Practical tips for reptile fecal exams

by samantha_ashenhurst | September 28, 2022 12:01 pm

[1]

While a growing number of reptiles are being kept as pets,¹ there is still much to be learned about the parasitic infections these creatures may harbour. Parasites are organisms that have adapted to live on or within a host organism, often causing some amount of detriment. Indeed, these pathogens may affect hosts in numerous ways, whether through impairment to locomotion, thermoregulation, reproduction, or survival. Further, infections may also impact population dynamics.² Recent reviews have been published on various taxonomic groups of parasites which utilize reptiles as definitive hosts; however, these checklists are often narrowly focused on specific geographic regions.^1,3,4

[2]

In contrast to domestic companion animals, parasites tend to be relatively common in pet reptiles, and some of these species are also known to be zoonotic.^1,5 More than 20 different species of nematodes have been reported to occur in reptiles,³ and infections with acanthocephalans,⁴ coccidia,⁶ other protozoans,⁵ numerous species of trematodes,⁷ and pentastomids have also been described.

Similar to what is done in small animal medicine, many internal parasites can be detected in reptiles through examination of the host’s feces. While sedimentation is commonly employed for the detection of relatively heavier eggs (e.g. trematodes), and the Baermann technique is used for the detection of larvae, fecal flotation remains a highly effective technique for the detection of common eggs, cysts, and oocysts, especially when centrifugation is incorporated.

Sample collection procedures

Diagnostic testing on fecal samples can either be performed in-house or through submission to an outside laboratory. Samples should be processed as soon as possible, as impaired recovery of parasite stages cannot be ruled out after a prolonged period of time.⁵

It is important that fecal samples are fresh and stored properly in a screw-top cup or in a bag devoid of air while they are being shipped or prior to processing. If these samples are not fresh, there is an increased chance eggs will develop and hatch and desiccate; or, alternatively, non-parasitic, free-living nematodes and fly larvae will contaminate the sample. To ensure the sample is fresh, it should optimally be collected within one to two hours of defecation, and, if shipping is required, it should be sent on ice, but not frozen. Once the sample has been received, it is best to keep it refrigerated (at 4 C) until evaluation.

[3]

MDCF overview

[4]

There are multiple coprological examination techniques used to detect internal parasites in reptiles. While wet mounts or direct preparations have their utility in certain situations, these methods have some inherent drawbacks. Particularly, the technique is limited by the fact very little fecal material is used, and no concentration step is performed. Additionally, as movement is one of the principal characteristics allowing recognition of trophozoites in this procedure, if the fecal layer is too thick, it is challenging to appreciate small, colourless protozoa moving in the field.⁵

 

In the U.S., one technique employed at the Colorado State University Veterinary Diagnostic Laboratories (CSU-VDL) is a modified double centrifugal technique (MDCF), which uses the principles of centrifugation and specific gravity (SpG) to detect parasitic eggs, cysts, and oocysts in a sample. In the MDCF procedure, the reptile fecal sample is first centrifuged in water. During this process, parasitic elements and debris will form a pellet at the bottom of a conical vial. This is followed by a second centrifugal spin in a sugar or salt solution, which allows for any parasitic elements present to float to the top of the vial and adhere to a seated coverslip. This coverslip is examined under a compound microscope on 100x magnification (Table 1).

[5]

In CSU-VDL’s Parasitology section, Sheather’s sugar (SpG 1.27), a commercially available media, serves as the fecal flotation media of choice and is used for all host groups, including reptiles. (There are, however, several types of fecal flotation media which can be made in-house, detailed in Table 2.) After preparing any type of media, the correct SpG may be verified using a heavy fluid hydrometer (Figure 2). After placing the media in a graduated cylinder, this instrument is used by placing it in the solution and recording the value at which the media crosses the graduated stem. Most commercial and homemade solutions should have a SpG of 1.18 to 1.30; however, a SpG of greater than 1.25 is able to reliably detect most eggs, cysts, and oocysts of veterinary importance across multiple host groups.

With regards to centrifugation, it is appropriate to use either a swinging bucket or fixed-bucket centrifuge (Figure 3). The advantage of a swinging-head centrifuge that has a slow start and slow stop is that vials can often be spun with coverslips seated in place. If only the fixed-bucket style is available, it is advisable the following steps be carried out:

1. Add additional media to the conical vial (following the second spin in a sugar or salt solution) to bring the volume flush with the top of the tube.
2. Place a coverslip on top of the tube.
3. Wait an additional 10 minutes after centrifugation has ended to allow parasitic elements to float and adhere to the coverslip prior to evaluating the sample under the microscope (Table 1).

[6]

Calibration and measuring

[7]

As highlighted in the Veterinary Parasitology Reference Manual, proper calibration is an important step in accurately measuring parasites under the microscope.⁸ Before viewing fecal slides on a microscope, it is important to ensure the microscope has been properly calibrated to allow for proper measurement and identification of parasite eggs, cysts, and oocysts.

To calibrate, an ocular micrometer is superimposed over a stage micrometer while the lowest objective (usually 4x) is in place. While the two micrometers are superimposed, line up the left edge of each and locate the position in which one line falls directly over another. At this point, the number of ocular units are compared to the ‘known’ divisions on the stage micrometer so a conversion factor can be obtained.  There are numerous videos and resources on the web which provide sample calculations and visual representation of this process.

Tips on assessing flotation results

Pinworms are common nematode worms detected in reptiles (Figures 4A-E). It is also common to find other nematode eggs, as well as trematode eggs and coccidian oocysts belonging to the genus Eimeria (Figures 4A and F). Information on various species of Eimeria infecting both domestic and wild hosts can be found on the Coccidia of the World Database (eimeria.unl.edu/table.html).

It is not unusual to find pseudo-parasites in fecal flotations, which, though they appear as parasitic elements, are not as such (Figure 5). Common pseudo-parasites include plant matter, arthropod appendages, pollen grains, fungal spores, and other inanimate objects.

The number of parasites found in feces is not always associated with the intensities found in the respective intestine or by the host identity⁵ and parasitic egg counts can often be rather randomly distributed in feces. Intestinal parasites may also be under-represented in the feces due to their location in the intestine, their intensities of infection, presence of adult females shedding eggs, and/or the erratic nature of feces formation and release.⁹

Overall, evaluation of reptile feces for internal parasitic infections should be approached in a similar manner to other taxonomic groups. With proper preparation of materials and calibration of diagnostic tools, the examiner can become adept and confident in identification and evaluation of reptile feces.

[8]

Ashley McGrew, DVM, PhD, DACVM, is an associate professor at Colorado State University’s College of Veterinary Medicine and Biomedical Sciences.  She graduated from the DVM/PhD dual-degree program at CSU and went on to become boarded in parasitology under the ACVM in 2015. She currently teaches veterinary parasitology and serves as Section Head of Parasitology within the CSU Veterinary Diagnostic Laboratories. She can be reached at ashley.mcgrew@colostate.edu[9].

Miranda Sadar, DVM, DACZM, is an assistant professor at Colorado State University’s College of Veterinary Medicine and Biomedical Sciences. She graduated from CSU with a BSc in zoology before earning a DVM degree from Colorado State University. Dr. Sadar completed an internship in zoological, zoological companion animal, and wildlife medicine at the University of Saskatchewan’s Western College of Veterinary Medicine. After finishing a two-year fellowship in wildlife medicine at the Wildlife Center of Virginia, she moved to Davis, Calif., and completed a residency in zoological medicine at the University of California. Sadar is a board-certified specialist in zoological medicine with a focus on zoological companion animals. She can be reached at miranda.sadar@colostate.edu[10].

Tyler Yamashita, BSc., is a research associate at Colorado State University’s Center for Vector-Borne Infectious Diseases (CVID) and Veterinary Diagnostic Laboratory. Yamashita, who graduated from CSU with a BSc. in biological sciences and a minor in entomology, is a rising first-year at CSU’s College of Veterinary Medicine. He can be reached at tyster45@colostate.edu[11].

References

¹ Ras-Norynska M & Sokol R. (2015). Internal parasites of reptiles. Ann Parasitol, 61(2):115-7.

² Galosi L, Attili AR, Perrucci S, Origgi FC, Tambella AM, Rossi G, Cuteri V, Napoleoni M, Mandolini NA, Perugini G, Loehr VJT. (2021). Health assessment of wild speckled dwarf tortoises, Chersobius signatus. BMC Vet Res, 17(1):102.

³ Castillo GN, Acosta JC, Gonzalezrivas CJ, & Ramallo G. (2020). Checklist of nematode parasites of reptiles from Argentina. Annal Parasitol, 66(4):425-432.

⁴ Smales LR. (2007). Acanthocephala in amphibians (Anura) and repiles (Squamata) from Brazil and Paraguay with description of a new species. Journal of Parasitology, 93(2):392-398.

⁵ Wolf D, Vrhovec MG, Failing K, Rossier C, Hermosilla C, Pantchev N. (2014). Diagnosis of gastrointestinal parasites in reptiles: comparison of two coprological methods. Acta Vet Scand, 56(1):44.

⁶ Schmidt V, Dyachenko V, Aupperle H, Pees M, Krautwald-Junghanns M-E & Daugschies A. (2008). Case report of systemic coccidiosis in a radiated tortoise (Geochelone radiata). Parasitol Res, 102:431-436.

⁷ Rom B, Kornas S, Basiaga M. (2018). Endoparasites of pet reptiles based on coprosopic methods. Ann Parasitol, 64(2):115-120.

⁸ Foreyt, W.J. (1989) Diagnostic parasitology. Veterinary Clinics of North America Small Animal Practice, 19 (5), 979-1000.

⁹ Jorge F, Carretero MA, Roca V, Poulin R, Perera A. (2013). What you get is what they have? Detectability of intestinal parasites in reptiles using faeces. Parasitol Res, 112(12):4001-7.

Source URL: https://www.veterinarypracticenews.ca/practical-tips-for-reptile-fecal-exams/

---