Genetic Diversity of Cryptosporidium spp. in Captive Reptiles (2024)

Samples.

A total of 123 diagnostic samples obtained from captive snakes, lizards, and tortoises from the United States, Switzerland, the Czech Republic, Ghana, and Australia were used in this study (Tables ​(Tables11 through ​through3).3). These included 88 samples from snakes, 26 samples from lizards, and 9 samples from tortoises. Another 19 samples from cross-transmission studies (11 snake samples and 8 lizard samples) were also studied. With the exception of samples from the Saint Louis Zoo (78 snake samples, 7 lizard samples, and 3 tortoise samples), Louisville Zoo (6 tortoise samples), and transmission studies, all samples were previously diagnosed as Cryptosporidium positive by microscopy, and purified oocysts were used in molecular studies. Samples from the Saint Louis Zoo, Louisville Zoo, and National Zoological Park were mostly feces, with the exception of gastric washings from three snakes in the Saint Louis Zoo.

After initial diagnosis of cryptosporidiosis in snakes in the Saint Louis Zoo at the end of 1998, a cryptosporidiosis control program was initiated, which involved the diagnosis and differentiation of Cryptosporidium infection by PCR and euthanasia of C. serpentis- or C. saurophilum-infected snakes. To monitor the effectiveness of this cryptosporidiosis control measure, 8 to 17 snakes at the zoo were examined for Cryptosporidium infection periodically for 1 year from May 1999 to April 2000, with continuous euthanasia of infected animals (Table ​(Table4).4). To identify the source of isolates belonging to the C. parvum mouse genotype and C. muris in snakes, fecal samples were also collected from 11 of the feeder mice used in the Saint Louis Zoo and examined for Cryptosporidium species and genotypes.

Cross-transmission studies.

To evaluate the infectivity of C. saurophilum to snakes and lizards, two corn snakes and leopard geckoes were inoculated with 10,000 oocysts originating from a bull snake (sample 815). Forty-five days after the experimental infection, animals were euthanized, and the stomach and intestine and their contents were examined for Cryptosporidium oocysts by PCR-RFLP and DNA sequence analysis of the SSU rRNA gene. The infectivity for snakes of C. serpentis isolated from lizards was assessed by experimental infection of a captive-born Burmese python (Python mollurus) with oocysts isolated from a wild imported juvenile Nile monitor (Varanus niloticus) from Togo. Cryptosporidium oocysts in the python's feces were genotyped by DNA sequencing of the SSU rRNA gene 88 days after the inoculation. The cross-transmission of Cryptosporidium spp. between snakes and lizards was further evaluated by the differentiation of Cryptosporidium spp. and genotypes in a group of six snakes and four lizards that were housed in the same room by the SSU rRNA PCR-RFLP analysis (Table ​(Table33).

Morphometric measurements.

Oocysts of C. serpentis from a desert monitor (sample 806) and of C. saurophilum from a bull snake (sample 815) were measured under a differential interference contrast microscope at a magnification of ×1,000. Twenty oocysts were measured for C. saurophilum organisms, and 37 oocysts were measured for C. serpentis organisms. Mean length and width and the shape index were calculated along with the 95% confidence limits (CL) for each species.

DNA extraction.

Purified oocysts or fecal samples containing oocysts were used in DNA extraction. DNA was extracted from stool samples by alkaline digestion and phenol-chloroform extraction, followed by DNA purification with a commercial kit. Briefly, 33.3 μl of 1 M KOH and 9.3 μl of 1 M dithiothreitol were added to a 1.5-ml microcentrifuge tube containing 100 μl of stool or oocyst suspension. After incubation at 65°C for 15 min, the solution was neutralized with 4.3 μl of 25% hydrochloric acid and buffered with 80 μl of 2 M Tris-HCl (pH 8.3). The DNA was extracted with 250 μl of phenol-chloroform-isoamyl alcohol (Invitrogen, Carlsbad, Calif.) after thorough mixing and centrifugation in an Eppendorf (Hamburg, Germany) microcentrifuge at 5,000 × g for 5 min. The supernatant was transferred to a 2.0-ml Eppendorf tube containing 1.0 ml of ASL buffer from the QIAamp DNA Stool Mini Kit (QIAGEN, Valencia, Calif.). The DNA was further purified following the manufacturer-suggested procedures. DNA was stored at −70°C before it was used in molecular analysis.

Species differentiation and genotyping.

Cryptosporidium spp. and C. parvum genotypes present were diagnosed by a PCR-RFLP technique (13, 16, 17). In this method, a segment (∼833 bp) of the Cryptosporidium SSU rRNA gene was amplified by nested PCR. Species and genotype diagnosis was made by restriction digestion of the secondary PCR product with SspI (New England BioLabs, Beverly, Mass.) and VspI (Promega, Madison, Wis.). Each sample was examined at least twice by independent PCR-RFLP analyses. To confirm the diagnosis of new Cryptosporidium spp. and to identify genetic heterogeneity within C. serpentis and C. saurophilum, secondary PCR products were sequenced in both directions on an ABI Prism 3100 analyzer (Applied Biosystems, Foster City, Calif.) by using forward and reverse primers, after PCR products had been purified with the Wizard PCR Prep Kit (Promega). Nucleotide sequences obtained from this study were aligned against each other by using the ClustalX (11) program and manual adjustment. A neighbor-joining tree was constructed from the aligned sequences as previously described by using the Treecon program, and genetic distances were calculated with the Kimura 2-parameter model (17).

Because of the presence of mixed Cryptosporidium species in samples from snakes and lizards that were housed together in cross-transmission studies, PCR products from one of the snakes (sample 938) and one of the lizards (sample 944) were cloned into a pGEM-T vector (Promega). Eight (for sample 944) or 15 (for sample 938) clones were sequenced for each PCR product to confirm the diagnosis.

Nucleotide sequence accession numbers.

The nucleotide sequences of the partial SSU rRNA gene have been deposited in the GenBank database under accession numbers AF093499, AF093501, AF112573, AY120913 through AY120915, and AY268581 through AY268584.

Cryptosporidium spp. in reptiles.

SSU rRNA PCR confirmed that all snake and lizard samples from Maryland (three snake samples), Washington, D.C. (one snake sample and four lizard samples), Kansas (one snake sample), Ghana (one lizard sample), the Czech Republic (five lizard samples), Switzerland (one snake sample and nine lizard samples), and Australia (one snake sample and one lizard sample) were positive for Cryptosporidium. The samples were previously diagnosed as Cryptosporidium positive by microscopy. In contrast, 3 of the 6 tortoise samples from the Louisville Zoo, 36 of 81 snake samples, 4 of 7 lizard samples, and 0 of 3 tortoise samples from the Saint Louis Zoo were positive for Cryptosporidium by PCR. These animals had not been previously screened for Cryptosporidium oocysts by microscopy.

PCR-RFLP analysis of SSU rRNA PCR revealed banding patterns distinctive for C. serpentis and the Cryptosporidium desert monitor genotype (Tables ​(Tables11 and ​and2).2). Cryptosporidium serpentis was found in 28 of 48 positive snake samples and 11 of 25 positive lizard samples, and the Cryptosporidium desert monitor genotype was identified in 3 of 48 positive snake samples and 9 of 24 positive lizard samples. Sequence analyses confirmed the results of RFLP analyses but also revealed genetic diversities within C. serpentis and the Cryptosporidium desert monitor genotype. Most desert monitor genotype isolates had identical SSU rRNA sequences, but isolates 1343 and 1786 had one single nucleotide polymorphism (SNP). Two genotypes (A and B) were seen in C. serpentis isolates, which differed from each other by one SNP (Fig. ​(Fig.1).1). One isolate from a lizard (sample 1665) had the C. serpentis RFLP banding pattern, was related to the two C. serpentis genotypes, but had significant differences in nucleotide sequence (Fig. ​(Fig.11).

Several other Cryptosporidium spp. were also identified in reptiles. RFLP analysis showed banding patterns of the C. parvum mouse genotype in 12 snakes and 1 lizard, of the C. parvum bovine genotype in 6 lizards, and of C. muris in 3 snakes. DNA sequence analysis confirmed the identifications, as the sequences obtained were identical to previously reported sequences (16, 17). Two other new Cryptosporidium spp. were isolated in reptiles, as shown by both RFLP and sequence analyses. In RFLP analyses, one isolate from a snake (sample 2162) had an SspI band of over 800 bp but had a VspI band similar to the size of the band of the desert monitor genotype (between 600 to 700 bp). In contrast, three tortoise isolates had a slightly smaller SspI band size (near 800 bp) but had a VspI band similar to the band of C. serpentis (over 700 bp). DNA sequencing revealed that the sequences represented two new Cryptosporidium spp. A neighbor-joining analysis indicated that the tortoise genotype was related to C. serpentis and C. muris, whereas the new snake genotype was related to C. parvum (Fig. ​(Fig.22).

Because C. muris and the C. parvum mouse genotype were seen in high frequency in snakes and lizards from the Saint Louis Zoo, whose diet contained mice, fecal samples were taken from 11 feeder mice and analyzed for Cryptosporidium. Three such samples were positive for Cryptosporidium by PCR analysis of the SSU rRNA gene. RFLP and sequence analyses showed the presence of the C. parvum mouse genotype in two mice and a mixed infection of C. muris and the C. parvum mouse genotype in one mouse.

The identification of the desert monitor genotype as C. saurophilum.

Previous characterization of the Cryptosporidium desert monitor genotype showed that the parasite is closely related to intestinal Cryptosporidium spp. Because the only known intestinal Cryptosporidium parasite in reptiles is C. saurophilum, morphometric measurements were done on the desert monitor genotype, and the data obtained were compared with those from C. serpentis and those previously reported for C. saurophilum. Oocysts of the desert monitor genotype were visibly smaller than those of C. serpentis (Fig. ​(Fig.3),3), with a mean length of 4.94 μm, a mean width of 4.49 μm, and a shape index (the length/width ratio) of 1.14. In comparison, oocysts of C. serpentis were 5.94 μm in length and 5.11 μm in width and had a shape index of 1.17 (Table ​(Table5).5). Thus, the morphometric measurements of the desert monitor genotype were similar to those previously reported for C. saurophilum (mean, 5.0 × 4.7 μm; range, 4.4 to 5.6 × 4.2 to 5.2 μm) (6).

Cross-transmission of Cryptosporidium between snakes and lizards.

To assess the ability for cross-transmission of Cryptosporidium spp. between snakes and lizards, an isolate of the Cryptosporidium desert monitor genotype originating from a snake was used to inoculate two corn snakes and two leopard geckoes that were free of Cryptosporidium infection by microscopy. Both geckoes started to shed oocysts 21 days after inoculation, but both snakes remained negative through the observation period. All four inoculated animals were euthanized 45 days after the inoculation, and tissue sections were taken from the stomach and intestine for histology and PCR-RFLP analysis. Cryptosporidium was not found in any of the tissue sections from the snakes. In contrast, Cryptosporidium in developmental stages was seen in hematoxylin and eosin-stained gastric tissues. PCR analysis of DNA from gastric and intestinal fragments confirmed that both snakes were negative for Cryptosporidium. Positive PCR amplifications, however, were obtained with DNA from the intestine of one gecko and from both the stomach and intestine of the other gecko. RFLP analysis of the PCR products with SspI and VspI indicated that the parasites present belonged to the Cryptosporidium desert monitor genotype (Table ​(Table3).3). The python inoculated with C. serpentis oocysts from a Nile monitor started to shed Cryptosporidium oocysts 88 days after inoculation. PCR-RFLP analysis of oocysts isolated from the snake confirmed that it belonged to C. serpentis (Table ​(Table3).3). Sequence analysis produced an SSU rRNA sequence identical to that of the isolate from the monitor, C. serpentis type B (Fig. ​(Fig.22).

The cross-transmission of Cryptosporidium spp. between snakes and lizards was further assessed by the differentiation of Cryptosporidium spp. in a group of six snakes and four lizards that were housed in the same room. All snakes and lizards were positive for Cryptosporidium. PCR-RFLP analysis indicated that all animals were infected with multiple Cryptosporidium spp.; the six snakes were all infected with C. serpentis, a new Cryptosporidium genotype, and low levels of C. saurophilum, whereas the four lizards were all infected with C. serpentis and C. saurophilum (Fig. ​(Fig.44 and Table ​Table3).3). Analysis of the cloned PCR products confirmed the diagnosis of C. serpentis and the new Cryptosporidium genotype in snakes and C. serpentis and C. saurophilum in lizards. Among the 15 clones of a PCR product from the snake isolate 938, 4 clones were identified as C. serpentis, and 11 clones were identified as the new Cryptosporidium genotype. Likewise, four of the eight clones of a PCR product from lizard isolate 944 belonged to C. saurophilum, and the remaining four clones belonged to C. serpentis. The new Cryptosporidium genotype was genetically related to the intestinal Cryptosporidium spp. and had a large SspI band (about 800 bp), similar to the tortoise genotype (Fig. ​(Fig.4,4, filled arrow), but had a VspI upper band (just over 600 bp) similar to that of C. saurophilum (Fig. ​(Fig.4,4, open arrow).

Effectiveness of the diagnosis-euthanasia control strategy.

At the Saint Louis Zoo, a diagnosis-euthanasia program was initiated in March 1999 after the identification between December 1998 and January 1999 of chronic cryptosporidiosis in snakes. To monitor the effectiveness of the control measures, samples were periodically taken from snakes for 1 year. Right after the initiation of the control measure, 5 of 10 and 8 of 17 snakes sampled were positive for C. serpentis or C. saurophilum in May and June 1999, respectively. Afterwards, only 1 of 45 snake samples taken at five different time periods was positive for C. serpentis (Table ​(Table44).

We thank Roy Burns of the Louisville Zoo and Richard J. Montali of the National Zoological Park for providing samples and Michael Cranfield of the Baltimore Zoo for facilitating the cross-transmission experiment.

This study was supported in part by grant no. 524/00/P015 of the Grant Agency of the Czech Republic

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