Internal Parasite Diagnosis in Small Animals
Diagnosis of internal parasites in small animals is typically performed by examination of feces for parasite eggs. Fecal samples should be fresh, preferably collected from the rectum using a fecal loop. Specimens should be submitted to a diagnostic laboratory in a sealed container, labeled with proper identification. Specimens should be fixed in 10% formaldehyde solution or sent chilled. Other preservative solutions (eg, sodium acetate formalin, polyvinyl alcohol, available in commercial mailer kits) better preserve protozoa and facilitate special staining (eg, trichrome, iron hematoxylin).
Routine examinations should be done by both direct fecal smear and fecal flotation. Direct smears are prepared by mixing feces (an amount of feces fitting on one-half the tip of a wooden applicator stick) with 1 drop each of saline (for motile organisms), and Lugol's iodine stain (to see internal structures, eg, those of Giardia), covering with separate coverslips on the same slide. Flotation methods concentrate diagnostic stages and provide a cleaner final preparation, using about 2 g feces. Sugar (specific gravity [sg] 1.27) and sodium nitrate (sg 1.39) are commonly used flotation media. Zinc sulfate flotation (sg 1.18), repeated on 3 consecutive days, is the method of choice for revealing Giardia cysts, which are intermittently shed in feces. Motile nematode larvae may be collected and concentrated by Baermann sedimentation; a convenient method for small animals is to place ~2 g feces on several sheets of cotton gauze in a tea strainer suspended in the wide end of a conical funnel fitted with a rubber tube and stopcock. The funnel is filled with warm tap water or saline for 1 hr before sediment examination. Larvae descend due to gravity and are collected from the end of the funnel for microscopic examination. Special procedures, such as formalin-ethyl acetate sedimentation may be used for parasitic larval stages that do not float well. Sheather's sugar (sg 1.30) can be used to detect small oocysts of Cryptosporidium (4–6 μ) or Toxoplasma (12 μ) by focusing just under the coverslip. Oocysts “float up” to the under-surface of a coverslip placed on the slide for 10 min before examination.
A direct smear using 1 drop of blood can be used to detect motile microfilariae of Dirofilaria immitis, but should not be the sole means of detection. More accurate examination of blood may be done by a modified Knotts' test: 1 mL blood is added to 9 mL of 2% formalin solution in a 15 mL tube and centrifuged at 1,500 rpm. The supernatant is discarded and a drop of methylene blue stain is mixed with the sediment “button,” which is pipetted onto a microscope slide, covered with a coverslip, and examined to differentiate microfilariae of D immitis from those of the nonpathogenic Dipetalonema reconditum. Commercial filtration and staining procedures are effective alternatives to the modified Knotts' test. Animals on heartworm preventive become amicrofilaremic, and an occult heartworm test using a commercially available ELISA for circulating uterine antigen of adult female Dirofilaria is the method of choice for treated dogs. Feline dirofilariasis cannot be reliably diagnosed by microfilaremia or antigenemia tests because heartworm numbers are typically too low; antibody titers to D immitis are used to detect prior exposure and possible current infection in cats.
Internal Parasite Diagnosis in Livestock
Fresh fecal samples from livestock should be collected from pasture or, preferably, per rectum using plastic gloves. Samples should be placed in a properly identified, sealed specimen jar. A representative number of herd samples should be collected from a minimum of 10 animals to account for the typical high individual variation in numbers of eggs shed. Samples can be combined after thorough mixing to enable examination of a single herd composite sample.
Quantitative fecal egg counts by the modified Wisconsin centrifugal flotation procedure or similar methods can be used to estimate relative infection burden for nematodes of the “GI parasite complex” (eg, trichostrongylid larvae of cattle), while also detecting coccidia and other parasites such as lungworm larvae and tapeworms. Three grams of feces are placed in a container, suspended in ∼15 mL water, strained through a gauze square into a 15 mL tube, and centrifuged (1,500 rpm for 3 min). The supernatant is decanted and the sediment mixed with saturated sugar solution, filling the tube enough to form a positive meniscus before placing a 22 × 22 mm coverslip on the lip of the test tube. The tube is centrifuged again at low speed (1,500 rpm for 5 min). The coverslip, with the surface film containing eggs, is removed and transferred to a microscope slide for counting of trichostrongylid and strongylid eggs. The total is divided by 3 to derive eggs/g (EPG). Other parasites are noted, if present, with a general abundance designation of +1 (few), +2 (small number), +3 (large number), or +4 (too numerous to count).
A saturated solution of table salt (sg 1.20) is an inexpensive alternative medium for diagnosis of livestock parasites, although salt may be corrosive to metal. Magnesium sulfate (sg 1.20) is the preferred medium for swine feces. Special slides containing chambers with etched areas of known volume are also used for estimating EPG, especially for small ruminants. Feces in a strained solution (usually saturated salt) are introduced into each chamber with a Pasteur pipette, and the eggs are counted under low-power magnification. A commonly used counting slide is the McMaster slide, which has 2 chambers, each with a volume of 0.15 mL under the etched area. For example, if 3 g of feces are mixed with 42 mL of concentrate solution, then each egg counted is multiplied by 50 to yield the number of EPG in the fecal sample. Acceptable correlation between the EPG and the relative worm burden is often possible in young animals, although low (<5 EPG) or negative counts are typically found in adult animals. In young cattle, which generally have EPG counts 10 times that of adult animals, EPG counts >50 reflect a moderate infection, and EPG counts >500 indicate a heavy burden and a need for treatment.
Because fluke eggs do not float readily, quantitative fecal sedimentation procedures are usually used. Two grams of feces are mixed with 35 mL soapy solution (2% liquid detergent) and strained through gauze into a 50-mL centrifuge tube. The tube is filled with soapy water and allowed to stand for 3 min, after which ½ of the supernatant is discarded. This is repeated 2–3 times until the supernatant is clean. All but 15 mL is poured off, 2 drops of new methylene blue are added, and the eggs counted with a dissecting microscope in a gridded Petri dish or by examining several coverslipped microscope slides. The eggs of the liver fluke, Fasciola hepatica, can be differentiated from those of Paramphistomum spp, the rumen fluke, by the golden color, more barreled shape, and slightly larger size of eggs of F hepatica versus the gray color, more pointed end, and smaller size of the usually nonpathogenic rumen fluke. Commercial sieve-sediment kits can reduce sample preparation time by 50%. In cattle, Fasciola EPG counts >3 suggest economic losses; EPG >10 may be associated with clinical signs.
Examination for Ectoparasites
Animals with dermatoses should be evaluated by examining for ectoparasites or evidence of their presence. For example, fleas may not be seen on a cat or dog, but small black flecks of flea excrement that produce a reddish stain when placed on a wet paper towel may be noted. Skin must sometimes be scraped to diagnose parasites. A scalpel blade is used for the deep scrapings (until blood oozes) needed to demonstrate parasites that live in burrows (eg, Sarcoptes) or hair follicles (Demodex spp). The scraped material is placed in a drop of mineral oil on a slide, and the entire area under the coverglass is scanned under low-power magnification. A few drops of 10% potassium hydroxide solution may be added to clear debris and allow better visualization.
Last full review/revision March 2012 by Charles M. Hendrix, DVM, PhD