsahy1, Eman D El Kerdany1, Radwa G Diab1, Maha R Gaafar1, Iman H Diab2,  
1 Department of Parasitology, Faculty of Medicine, Alexandria University, Alexandria, Egypt
2 Department of Biochemistry, Faculty of Medicine, Alexandria University, Alexandria, Egypt

Correspondence Address:
Maha R Gaafar
MD, Departments of Parasitology, Faculty of Medicine, Alexandria University, Alexandria


Background Cryptosporidium and Cyclospora spp. are worldwide emerging parasites in both immunocompetent and immunocompromised individuals. Objective This study was designed to compare Cryptosporidium and Cyclospora spp. detected in fish with the corresponding species isolated from humans, morphologically and genetically. Detection of any similarity could be considered of potential epidemiological importance. Materials and methods Intestinal contents of 35 Tilapia zillii fish and 50 human stool samples were stained and examined to identify Cryptosporidium and Cyclospora oocysts. Thirty male Swiss albino mice were divided into the control group (I) and the experimental group (II), which was further subdivided into four equal subgroups (IIa, IIb, IIc, and IId), that were infected with fish and human Cryptosporidium and Cyclospora spp., respectively. Two weeks later, all mice were killed; parts of their small intestines were subjected to histopathological examination and processed for transmission electron microscopy (TEM). DNA was extracted from frozen oocysts present in human stool samples and fish intestinal samples, and amplification was performed using specific primers for Cryptosporidium parvum and Cyclospora cayetanensis. Results Cryptosporidium and Cyclospora spp. of both fish and humans detected in mice intestinal sections were morphologically similar by both light and electron microscope. However, failure of DNA amplification of oocysts of both parasites in fish intestinal samples, following application of the specific primers, indicates that fish species were not identical to human species. Conclusion It can be deduced that species identified in fish are apparently not infectious to humans.

How to cite this article:
El Temsahy MM, El Kerdany ED, Diab RG, Gaafar MR, Diab IH. Comparison between human and fish species of Cryptosporidium and Cyclospora.Parasitol United J 2014;7:47-55

How to cite this URL:
El Temsahy MM, El Kerdany ED, Diab RG, Gaafar MR, Diab IH. Comparison between human and fish species of Cryptosporidium and Cyclospora. Parasitol United J [serial online] 2014 [cited 2023 Nov 29 ];7:47-55
Available from:

Full Text


Cryptosporidium and Cyclospora spp. are worldwide emerging parasites that cause acute, self-limiting diarrhea in immunocompetent individuals, and pronounced chronic diarrhea in immunocompromised patients. Hundreds of human infections have been reported, including epidemics in several urban areas [1],[2],[3] . Cryptosporidium spp. affects the intestinal tracts of mammals, birds, reptiles, and amphibians as well as fish [2] . There is a great diversity of Cryptosporidium spp. recorded from all these hosts [3],[4] . In 1995, Cryptosporidium oocysts recovered from human stool, referred to as Cryptosporidium parvum, were found to be infectious for fish, amphibians, and reptiles. Moreover, oocysts recovered from C. parvum-inoculated animals were infectious to mice [5] . Considerable information is available for avian cryptosporidiosis, which was not transmissible to mammals [6] . However, scanty information is available for Cryptosporidium spp. infections in fish, amphibians, and reptiles [6],[7] .

In the last decade, Cyclospora cayetanensis has gained prominence as a water-borne and food-borne pathogen. Their oocysts are shed unsporulated with the feces of infected individuals. Therefore, the infections are unlikely to be passed directly [8],[9] . A previous study considered human to be the only host for Cyclospora spp. with the exception of some monkeys and baboons [10] . Meanwhile, other studies reported its presence in rodents, poultry, and reptiles principally snakes [11],[12],[13] . Furthermore, Cyclospora spp. was demonstrated in aquatic animals such as freshwater clams and bivalves [14],[15] . Eventually in 2007, Cyclospora spp. was reported in a woman consuming nonpotable water and raw fish. This suggested that the raw fish may be the source for the infection [16] . To date, there is no convincing evidence that Cyclospora spp. is a zoonotic infection. The present study was undertaken to compare, morphologically and genetically, Cryptosporidium and Cyclospora spp. detected in fish with the corresponding species isolated from infected humans. Detection of any similarity could be considered of potential epidemiological importance.

 Materials and methods

Type of study

This is a descriptive comparative study.

Collection of parasites from fish and human samples

Thirty-five Tilapia zillii were obtained from fish markets of different districts of Alexandria from July 2012 to August 2012. The fish intestinal contents were prepared [17] , and intestinal smears were stained with modified Ziehl-Neelsen, safranin, and modified trichrome stain [18] . Mixed infections were discarded and all single-positive samples for Cryptosporidium and Cyclospora spp. detected in fish were pooled separately. Stool samples were collected from 50 patients suffering from gastrointestinal problems, attending the outpatient clinic of Alexandria University Hospital from July 2012 to August 2012. All samples were screened conventionally and stained with the same special stains used for fish samples [18] . Positive Cryptosporidium or Cyclospora spp. samples were pooled separately. One pooled sample of each parasite from both fish and human was suspended in 4 ml PBS. Each pooled sample was divided into 2 ml; the first was preserved in 2.5% potassium dichromate at 4°C for infecting mice [19] and the second was fixed in an equal volume of glycerol and PBS and stored at −20°C until use for the molecular studies [20] .

According to a pilot study, one pooled sample was selected for each parasite depending on the heaviness of infection and the histopathological intestinal changes in small groups of experimental mice. The parasites were isolated from the fecal debris by repeated washing in PBS according to Lumb's technique [21] .

Infection of experimental mice

Cyclospora oocysts were allowed to sporulate before infection of experimental mice [22] . Just before infection by intragastric gavages [19] , Cryptosporidium oocysts and sporulated Cyclospora oocysts were washed in distilled water, centrifuged, and the dose of infection was adjusted to 10 4 oocysts/0.1 ml for each parasite. Male Swiss albino mice, aged 3-5 weeks, weighing 20-25 g were housed in well-ventilated cages, supplied with standard pellet food and water [23] . Mice stools were examined conventionally to exclude the presence of other parasites [18] . Thirty mice were divided into two main groups. Group I represented the control noninfected group (six mice) and group II representing the experimental infected group (24 mice) was further divided into four equal subgroups: IIa and IIb were infected by Cryptosporidium spp. isolated from fish and humans, respectively; IIc and IId were infected by Cyclospora spp. isolated from fish and humans, respectively. Mice stools were collected every other day, starting a week following inoculation until the day of killing. Smears were stained with modified Ziehl-Neelsen and examined microscopically. Shed oocysts were counted in oil immersion field [24] . Two weeks following infection, all mice were killed by overdose of ether [19] . Their small intestines were divided into two parts. The first was fixed in 10% formalin, stained by hematoxylin and eosin, and subjected to histopathological examination by light microscopy (LM) (HM-LUX; Leitz Wetzlar, Stuttgart, Germany) to detect the parasites and the associated pathological lesions [25] . The second part was fixed in 2.5% glutaraldehyde at 4°C and processed for TEM (100 CX; Joel, Tokyo, Japan) to study the ultrastructural morphology of both fish and human isolates [26] . Fixed oocysts in a mixture of glycerol and PBS were subjected to molecular study for the determination of the genetic characters of both fish and human isolates [20] .

DNA extraction and PCR amplification

DNA was extracted from four positive pooled, preserved, and frozen samples of Cryptosporidium and Cyclospora oocysts (two from fish intestinal contents and two from human stool samples, respectively) using the QIAamp DNA extraction kit (Qiagen, Mississauga, Ontario, Canada). Five μl of the extracted DNA from each sample was loaded on 1% agarose gel stained with ethidium bromide, and the rest was stored at −20°C until use for PCR amplification [27],[28] . Inhibitors present in human stool samples and fish intestinal contents were removed by an agarose-embedded DNA preparation [29],[30],[31] . Genomic DNA (2 μg) was suspended in PCR buffer containing 40 μmol/l of each of two oligonucleotide primers, 200 μmol/l (each) deoxyribonucleoside triphosphates, 50 mmol/l KCl, 10 mmol/l Tris-HCl (pH 8.3), 1.5 mmol/l MgCl 2 , 0.01% gelatin, and 2.5 U Taq DNA polymerase in a total volume of 50 μl [27] . For C. parvum, amplification was performed using two sets of primers; F: 5′-CCGAGTTTGATCCAAAAAGTTACGAA-3′ and R: 5′-TAGCTCCTCATATGCCTTATTGA-GTA-3′. Thermocycling conditions using an automated thermal cycler (PTC-200; MJ Research, California, USA) consisted of denaturation at 94°C for 10 min, followed by 50 cycles of denaturing for 60 s at 94°C, annealing for 90 s at 56°C, and extension for 90 s at 72°C, and then a 10 min extension at 72°C [28] . For C. cayetanensis, amplification was performed using two sets of primers; F: 5′-GCAGTCACAGGAGGCATATATCC-3′ and R: 5′-ATGAGAGACCTCACAGCCAAAC-3′. Thermocycling conditions consisted of denaturation at 95°C for 2 min, followed by 40 cycles of denaturing at 95°C for 30 s, annealing at 59°C for 30 s, and extension at 72°C for 30 s, and a final extension for 5 min at 72°C [27] . PCR products were electrophoresed in 2.5% (w/v) agarose gel. The gels were stained by immersion in 1 μg/ml ethidium bromide for 15 min and photographed under 300 nm ultraviolet light [27],[28] .

Statistical analysis

Descriptive measures included count, percentage, arithmetic mean, and SD. Statistical tests included the Mann - Whitney Z and Friedman tests. They were used for discrete variables [32] . The level of significance was at P value equal to or less than 0.05.

Ethical consideration

Stool samples were collected after obtaining the patients' informed consents. Both human sampling and mice handling got the approval of the Ethics Committee of Alexandria University.


Examination of fish and human samples

Of the 35 fishes, nine were infected with Cryptosporidium spp. (25.7%) [Figure 1] and four were infected with Cyclospora spp. (11.4%) [Figure 2]. Microsporidium spores were also detected in 13 samples stained by modified trichrome stain (37.1%). Of the 50 human samples, 14 were positive for intestinal parasites. Cryptosporidium spp. was detected in four samples (8%) and Cyclospora spp. in two samples (4%). The size and shape of both organisms detected in human stool samples were similar to those detected in fish intestinal contents. Other intestinal parasites diagnosed were Giardia lamblia (n = 3), G. lamblia and Blastocystis hominis (n = 1), and Microsporidium spp. (n = 4).{Figure 1}{Figure 2}

Mice infection

Mice stools revealed progressive increase in the number of shed oocysts from day 7 to 14, with statistical significance using the Mann - Whitney Z-test and the Friedman test [Table 1], except in subgroup IId infected by Cyclospora oocysts from humans. No oocysts were detected in the stool of the control group.{Table 1}

Histopathological findings

LM examination of hematoxylin and eosin-stained intestinal sections of mice infected by oocysts isolated from fish and humans demonstrated similar Cryptosporidium oocysts on the brush border of columnar epithelial cells [Figure 3]. Similar Cyclospora oocysts were attached to the brush borders and were intracellularly located in a supranuclear location. Multiple immature oocysts were also observed within the gut lumen [Figure 4]. The infected intestinal sections showed altered mucosal architecture, with shortening and widening of the intestinal villi [Figure 5].{Figure 3}{Figure 4}{Figure 5}

Transmission electron microscopic findings

Cryptosporidium spp. organisms were able to undergo a complete reproductive cycle, forming both asexual and sexual stages, located within parasitophorous sacs at the intracellular, extracytoplasmic part of the intestinal epithelium. A typical dense band was present within the affected host cells. Most of the stages exhibited typical apicomplexan ultrastructural characteristics. In some mice infected by Cryptosporidium spp. isolated from fish, trophozoite stages were detected at the microvillus surface of the epithelium. They were rounded, enveloped by a parasitophorous sac, and each contained a nucleus with a prominent nucleolus. Multiple spherical ribosomes surrounded the nucleus. Great distortion of the microvilli was observed [Figure 6]. In other intestinal sections of the same subgroup (IIa), macrogamont stages located within the parasitophorous sacs possessed a compact nucleus, vacuolated cytoplasm, several dense bodies, and amylopectin granules [Figure 7]. In mice infected also by Cryptosporidium spp. isolated from humans, trophozoite and macrogamont stages were detected. Their ultrastructural characteristics were identical to the corresponding stages observed in subgroup IIa [Figure 8]. Mature Cryptosporidium oocysts with a large residual body, containing sectioned sporozoites were detected in the intestinal lumen of subgroup IIa. The nearby intestinal microvilli showed distortion [Figure 9]. A single rhoptry in the apical region was detected in few sporozoites. A free Cryptosporidium sporozoite observed in the intestinal lumen of subgroup IIb was morphologically similar to the sporozoite detected in the sections of subgroup IIa. It was oval in shape with an anteriorly located vacuole, micronemes, and dense granules and a posteriorly located nucleus. Major alteration was noticed in the intestinal microvilli [Figure 10].{Figure 6}{Figure 7}{Figure 8}{Figure 9}{Figure 10}

The extracellular immature Cyclospora oocysts detected in the intestinal sections of mice infected by both fish and human species were seen either in the gut lumen or attached to the enterocyte microvilli [Figure 11]. The immature oocysts of subgroups IIc and IId were typical. They appeared as roughly spherical organisms, 8-10 μm in diameter, comprised of an outer fibrillar coat, a thinner cell wall, and a cell membrane enclosing the cytoplasm. The internal cytoplasmic mass contained light and dark granules [Figure 12]a and b.{Figure 11}{Figure 12}

Therefore, by TEM, identification of the coccidian parasites to the genus level was achieved as genus Cryptosporidium and genus Cyclospora. However, differentiation between species isolated from fish and humans was not achieved.

Molecular findings

DNA was extracted successfully from the four positive samples [Figure 13]. The specific primers used in this study were found to amplify C. parvum and C. cayetanensis from human stool specimens at 452-bp for the first and 116-bp for the second [Figure 14] and [Figure 15]. The specific primers used failed to amplify the Cryptosporidium and Cyclospora spp. isolates from fish intestinal contents. The application of the agarose-embedded DNA preparation method canceled the possibility that the absence of amplification in the fish samples was related to the PCR inhibitors.{Figure 13}{Figure 14}{Figure 15}


Various community outbreaks due to contamination of water or food with Cryptosporidium and Cyclospora spp. have further highlighted their importance in public health [1,8,33]. This study was undertaken to compare Cryptosporidium and Cyclospora spp. detected in fish with the corresponding species isolated from infected humans, morphologically by LM and TEM and genetically by DNA extraction and amplification using specific primers for C. parvum and C. cayetanensis.

Morphological identification was supported by the study of Jirku et al. [26] , who claimed that species identification of apicomplexan parasites is traditionally based on host specificity, morphology of life-cycle stages, and pathology associated with an individual host. Our LM findings revealed that Cryptosporidium and Cyclospora oocysts of both fish and humans were identical. Moreover, similar intestinal pathological changes were demonstrated for both parasites. Recorded morphology for Cryptosporidium spp. was as reported by other studies [34],[35] stating that oocysts in fish were ˜4.6×4.4 μm, located along the epithelial lining of some areas of the fish stomach but seldom found in the intestine.

The documented histological damage was also in accordance with the study by Jirku et al. [26] , who claimed that infection in amphibian hosts (toads) by Cryptosporidium spp. produced pathological changes in the mucosa and within the muscular tunic and serosa. Furthermore, infectivity to experimental mice by Cryptosporidium and Cyclospora oocysts from T. zilli was confirmed by Diab et al. [36] . Cyclospora oocysts were not commonly encountered in our study. This may be attributed to their immature developmental condition when excreted in the environment or because of their low prevalence in the aquatic environment [14,15,36]. In contrast, the high detection rate of Microsporidium spp. encountered in fish may be related to the stability of their spores in the environment and their capability of remaining infective for weeks outside their hosts [37] .

TEM findings for mice intestinal sections showed morphological features consistent with previously reported studies [34],[38] . These findings could be explained by understanding the invasion strategy of Cryptosporidium spp. [38],[39] . It was shown that, when sporozoites attach to the host cell membrane, their rhoptries extend to the attachment sites, whereas both micronemes and dense granules are recruited to the apical complex regions of the attached parasites. During internalization, numerous vacuoles covered by the parasites' plasma membranes are clustered together to establish a preparasitophorous vacuole. This vacuole comes in contact with the host cell membrane to form a host cell-parasite membrane interface, beneath which an electron-dense band begins to appear within the host cell cytoplasm. Simultaneously, host cells display membrane protrusions, resulting in the formation of mature parasitophorous vacuoles covering the parasite [38],[39] . By TEM, we were able to demonstrate this electron-dense band within the Cryptosporidium spp.-infected host cells. An anteriorly located vacuole noticed in some sporozoites was also reported by Huang et al. [38] , who supposed that during internalization of host epithelial cells by C. parvum, vacuole-like structures appear in the apical complex region of the sporozoites. Besides, O'Hara et al. [40] stated that through the stages of internalization of C. parvum upon the host cell, the apically segregated micronemes were opposed to a central microtubule-like filamentous structure, and the more distal micronemes formed the anterior vacuole. TEM findings of mice intestinal sections of subgroups IIc and IId showed immature spherical Cyclospora oocysts. Two other studies [41],[42] reported similar structural details as those noticed in the present study.

Our study revealed that the molecular methods were apparently the only techniques capable of achieving successful differentiation between human and fish species of Cryptosporidium and Cyclospora. On the basis of failure of PCR amplification of oocysts from infected fish samples, it was revealed that fish species were not identical to human species. This was supported by other studies [43],[44] , which recorded that PCR offers alternatives to the traditional diagnosis for both clinical and environmental samples. It was formerly implied that application of PCR to stool specimens is limited because of the presence of substances inhibiting the reaction [31] , even when present at low concentrations. These occur in stools, tissue extracts, and body fluids, and include bilirubin, bile salts, and complex polysaccharides [45],[46] . In our study, by following the application of agarose-embedded DNA preparation technique, we avoided the possibility of failure of amplification related to the presence of PCR inhibitors. This was in accordance with the study by Moreira [30] , who proposed that agarose-embedded DNA is a simple, inexpensive, and useful method for the removal of soluble PCR inhibitors present as natural contaminants of DNA samples. By applying this method, Monteiro et al. [29] also obtained a positive PCR proving that inhibitors present in the original DNA samples were completely removed.

The amplification of the 452-bp band specific for C. parvum performed in our study was in accordance with other studies [28,47,48], and it was successfully applied for infected specimens from humans, calves, and goats [28] . Amplification of 116-bp specific for C. cayetanensis coincided with the study by Lalonde and Gajadhar [27] , who used the same specific primers for the detection of C. cayetanensis in fecal human samples, and concluded that these primers provided highly sensitive and reliable results. As Cyclospora spp. is an emerging parasite that cannot be cultured in vitro, there is a very limited amount of specific species sequence information available in GenBank. To our knowledge, very few studies have reported validation of a C. cayetanensis DNA extraction and the PCR detection technique for use with fecal samples [27],[49] .


Therefore, we concluded that, by comparing the three methods applied in the present study for differentiation between Cryptosporidium and Cyclospora spp. of fish and humans, the molecular methods proved to be the best. They indicated that species detected in human and fish were not similar. Thus, from our epidemiological point of view, fish are not suitable hosts for these water-transmitted human pathogens. It is recommended that further genetic studies and molecular specificity testing should be carried out to obtain acceptable sequence information of fish Cryptosporidium and Cyclospora spp.

 Author Contribution

M.M. El Temsahy and E.D. El-Kerdany proposed the research idea and shared in the assessment of the light and electron microscope and histopathology findings. M.R. Gaafar and R.G. Diab shared in the study design, performed the experiments, and shared in the assessment of the light and electron microscope, histopathology, and molecular findings. I.H. Diab performed the molecular studies. All authors contributed in the writing of the manuscript.



1Ribes JA, Seabolt JP, Overman SB. Point prevalence of Cryptosporidium, Cyclospora, and Isospora infections in patients being evaluated for diarrhea. Am J Clin Pathol 2004; 122:28-32.
2 Chalmers RM, Davies AP. Minireview: clinical cryptosporidiosis. Exp Parasitol 2010; 124:138-146.
3 Fayer R. Taxonomy and species delimitation in Cryptosporidium. Exp Parasitol 2010; 124:90-97.
4 Šlapeta J. Centenary of the genus Cryptosporidium: from morphological to molecular species identification. In: Ortega-Pierres MG, et al., editors. Giardia and Cryptosporidium: from molecules to disease. Wallingford, UK: CAB International; 2009, 31-50.
5 Arcay L, Baez De Bordes E, Bruzal E. Cryptosporidiosis experimental en la escala de vertebrados. I. Infections experimentales II. Estudio histopathologico. Parasitol Al Dia 1995; 19:20-29.
6 Ryan U. Cryptosporidium in birds, fish and amphibians. Exp Parasitol 2010; 124:113-120.
7 Zanguee N, Lymbery A, Lau J, et al. Identification of novel Cryptosporidium species in aquarium fish. Vet Parasitol 2010; 174:43-48.
8 Mansfield LS, Gajadhar AA. Cyclospora cayetanensis, a food- and waterborne coccidian parasite. Vet Parasitol 2004; 126:73-90.
9 Blans MCA, Ridwan BU, Verweij JJ, Rozenberg-Arska M, Verhoef J. Cyclosporiasis outbreak, Indonesia. Emerg Infect Dis 2005; 11:1453-1455.
10Eberhard ML, DaSilva AJ, Lilley BG, Pieniazek NJ. Morphologic and molecular characterization of new Cyclospora species from Ethiopian monkeys: C. ceropitheci sp. n., C. colobi sp. n., and C. papionic sp. n. Emerg Infect Dis 1999; 5:651-658.
11Ford PL, Duszynski DW, McAllister CT. Coccidia (Apicomplexa) from heteromyid rodents in the Southwestern United States, Baja California, and Northern Mexico, with three new species from Chaetodipus hispidus. J Parasitol 1990; 76:325-331.
12García López HL, Rodríguez Tovar LE, Garza CEM. Identification of Cyclospora in poultry. Emerg Inf Dis 1996; 2:356-357.
13Lainson R. The genus Cyclospora (Apicomplexa: Eimeriidae), with a description of Cyclospora schneideri n. sp. in the snake Anilius scytale scytale (Aniliidae) from Amazonian Brazil: a review. Mem Inst Oswaldo Cruz 2005; 100:103-110.
14Graczyk TK, Ortega YR, Conn DB. Recovery of water-borne oocysts of Cyclospora cayetanensis by Asian freshwater clams (Corbicula fluminea). Am J Trop Med Hyg 1998; 56:928-932.
15Negm AY. Human pathogenic protozoa in bivalves collected from local markets in Alexandria. J Egypt Soc Parasitol 2003; 33:991-998.
16Marin-Leonett M, Figuera L, Nessi A, et al. Diarrhea due to Cyclospora-like organism in an immunocompetent patient. J Infect Dev Ctries 2007; 1:345-347.
17El Azzouni MZ. Heterologous immunity in schistosomiasis using heterophyid antigens [MD thesis]. Alexandria: Faculty of Medicine, University of Alexandria; 1988.
18Giarcia LS, Bruckner DA. Macroscopic and microscopic examination of fecal specimens. In: Giarcia LS, Bruckner DA, editor. Diagnostic medical parasitology. 3rd ed. Washington, DC: ASM Press; 1997. 608-649.
19Gaafar MR. Effect of solar disinfection on the viability of intestinal protozoa in drinking water. J Egypt Soc Parasitol 2007; 37:65-86.
20Carnevale S, Velasquez JN, Labbe JH, Cabrera MG, Rodrigez MI. Diagnosis of Enterocytozoon bieneusi by PCR in stool samples eluted from filter paper disks. Clin Diagn Lab Immunol 2000; 7:504-506.
21Lumb R, Swift J, James C, Papanaoum K, Mukh-erjee T. Identification of the microsporidian parasite, Enterocytozoon bieneusi in fecal samples and intestinal biopsies from an AIDS patient. Int J Parasitol 1993; 23:793-801.
22Sadaka HA, Zoheir MA. Experimental studies on cyclosporiasis. J Egypt Soc Parasitol 2001; 31:65-77.
23El-Fakhry Y, Achbarou A, Desportes I, Mazier D. Encephalitozoon intestinalis: humoral responses in interferon-y receptor knockout mice infected with a microsporidium pathogenic in AIDS patients. Exp Parasitol 1998; 89:113-121.
24Allam SR, Sadaka HA, Eissa MM, Baddour NM. A novel macrolide in mixed protozoal infection in immunosuppressed mice. J Med Res Inst 1999; 20:149-159.
25Drury RAB, Wallington EA. Carleton's histological technique. 5th ed. Oxford, New York, Toronto: Oxford University press; 1980.
26Jirku M, Valigurova A, Koudela B, Køížek J , Modrý D, Slapeta J. New species of Cryptosporidium Tyzzer, 1907 (Apicomplexa) from amphibian host: morphology, biology and phylogeny. Folia Parasitol 2008; 55:81-94.
27Lalonde LF, Gajadhar AA. Highly sensitive and specific PCR assay for reliable detection of Cyclospora cayetanensis oocysts. Appl Environ Microbiol 2008; 74:4354-4358.
28Gobet P, Buisson JC, Vagner O, et al. Detection of Cryptosporidium parvum DNA in formed human feces by a sensitive PCR-based assay including uracil-N-glycosylase inactivation. J Clin Microbiol 1997; 3:254-256.
29Monteiro L, Gras N, Vidal R, Cabrita J, Mégraud F. Detection of Helicobacter pylori DNA in human feces by PCR: DNA stability and removal of inhibitors. J Microbiol Methods 2001; 45:87-94.
30Moreira D. Efficient removal of PCR inhibitors using agarose-embedded DNA preparations. Nucleic Acids Res 1998; 26:3309-3310.
31Monteiro L, Gras N, Megraud F. Magnetic immuno-PCR assay with inhibitor removal for direct detection of Helicobacter pylori in human feces. J Clin Microbiol 2001; 39:3778-3780.
32Knapp RG, Miller MC. Clinical epidemiology and biostatistics. Baltimore: Williams and Wilkins; 1992. 43-45.
33Thompson RCA, Olson ME, Zhu G, Enomoto S, Abra-Hamsen MS, Hijjawi NS. Cryptosporidium and cryptosporidiosis. Adv Parasitol 2005; 59:77-158.
34Alvarez-Pellitero P, Sitjà-Bobadilla A. Cryptosporidium molnari n. spp. (Apicomplexa: Cryptosporidiidae) infecting two marine fish species, Sparus aurata L. and Dicentrarchus labrax L. Int J Parasitol 2002; 32:1007-1021.
35Ryan U, O'Hara A, Xiao L. Molecular and biological characterization of a Cryptosporidium molnari-like isolate from a guppy (Poecilia reticulata). Appl Environ Microbiol 2004; 70:3761-3765.
36Diab RMG, El Temsahy MM, El Kerdany ED, Gaafar MR, Nasr MA. A study of medically important fish-transmitted parasites in Alexandria. Int J Inf Dis 2010; 14:e290.
37Slifko TR, Smith HV, Rose JB. Emerging parasite zoonosis associated with water and food. Int J Parasitol 2000; 30:1379-1393.
38Huang BQ, Chen X, LaRusso NF. Cryptosporidium parvum attachment to and internalization by human biliary epithelia in vitro: a morphological study. J Parasitol 2004; 90:212-221.
39Umemiya R, Fukuda M, Fujisaki K, Matsui T. Electron microscopic observation of the invasion process of Cryptosporidium parvum in severe combined immunodeficiency mice. J Parasitol 2005; 91:1034-1039.
40O'Hara SP, Huang BQ, Chen X, Nelson J, LaRusso NF. Distribution of Cryptosporidium parvum sporozoite apical organelles during attachment to and internalization by cultured biliary epithelial cells. J Parasitol 2005; 91:995-999.
41Ortega YR, Sterling CR, Gilman RH, Cama VA, Diaz F. Cyclospora species: a new protozoan pathogen of humans. N Eng J Med 1993; 6:1308-1312.
42Satheeshkumar S, Ananthan S. Electron microscopy identification of microsporidia (Enterocytozoon bieneusi) and Cyclospora cayetanensis from stool samples of HIV infected patients. Ind J Med Microbiol 2004; 22:119-122.
43Morgan UM, Thompson RC. Molecular detection of parasitic protozoa. Parasitology 1998; 117:73-85.
44Fanzen C, Muller A. Molecular techniques for detection, species differentiation, and phylogenetic analysis of microsporidia. Clin Microbiol Rev 1999; 12:243-285.
45Morgan UM, Pallant L, Dwyer BW, Forbes DA, Rich G, Thompson RC. Comparison of PCR and microscopy for detection of Cryptosporidium parvum in human fecal specimens: clinical trial. J Clin Microbiol 1998; 36:995-998.
46Widjoatmodjo MN, Fluit AC, Torensma R, Verdonk GP, Verhoef J. The magnetic immunopolymerase chain reaction assay for direct detection of Salmonella in fecal samples. J Clin Microbiol 1992; 30:3195-3199.
47Laxer MA, D'Nicuola ME, Patel RJ. Detection of Cryptosporidium parvum DNA in fixed paraffin-embedded tissue by the polymerase chain reaction. Am J Trop Med Hyg 1992; 47:450-455.
48Laxer MA, Timblin BK, Patel RJ. DNA sequences for the specific detection of Cryptosporidium parvum by the polymerase chain reaction. Am J Trop Med Hyg 1991; 45:688-694.
49Relman DA, Schmidt TM, Gajadhar AA, et al. Molecular phylogenetic analysis of Cyclospora, the human intestinal pathogen, suggests that it is closely related to Eimeria species. J Infect Dis 1996; 173:440-445.