1: Fecal Examination for the Diagnosis of Parasitism

Fecal Examination for the Diagnosis of Parasitism

The fecal examination for diagnosis of parasitic infections is one of the most common laboratory procedures performed in veterinary practice. Relatively inexpensive and noninvasive, fecal examination can reveal the presence of parasites in several body systems. Parasites inhabiting the digestive system produce eggs, larvae, or cysts that leave the body of the host by way of the feces. Occasionally, even adult helminth parasites may be seen in feces, especially when the host has enteritis. Parasitic worm eggs or larvae from the respiratory system are usually coughed into the pharynx and swallowed, and they too appear in feces. Mange or scab mites may be licked or nibbled from the skin, thus accounting for their appearance in the feces. Many parasitic forms seen in feces have characteristic morphologic features that, when combined with knowledge of the host, are diagnostic for a particular species of parasite. On the other hand, certain parasites produce similar eggs or oocysts, and cannot be identified to the species level (e.g., many of the strongylid‐type eggs from livestock). Fecal examination may also reveal to a limited extent the status of digestion, as shown by the presence of undigested muscle, starch, or fat droplets.


Fecal exams should be conducted on fresh fecal material. If fecal samples are submitted to the laboratory after being in the environment for hours or days, fragile protozoan trophozoites will have died and disappeared. The eggs of some nematodes can hatch within a few days in warm weather, and identification of nematode larvae is far more difficult than recognizing the familiar eggs of common species. Also, free‐living nematodes rapidly invade a fecal sample on the ground, and differentiation of hatched parasite larvae from these free‐living species can be time‐consuming and difficult.

Owners of small animals should be instructed to collect at least several grams of feces immediately after observing defecation. This will ensure the proper identification of the sample with the client’s pet (i.e., a sample from a stray animal will not be collected) and that feces rather than vomitus or other material is collected. The limited amount of feces recovered from the rectum on a thermometer or fecal loop should not be relied on for routine parasitologic examination, since many infections that produce only small numbers of eggs will be missed. Owners should be instructed to store fecal samples in the refrigerator if the sample will not be submitted for examination for more than an hour or two after collection.

Feces should be collected directly from the rectum of large animals. This is particularly important when identification of individual animals is needed. Rectal samples are also needed when the sample is to be examined for lungworm larvae or cultured for identification of third‐stage larvae, since contaminating free‐living nematodes and hatched first‐stage larvae of gastrointestinal nematodes may be confused with lungworm larvae. If rectal samples are unavailable, owners should be asked to collect feces immediately after observing defecation. The process of development and hatching of common strongylid eggs can be slowed by refrigeration. Development is also reduced when air is excluded from the sample by placing the collected feces in a plastic bag and evacuating or pressing out the air before sealing the bag.


If collected feces cannot be examined within a few hours, the sample should be refrigerated until it can be tested. Feces should not be frozen, because freezing can distort parasite eggs. If a sample needs to be evaluated for the presence of protozoan trophozoites like Giardia and trichomonads, it should be examined within 30 minutes after collection. The trophozoite is the active, feeding form of the parasite and is not adapted to environmental survival; it dies soon after being passed in the feces.

Increasingly, veterinary practitioners in the United States are using reference laboratories for routine diagnostic tests for parasite infection. Specific laboratory instructions for age, storage and transportation of samples to commercial labs should be followed. In general, when fresh fecal material is submitted to another laboratory for examination, it should be packaged with cold packs. In some cases, preservation of samples may be preferred. Helminth eggs can be preserved with a volume of 5%–10% buffered formalin equal to that of the sample. Formalin fixation also inactivates many other infectious organisms that may be present. Special fixatives, such as polyvinyl alcohol (PVA), are required to preserve protozoan trophozoites and are not routinely used in veterinary practices.

Slides prepared from flotation tests do not travel well, even if the coverslip is ringed with nail polish, since hyperosmotic flotation solutions will usually make parasite eggs or larvae unrecognizable within hours of preparing the slide. However, slides from flotation tests can be preserved for several hours to several days by placing them in a refrigerator in a covered container containing moist paper towels to maintain high humidity. It is best to place applicator sticks under the slide to prevent it from becoming too wet.


Before performing specific tests on the fecal sample, its general appearance should be noted; consistency, color, and the presence of blood or mucus may all be indicative of specific parasitic infections. Hookworm disease in dogs, for example, commonly produces dark, tarry feces, whereas diarrheic feces caused by whipworms may contain excess mucus and frank blood. The presence of adult parasites or tapeworm segments should also be noted.

Fecal Flotation

The technique most commonly used in veterinary medicine for examination of feces is the fecal flotation test. This procedure concentrates parasite eggs and cysts while separating them from much of the sample debris. Fecal flotation is based on the principle that parasite material present in the feces is less dense than the fluid flotation medium and thus will float to the top of the container, where it can be collected for microscopic evaluation. Flotation tests are easy and inexpensive to perform, but in busy practices the choice of flotation solution and test procedure often does not receive much consideration, despite the substantial effect these choices can have on the sensitivity of flotation exams.

Choice and Preparation of Flotation Solutions

Many different substances can be used to make flotation solutions. The higher the specific gravity (SPG) of the flotation solution, the greater the variety of parasite eggs that will float. Additionally, studies have shown that fecal flotation tests recover only a portion of each type of parasite egg/cyst in a sample because of variation in individual eggs, binding to debris, and so on. As SPG increases the portion recovered increases, which is an important consideration when the number of eggs in the sample is low. However, as SPG increases, more debris will also float, and the risk of damage to eggs from the hyperosmotic solution also increases. These factors limit the range of useful flotation solutions to SPG ranging from approximately 1.18 to 1.3. Both salt and sugar flotation solutions are commonly used in veterinary parasitology and provide flotation for common parasites with lower specific gravities than the flotation solution (Table 1.1).

Salt solutions are widely used in flotation procedures. A common flotation solution used in the United States is a commercially available sodium nitrate solution (SPG 1.20). This solution will float common helminth eggs and protozoan cysts. The commercial solution is not a saturated solution of sodium nitrate (SPG 1.33). Slides prepared with any salt solution need to be examined relatively quickly after they are prepared because crystals form as slides dry and parasites may be damaged, making them more difficult to identify.

Zinc sulfate (ZnSO4) solution at a SPG of 1.18–1.2 is another salt flotation solution. It is preferred at SPG 1.18 for recovery of Giardia, but recovers a higher proportion of other parasites at SPG 1.2 and is probably used more frequently at this SPG. It is commercially available and when water is added to the purchased salt solution as directed, the resulting SPG is 1.2.

Table 1.1. Approximate specific gravity of some common helminth eggs

Sources: From 1David E., and Lindquist W. 1982. Determination of the specific gravity of certain helminth eggs using sucrose density gradient centrifugation. J. Parasitol. 68:916–919; 2Norris J., Steuer A. et al. 2018. Determination of the specific gravity of eggs of equine strongylids, Parascaris spp., and Anoplocephala perfoliata. Vet. Parasitol. 260:45–48.

Species Specific gravity
Ancylostoma caninum 1.061
Toxocara canis 1.091
Toxocara cati 1.101
Trichuris vulpis 1.151
Taenia sp. 1.231
Physaloptera 1.241
Parascaris spp. 1.092
Equine strongyles 1.052
Anoplocephala perfoliata 1.062

When detection of Giardia is required, a 33% zinc sulfate solution (SPG 1.18) is recommended because it does not cause the same rapid collapse of cysts seen with other flotation solutions and they are more easily recognized in flotation preparations.

Saturated solutions of sodium chloride (SPG 1.20) and magnesium sulfate (Epsom salts, SPG 1.32) are less widely used but can be easily prepared, are inexpensive, and are effective in floating common helminth eggs and protozoan cysts.

None of these salt flotation solutions will reliably float most trematode eggs, some tapeworm eggs, and very dense nematode eggs or larvae.

Another common solution used in routine flotation exams is Sheather’s sugar solution (SPG 1.25). Because of its relatively higher SPG, Sheather’s solution is also more efficient in recovering helminth eggs than common salt solutions, especially tapeworm and more dense nematode eggs. In addition, it does not distort eggs as rapidly as the salt solutions. Sheather’s solution is specifically recommended for recovery of Cryptosporidium oocysts in fecal samples, but it does not appear to be as effective as 33% ZnSO4 solution for detection of Giardia. Sheather’s solution is inexpensive and easy to prepare and is also commercially available in the United States, but it is more viscous and sticky than salt solutions. The advantages and disadvantages of these solutions are shown in Table 1.2 and instructions for preparing them are given below.

Although the SPG of flotation solutions is not often measured in practices, it can be easily determined with an inexpensive hydrometer from a scientific supply company. A hydrometer will last indefinitely and should be considered part of quality control for the veterinary practice laboratory.

Table 1.2. Comparison of commonly used flotation solutions

Flotation solution Specific gravity Advantages Disadvantages
Sodium nitrate (NaNO3) Commercial product 1.18–1.2 Floats common helminth and protozoa eggs and cysts Does not float most fluke and some tapeworm and nematode eggs
Saturated NaNO3 1.33
Zinc sulfate (ZnSO4) 1.18–1.2 Floats common helminth and protozoa eggs and cysts; preferred for Giardia and some lungworm larvae When used at SPG 1.18 recovers lower proportion of common helminth eggs; does not float most fluke and some some tapeworm and nematode eggs
Saturated sodium chloride (NaCl) 1.2 Floats common helminth and protozoa eggs and cysts Does not float most fluke and some tapeworm and nematode eggs
Saturated magnesium sulfate (Epsom salts) 1.32 Floats common helminth and protozoa eggs and cysts; higher SPG recovers parasites more efficiently Higher SPG will float more debris; does not float most fluke and some tapeworm and nematode eggs
Sheather’s sugar solution 1.20–1.28 Floats common helminth and protozoa eggs and cysts; higher SPG recovers parasites more efficiently; preferred for Cryptosporidium oocysts; generally less damaging than salt solutions Does not float most fluke and some tapeworm and nematode eggs; creates sticky surfaces


  1. Combine 330 g zinc sulfate with water to reach a volume of 1000 mL.
  2. Additional water or zinc sulfate can be added to produce an SPG of 1.18. If zinc sulfate solution is used with formalinized feces, the SPG should be increased to 1.20 and a SPG of 1.2 is often preferred for general use.
  3. Check the SPG with a hydrometer.


  1. Add salt to warm tap water until no more salt goes into solution and the excess settles at the bottom of the container.
  2. To ensure that the solution is fully saturated, it should be allowed to stand overnight at room temperature. If remaining salt crystals dissolve overnight, more can be added to ensure that the solution is saturated. Table salt contains an anticaking compound that does not dissolve and should not be confused with residual sodium chloride crystals. Pickling salt does not contain this compound.
  3. Check the SPG with a hydrometer, recognizing that the SPG of saturated solutions will vary slightly with environmental temperature.


  1. Combine 355 mL (12 fl oz) of water and 454 g (1 lb) of granulated sugar (sucrose). Corn syrup and dextrose are not suitable substitutes.
  2. Dissolve the sugar in the water by stirring over low or indirect heat (e.g., the top half of a double boiler). If the container is placed on a high direct heat source, the sugar may caramelize instead of dissolving in the water.
  3. After the sugar is dissolved and the solution has cooled to room temperature, add 6 mL formaldehyde USP to prevent microbial growth (30 mL of 10% formalin can also be used, with the volume of water reduced to 330 mL).
  4. Check the SPG with a hydrometer.

Flotation Procedures

No matter how the flotation procedure is performed, the principle is the same. After mixing the flotation solution and the fecal sample together, the less dense material eventually floats to the top. This process can occur either by letting the mixture sit on the benchtop for a specified time (passive flotation) or by centrifuging the mixture. Centrifugation makes the flotation occur more rapidly and efficiently, regardless of the flotation solution used. Many practitioners like to use convenient commercial flotation kits that provide a container for collection of the sample and performing the test. However, the convenience of the kits is offset by the loss of sensitivity in the fecal exam procedure. A smaller amount of feces is used and the test cannot be centrifuged.

The increased sensitivity of the centrifugation procedure is particularly important in infections where the diagnostic form of the parasite may be present in low numbers (e.g., Trichuris and Giardia infections in dogs and cats). Centrifugation is also necessary when using 33% ZnSO4 or sugar solution because of the slightly lower SPG of ZnSO4 solution and the high viscosity of sugar solution, both of which retard the flotation process. A veterinary practice that does not centrifuge flotation tests and relies on the traditional benchtop technique substantially reduces the sensitivity of its fecal exams.


This is the best technique for the standard fecal flotation test regardless of the flotation solution used. It is particularly important to use this procedure with Sheather’s sugar and 33% ZnSO4 flotation solutions to ensure that the flotation is effective:

  1. Mix 3–5 g (about 1 teaspoonful) of feces with a small amount of flotation solution in a paper or plastic cup. Cat feces and small ruminant pellets, which are sometimes too hard to break up easily, can be ground with a mortar and pestle or allowed to soak in water until they become softer.

If the sample appears to contain a large amount of fat or mucus, an initial water wash is performed, and water should be used in Step 1. The water wash may be eliminated for most fecal samples of normal appearance.

  1. Strain the mixture of feces and flotation solution (or feces and water if a water wash is performed) through a double layer of cheesecloth or gauze. A tea strainer can also be used.
  2. Pour the mixture into a 15‐mL centrifuge tube. If the rotor on the centrifuge is not angled (i.e., if the tubes hang straight when not spinning), the centrifuge tube can be filled with flotation solution until a reverse meniscus is formed and a coverslip is added (Fig. 1.1). The tube is spun with the coverslip. The centrifugal force generated by the centrifuge will hold the coverslip in place. If the centrifuge has an angled rotor, fill the tube to approximately 10–12 mL (amount that will prevent spilling) and place in the centrifuge.
  3. Spin the mixture in a benchtop centrifuge for about 5 minutes at approximately 500–650 × g (650 × g is 2500 rpm on a 4‐in. rotor), regardless of whether the feces have been mixed with water or with the flotation solution. If a specific g force and speed setting cannot be determined, spinning the tube at the same speed used to separate serum from blood cells is sufficient.

If the initial spin is a water wash, the supernatant should be discarded, the sediment resuspended with flotation solution, and Steps 3 and 4 repeated.


Fig. 1.1 Centrifuge tube filled with flotation solution to the top and coverslip placed in contact with the fluid column.

  1. Allow the centrifuge to stop without using the brake. The slight jerking that results from the use of the brake may dislodge parasites from the surface layer. If preferred, the tube can be allowed to sit for an additional 5–10 minutes to maximize recovery of parasite material that may not have completed flotation through the liquid column to the surface.
  2. Following centrifugation, there are several ways to harvest the surface layer of fluid containing parasite eggs. If the tube has been spun with the coverslip in place, lift the coverslip off the tube and quickly place it on a microscope slide. When the tube is spun without the coverslip, remove the tube from the centrifuge after spinning and place in a test tube rack. Fill the tube with additional flotation solution to form a reverse meniscus. Place a coverslip on the tube and allow it to sit for an additional 5–10 minutes before removing the coverslip and placing it on a slide.

Alternatively, after the centrifuge comes to a stop, gently touch the surface of the fluid in the tube with a glass rod, microbiologic loop, or base of a small glass tube and then quickly touch the rod to a microscope slide to transfer the drop or two of adhering fluid. This procedure will be less efficient than allowing the tube to stand with a coverslip in place.


When a centrifugal flotation procedure cannot be performed, sodium nitrate and saturated salt solutions can be used in a benchtop flotation test, although a number of studies have shown that this procedure will detect significantly fewer parasite infections than the centrifugal flotation. This technique is not recommended for 33% ZnSO4 or sugar flotation tests:

  1. Mix several grams (a teaspoonful) of feces with the flotation solution in a cup.
  2. Strain the mixture through cheesecloth or a tea strainer.
  3. Pour into a test tube, pill vial, or container provided in a commercial kit. Add enough mixture or additional flotation fluid until there is a reverse meniscus on the top of the container. Place a coverslip on the fluid drop at the top.
  4. Allow the flotation to stand for at least 10 minutes, remove the coverslip, place it on a slide, and examine. If the test is allowed to stand for too long, the salt may crystallize on the edges of the coverslip so that it will not lie flat on the slide.


Fecal flotation slides should be scanned using the 10× objective lens of the microscope (since most microscopes also have an eyepiece magnification of 10×, using the 10× objective gives a total magnification of 100×). Although most helminth eggs can be detected with the 4× objective, protozoan parasites are easily missed and this low power should not be used for scanning. The 40× lens should be used when there is uncertainty about the identity of structures on the slide and for scanning slides for Cryptosporidium oocysts. In some practices, to save expense, coverslips are not used. However, slides without coverslips dry out faster, do not have a flat plane of focus, and cannot be examined with the 40× lens, with the result that some parasites will be identified incorrectly or missed entirely.


Egg‐counting techniques are also flotation tests and have several uses in food animals and horses. They can be used to assess the degree to which individual animals or groups are contributing to pasture contaminations with parasites, to assess efficacy of drug treatment (discussed in a later section in this chapter) and, in some cases, to determine relative levels of individual susceptibility to parasite infection.

Egg counts are of limited value in making judgments about the clinical condition of individual animals because many factors affect egg production, including parasite species, individual host immunity, and stage of infection. Also, counts performed on combined samples from a number of animals may not accurately reflect parasitism within that herd.

The easiest quantitative test to perform is the modified McMaster test. This test requires the use of special reusable slides (Fig. 1.2), which can be purchased from several suppliers (including Chalex Corporation, Wallowa, OR [www.vetslides.com]; Focal Point, www.mcmaster.co.za; JA Whitlock & Co., Eastwood, New South Wales, Australia [www.whitlock.com.au]). The capacity of the counting chamber and the number of chambers counted per sample affect the detection level of the test. Saturated salt solutions are usually used as the flotation solution in this test.

As it is most commonly used, the modified McMaster test has a detection level of 25 or 50 eggs per gram (EPG) of feces. This level is acceptable in many situations since parasite control programs do not usually require detection of lower egg numbers. However, the accuracy of the McMaster test is reduced when egg counts are at the lower limits of detection. When egg counts of less than 200 EPG are expected in a group of animals being evaluated (e.g., in adult cattle or camelids) or when it is important to detect low numbers of eggs more accurately (e.g., in fecal egg count reduction tests [FECRTs]), a modification of the test is appropriate. The sensitivity and precision of the McMaster test can be improved by using slides that allow examination of larger quantities of the egg/flotation solution mixture, or by counting additional aliquots of the sample.

Alternatively, other procedures with lower detection limits can be used including the Wisconsin sugar flotation test (double centrifugation procedure) or a modified Stoll egg‐counting test. These procedures permit detection of fewer than 10 EPG of fecal material but are more time‐consuming to perform. Another procedure, the mini‐FLOTAC test, has been developed in Europe. Several studies have demonstrated greater precision and accuracy of the mini‐FLOTAC compared to McMaster and Wisconsin tests but at this time it has limited availability and has been used in the United States primarily in research.


Fig. 1.2 McMaster slide used in the modified McMaster procedure for quantitative egg counts.

Another commercially available test for determining equine fecal egg counts is Parasight (Lexington, KY [www.parasightsystem.com]). This system provides counts of equine strongyle and Parascaris spp. eggs. An equine sample is mixed with reagents and fluorescence‐imaging is used with a software counting algorithm to produce a fecal egg count.

Regardless of the procedure used to count parasites, the most important element is consistency. Each step of the procedure should be performed in the same way for every sample.

Modified McMaster test.

  1. For ruminants, combine 4 g of fecal material with 56 mL of flotation solution to yield a total volume of 60 mL. The test can also be performed with 2 g of feces and 28 mL of flotation solution when only small amounts of feces are available. For horses, it is standard in the United States to use 4 g of feces and 26 mL of flotation solution. See Note 1 below for modified calculations. Portable electronic scales that weigh in 0.1‐g increments are widely available and inexpensive. If a method of weighing feces is not available, adding manure to the measured flotation solution until the final desired volume is reached can also be used (e.g., add manure to 26 mL of fluid to a total volume of 30 mL). This method would be more accurate with horse or cattle manure compared to the pelleted manure of small ruminants or camelids.
  2. Mix well and strain through cheesecloth or a tea strainer. The mixture does not have to be strained, but it will be much easier to read the slide if large pieces of debris are removed. An alternative to straining is to use a filter pipette to transfer material to the counting chamber (a pipette with 12 mesh/cm wire mesh at the end is available from JA Whitlock & Co. [www.whitlock.com.au]).
  3. Immediately fill each chamber of the McMaster slide with the mixture using a pipette or syringe. The entire chamber must be filled, not just the area under the grid. If large air bubbles are present, remove the fluid and refill.
  4. Allow the slide to sit for at least 5 minutes before examining to allow the flotation process to occur. There has been limited investigation of the maximum amount of time slides can be allowed to sit before reading and there is no standard recommendation. Allowing slides to sit for an hour does not seem to alter results.
  5. Look at the slide with the 10× lens, focusing on the top layer, which contains the air bubbles. At this level, the lines of the grid will also be in focus. Count eggs, oocysts, and any other parasite stages, in each lane of both chambers. Each type of parasite should be counted separately. In some cases, eggs can be identified to genus or perhaps to species (e.g., Strongyloides, Trichuris, and Nematodirus), whereas others must be counted as a category of parasites (coccidia, strongylid eggs).

To determine the number of parasite EPG of feces, add the counts for both chambers for each parasite. The most commonly used McMaster slides in the United States are calibrated so that the number of eggs counted in a single chamber represents the number present in 0.15 mL of fecal mixture. If both chambers are counted and the results are added, the total represents the number of eggs present in 0.3 mL, which, for example, is 1/200th of a total volume of 60 mL; therefore, the number of eggs counted must be multiplied by 200. However, if a total of 4 g of feces was used in the test, the result must be divided by 4 to yield EPG of feces. Multiplying by 200 and dividing by 4 is equivalent to multiplying the number of eggs counted by 50. Therefore, each egg observed represents 50 EPG in the final count. The same level of detection can be achieved by using 2 g of fecal material and 28 mL of flotation solution. The smaller amount of feces may be preferred when evaluating small lambs or kids.

Additional Notes

  1. If a detection level of 25 EPG is desired, the McMaster test should be performed with 4 g of feces and 26 mL of flotation solution (results are then multiplied by 100 and divided by 4 as described earlier). Other combinations of manure and flotation solution can also be used with appropriate calculations using the formula
    where T = total volume of feces/flotation solution mixture, V = volume of aliquot examined in slide, and F = grams of feces used.
  2. If pelleted feces are very hard, 2–5 mL of water can be added first and left to soften manure for at least an hour. The flotation solution is then added to the softened manure (flotation solution volume reduced by the amount of water used to soften feces).
  3. A variety of methods can be used to homogenize feces and liquid. As alternatives to mixing with a tongue blade or other utensil, some laboratories use a mechanical or handheld kitchen mixer to homogenize feces and water/flotation solution. Shaking manure and fluid in a jar with glass beads has also been used, but as previously stated, whatever method is used, it should be consistent across samples.
  4. In another procedure for the McMaster test, feces is mixed initially with water, strained, and centrifuged, and the supernatant is discarded. Flotation solution is added to resuspend the sediment and mixed, and the mixture is used to fill the counting chambers. For example, 3 g of feces is mixed with 42 mL of water, strained and the fluid used to fill a 15‐mL centrifuge tube, which is then centrifuged at 300–650 × g for 2 minutes. The supernatant is then discarded and flotation solution is added to partially fill the tube, which is either shaken or stirred to resuspend the sediment. Additional flotation solution is added to fill the tube, and the counting chamber is filled. If the eggs in a volume of 0.3 mL are counted, the number of eggs seen is multiplied by 50 to give the number of eggs per g. The EPG can also be calculated using the formula described in Note 1. This procedure is the most effective for reducing debris but increases the time required to perform each test.


The Mini‐FLOTAC device and the Fill‐FLOTAC device, which are used together in the mini‐FLOTAC procedure, are available in North America from the University of Georgia. The test should be performed as described in the brochure provided with the test device. This procedure is generally more time‐consuming than the basic modified McMaster procedure but is more accurate and precise.

Wisconsin, Cornell‐Wisconsin egg‐counting test (double centrifugation procedure).

The Wisconsin egg‐counting test or double centrifugation flotation test is used to quantitate eggs when low EPG are expected. Unlike the McMaster test, where eggs are counted in an aliquot of the mixture of feces and flotation solution, the Wisconsin test preparation collects eggs from the entire fecal sample/flotation solution mixture. Because it is time‐consuming and difficult to accurately count numerous eggs on a slide with no grid this test is not suited to many circumstances where a quantitative count is needed. Any flotation solution can be used in this test:

  1. Combine 1–5 g feces and 12–15 mL of water in a cup. Mix and strain into another cup, rinsing first cup with 2–3 mL of water and straining, pressing the liquid through. Pour into a 15 mL centrifuge tube.
  2. Centrifuge (properly balanced) at 300–650 × g for 5–10 minutes.
  3. After spinning, discard the supernatant and resuspend the pellet in flotation solution.
  4. Either spin the tube with a coverslip in place or allow additional incubation after spinning as described for the centrifugal flotation procedure.
  5. Remove the coverslip and place on a glass slide.
  6. Examine with the 10× objective lens.
  7. Count and record the number of each type of parasite egg/cyst seen, systematically scanning the slide and counting eggs or cysts of each parasite species or group separately. Care must be taken to ensure that each microscope field on the coverslip is examined once but only once so that no eggs are missed or counted twice.

This technique allows the quantification of less than 1 EPG of feces.

Additional Notes

  1. If desired, the initial water wash can be omitted and the sample can be mixed directly with flotation solution and then centrifuged.
  2. Alternatively, 22 mL of flotation solution is mixed with 5 g of feces, and the resulting mixture is divided between two tubes. This modification increases the accuracy of the procedure.

Modified Stoll test.

There are many modifications for the Stoll test, depending on the level of detection desired. Like the McMaster test, the modified Stoll egg‐counting procedure is based on determining the number of eggs present in an aliquot of the prepared feces/flotation solution mixture. Any of the fecal flotation solutions can be used in this procedure.

  1. Combine 5 g of feces and 20 mL of water in a cup.
  2. Mix into a slurry and transfer 1 mL of the mixture to a centrifuge tube. If the mixture is not strained, a widemouthed or mesh‐covered filter pipette is needed to transfer the mixture.
  3. Fill the tube with flotation solution.
  4. Place the tube in the centrifuge and add flotation solution until a slight inverse meniscus is formed.
  5. Place a coverslip on top of the tube. It should contact the mixture without causing any to overflow.
  6. Centrifuge (properly balanced) at 300–650 × g for 10 minutes.
  7. Remove the coverslip and place on a glass slide.
  8. Examine with the 10× objective lens.
  9. Count and record the number of each type of parasite egg/cyst seen, systematically scanning the slide and counting eggs or cysts of each parasite species or group separately. Care must be taken to ensure that each microscope field on the coverslip is examined once but only once so that no eggs are missed or counted twice.

In this test, all the eggs present in 1 mL of the feces/flotation solution mixture were counted following centrifugation. This represents 1/25 of the volume of the mixture so the number of eggs counted multiplied by 25 represents the total number of eggs present, but this must be divided by 5 (grams of feces used) to yield EPG. In this situation, the multiplication factor is 5 and the minimum sensitivity of the test is 5 EPG. If the volume of fluid added to feces in the first step is 45 mL, the final multiplying factor would be 10.

For additional information on quantitative egg‐counting procedures, see references by Verocai et al. (2020), Neilsen and Reinemeyer (2018), Taylor et al. (2015), Coles et al. (2006), and the Ministry of Agriculture, Fisheries and Foods (1986).

Additional Procedures for Fecal Examination

The following procedures are used for identification of specific parasitic infections.

Direct Smear and Stained Fecal Smears

The direct smear is used to identify protozoan trophozoites (Giardia, trichomonads, amoebae, etc.) or other structures that float poorly or are readily distorted by flotation solutions. Because very little fecal material is used, the sensitivity of this test is low. It is not recommended for routine fecal examinations:

  1. Mix a very small amount of feces with a drop of saline on a microscope slide to produce a layer through which newsprint can be read. Saline should be used because water will destroy protozoan trophozoites.
  2. Use a coverslip to push large particles of debris to the side and place the coverslip on the slide. Examine with 10× and 40× magnification. The 100× lens (oil immersion) cannot be used effectively with fecal smears.
  3. If the fecal layer is too thick, it will be impossible to see small, colorless protozoa moving in the field. Movement is the principal characteristic that allows recognition of trophozoites in fresh fecal smears. A drop of Lugol’s iodine will enhance the internal structures of protozoan cysts but will also kill trophozoites present. To maximize the use of this test, it is best to look at an unstained smear before adding iodine.

Fecal smears can also be stained for identification of intestinal protozoa. Several stains can be used for identification of Cryptosporidium, including Ziehl–Neelsen, Kinyoun, carbol‐fuchsin, and Giemsa stains. Trichrome stain is widely used in human medicine for detection of Giardia cysts. In general, however, stains are not used extensively in veterinary practices and are not required for the identification of parasitic organisms. For details on performing these stains, a standard text on human parasitologic diagnosis should be consulted.

Fecal Sedimentation

A sedimentation procedure is used to isolate eggs of flukes, acanthocephalans, and some tapeworms and nematodes whose eggs do not float readily in common flotation solutions. In the simple sedimentation test, tap water is combined with feces and allowed to settle briefly before the supernatant is removed. This allows the removal of fine particulate material, but unlike the flotation exam, sedimentation tests have only limited concentrating ability. Fat and mucus can be removed from the fecal sample if a centrifugal sedimentation exam is performed using ethyl acetate. Unfortunately, ethyl acetate is toxic and very flammable. It should be stored in a flameproof cabinet and used only in well‐ventilated areas. An alternative to ethyl acetate is Hemo‐De, available through Fischer Scientific (www.fischerscientific.com), which is generally regarded as a safe compound and appears to give equivalent results in a centrifugal sedimentation procedure (see Neimester et al. 1987).

The Flukefinder® is a commercially available apparatus for performing sedimentation tests in the laboratory. It utilizes several screens to rapidly remove fecal debris. This device is very useful in practices conducting routine fecal examinations for flukes. Information on the Flukefinder can be obtained at www.flukefinder.com.


  1. Mix about 100 mL of water with about 10 g of feces, strain, and place in a beaker or other container.
  2. Allow mixture to sit for 1 hour and then decant the supernatant.
  3. Add more water, mix, and repeat the sedimentation procedure.
  4. Stir remaining mixture and place a few drops on a microscope slide. If desired, add one drop of 0.1% methylene blue. The methylene blue will stain the background debris blue but will not stain fluke eggs, which will stand out with a yellowish brown color.
  5. Coverslip and scan the slide using the 10× objective lens.

A smaller amount of feces and water can be used, placed in a test tube, and left to sit for 3–5 minutes between decantation steps. Addition of a drop of dishwashing soap to the water used in the test helps to free eggs from surrounding debris.


  1. Mix 1 g of feces with about 10 mL of 10% buffered formalin or water. Pour mixture into a 15‐mL centrifuge tube (with cap) until it is one‐half to three‐quarters full.
  2. Add ethyl acetate (see earlier discussion on safety) or Hemo‐De until the tube is almost full. Because organic solvents may dissolve some plastic centrifuge tubes, it is recommended that glass or polypropylene tubes be used for performing this test.
  3. Cap and shake the tube approximately 50 times.
  4. Centrifuge for 3–5 minutes at about 500 × g (as for centrifugal flotation procedure).
  5. When the tube is removed from the centrifuge, it will have three layers: (1) an upper layer containing ethyl acetate, fat, and debris; (2) a middle layer containing formalin or water and fine particulate matter; and (3) a bottom layer of sediment. Using an applicator stick, loosen the top debris plug that sticks to the sides of the tube, then decant the supernatant, leaving only the bottom sediment.
  6. Resuspend the sediment in a few drops of water or formalin, place one or two drops of the sediment on a slide, coverslip, and examine with the 10× microscope objective.

Baermann Test

The Baermann test is used to isolate larvae from fecal samples and is employed most often to diagnose lungworm infections. It is very important that the fecal sample be fresh. If feces of a grazing animal are being examined and an old sample is used, strongylid or Strongyloides eggs may have hatched, or free‐living nematodes may have invaded the sample, making nematode identification much more difficult. In small‐animal samples, hookworm eggs may hatch very quickly and can be confused with lungworm or Strongyloides larvae. Coprophagy and hunting can also result in larvae of spurious parasites being present in Baermann test preparations. For discussion of identification of nematode larvae see the section in this chapter: “Identification of nematode larvae recovered with fecal flotation or Baermann procedures.”

A further consideration for using a Baermann test in diagnosis is that metastrongyloid lungworms typically show erratic larval shedding patterns. Dramatic day‐to‐day variation in larval shedding increases the chance of false negative Baermann fecal examination results. Therefore, a single negative Baermann result is weak evidence for ruling‐out lungworm infection in an animal showing signs of respiratory disease. Detection sensitivity is increased by doing multiple (at least 3) Baermann examinations.

A Baermann test requires equipment to hold the fecal sample in water so that larvae can migrate out and be collected. This can now be most easily accomplished with the use of a plastic wine glass with a hollow stem. In the absence of disposable wine glasses, the original Baermann apparatus can be used. This consists of a funnel clamped to a metal stand. A short piece of tubing with a clamp is attached to the end of the funnel. Larvae in feces placed either in the bowl of the wine glass or in the funnel migrate out of the sample and fall down into the hollow stem or the tubing above the clamp, where they can be easily collected (Fig. 1.3):

  1. Place at least 10 g of feces in a piece of double‐layer cheesecloth. Gather the cheesecloth around the sample so that it is fully enclosed. Use a rubber band to fasten the cheesecloth, and pass through the rubber band two applicator sticks, a pencil or other object that will rest on the edges of the glass or funnel and suspend the sample. Alternatively, place the sample on a suspended piece of wire mesh or sieve.
  2. Fill the funnel or wine glass with lukewarm water. Make sure that the corners of the cheesecloth do not hang over the edge of the funnel or glass, because they will act as wicks for the water.
  3. Allow the sample to sit for at least 8 hours, preferably overnight.
  4. If using the disposable plastic glass, remove the fecal sample and collect the material at the bottom of the hollow stem using a Pasteur or transfer pipette or syringe. Transfer some of the fluid to a microscope slide, coverslip, and examine with the 4× or 10× objective lens.

    Fig. 1.3 (A) The traditional Baermann apparatus consisting of a suspended funnel with clamped tubing attached. For diagnostic testing of fecal samples, it is much more convenient to perform the Baermann exam with a disposable plastic wine glass (B).

  5. If using the funnel, release the clamp and collect the first 10 mL of fluid into a centrifuge tube. Spin as for a flotation exam, discard the supernatant, and examine the sediment. Alternatively, the very steady handed can carefully loosen the clamp and collect the first three or four drops onto a microscope slide.

Immunologic and Molecular Methods of Parasite Diagnosis

Immunologic methods have been important in the diagnosis of blood and tissue parasites for many years, and they are now being used increasingly for identification of specific parasites in fecal samples. Molecular diagnostic methods are also now being applied to detection of parasites in fecal samples. Although these techniques cannot currently replace morphologic exam of feces as a routine screening procedure for all parasites, they are useful for specific diagnosis of protozoan and helminth parasites that are detected in feces. For the discussion of these procedures, see Chapter 4.


Although the concept of quality control is not often applied to fecal exams, attention to both equipment and training will help ensure that fecal exams are consistently done correctly:

  1. Keep microscopes in good repair. Objective lenses and eyepieces should be routinely cleaned with lens cleaner and lens paper. Have microscopes professionally cleaned and checked every few years.
  2. Check the SPG of flotation solutions with a hydrometer when first prepared to ensure that they will recover parasites effectively. If a batch of solution is used over an extended period, SPG should be checked at least monthly.
  3. Use an ocular micrometer (see section on microscope calibration later in this chapter) to measure structures seen on fecal exams. If possible, recalibrate the microscope at regular intervals.
  4. Make sure that personnel performing the fecal exams are adequately trained. It is not unusual for untrained assistants to be given rudimentary instruction on performing flotation tests and then be assigned to do them. Under these circumstances, it is hardly surprising that air bubbles are identified as coccidia and that smaller parasites are missed entirely.
  5. As a check on the diagnostic accuracy of in‐clinic fecal exams, periodically submit duplicate portions of fecal samples to a diagnostic laboratory. Both negative and positive samples should be submitted.


There are several points to remember in using the compound microscope to examine preparations for parasites:

  1. Use the 10× objective lens of the microscope for scanning slides. This will provide a total magnification of 100× since most microscope eyepieces contain an additional 10× lens. Start in one corner and systematically scan the entire slide. The 40× objective lens (400× total magnification with eyepiece) is useful for closer examination or for looking for very small organisms such as Giardia or Cryptosporidium. The 100× (oil immersion) lens should not be used for flotation preparations. Not only is it likely that flotation solution will contact the lens and possibly damage it, but the pressure of the lens on the coverslip will create currents in the fluid on the slide, keeping everything in motion and making examination of structures difficult.
  2. Most parasite eggs and larvae have little or no color and do not stand out well, so it is important to maximize the contrast between the parasites and their backgrounds. If a microscope has a substage condenser it can be used to increase contrast by adjustment of the condenser diaphragm or placement of the condenser in a low position. Even when a substage condenser is not available, reducing the intensity of light projected on the slide is generally advisable, either by decreasing the microscope rheostat setting or by reducing the aperture of the iris diaphragm. A higher power used for close examination will, of course, require an increased amount of light.
  3. When reading a slide, it is helpful to focus up and down with the fine focus to change somewhat the plane of focus. Frequently, worm eggs will be at a slightly different level than protozoan cysts or oocysts, and a small manipulation of the fine focus may make structures more readily visible (Figs. 1.4–1.6).

Microscope Calibration

The ability to measure the size of parasitic organisms and structures is very helpful when identifying unusual parasites or where different organisms are similar in appearance but differ in size. For measurements, a micrometer disc, also known as a reticle (Fig. 1.7), is inserted into the ocular tube of the microscope and calibrated against a known reference in the form of a stage micrometer (Fig. 1.8). Each objective lens of the microscope must be individually calibrated with the ocular lens/micrometer combination to be used, and the calibrations should be posted close to the microscope for easy reference. The calibration will be accurate only for that particular microscope ocular and objective combination. Even if each lens is not calibrated with the stage micrometer, the ocular grid will provide a consistent reference against which to compare objects seen in fecal samples. These ocular micrometer discs and stage micrometers are not expensive and can be purchased from scientific catalogs that include microscope equipment.


Figs 1.4–1.6 The importance of small changes in the microscope focus can be seen in these three photos of the same field in a canine fecal flotation test. In each case, a slight manipulation of the fine focus brings a different parasite into clearer view (first Cystoisospora oocysts, then hookworm eggs, and finally Toxocara eggs) since each egg or oocyst type may be present on a slightly different level.


Fig. 1.7 A typical ocular micrometer of 50 divisions. The divisions have no meaning until calibrated against a stage micrometer.


Fig. 1.8 A typical stage micrometer of 1 mm total length. Each division represents 10 μm.


Fig. 1.9 Appearance at 40× of an ocular micrometer being calibrated with a 10 μm/division stage micrometer. Note the conjunction of line 11 of the stage micrometer with line 49 of the ocular micrometer.

Calibration of the 40× objective illustrates the procedure for calibration of the micrometer:

  1. To calibrate the 40× objective, place the stage micrometer on the stage of the microscope and focus until the lines are sharp. In the example (Fig. 1.9), the stage micrometer is 1 mm (1000 μm) long and is divided into 100 parts; thus, each small division of the stage micrometer represents 10 μm.
  2. Superimpose any convenient numbered line of the ocular micrometer (usually the 0 mark) on a convenient line of the stage micrometer (the first large line in the example). The field should now resemble Figure 1.9.
  3. Find the two lines that are exactly superimposed. In the example, line 49 of the ocular micrometer falls exactly on line 11 of the stage micrometer. Thus, 49 divisions of the unknown ocular micrometer represent 11 divisions, each 10 μm in length, for a total of 110 μm. To complete the calibration, divide 110 μm by 49 divisions, resulting in a calibration factor of 2.24 μm per division for the ocular micrometer in the example.
  4. Repeat this procedure for each objective lens to be calibrated on the microscope.

To use the calibrated microscope, superimpose the ocular micrometer scale on an egg or cyst and count the number of divisions subtended by the specimen, for example, 12. Multiply 12 by the calibration factor (2.24 for the 40 × lens in the example; 12 × 2.24 = 26.88 μm, the size of the object measured).


Fecal samples may contain deceptive “pseudoparasites” and “spurious parasites.” Pseudoparasites are ingested objects that resemble parasite forms; these include pollen grains, plant hairs, grain mites, mold spores, and a variety of harmless plant and animal debris (Figs. 1.101.15). “Spurious parasites” are parasite eggs or cysts from one species of host that may be found in the feces of a scavenger or predator host as the result of coprophagy or predation (Figs. 1.161.18). One of the best ways to avoid misidentifying these pseudo‐ and spurious parasites is to appreciate the variety of parasites that normally infect a host species. If a fecal sample contains a possible pseudoparasite or spurious parasite, it is best to repeat the examination with another sample collected at a later time. To limit opportunities for coprophagy or predation leading to ingestion of additional pseudo‐ and spurious parasites, small animals (dogs and cats) should be confined or leash walked only for 2–3 days prior to collection of the second sample.


Fig. 1.10 Examples of pseudoparasites. (A) Pine pollen is a common pseudoparasite found in fecal samples of many animals (400×). (B) Adult free‐living nematodes are also commonly found in fecal samples collected from the ground. These nematodes can rapidly invade fecal material. The presence of adults and variation in size and morphology (indicating different stages of the life cycle) are helpful in distinguishing these worms from parasite larvae.


Fig. 1.11 Examples of pseudoparasites. (A) In this ovine fecal sample, both a strongylid egg and a pseudoparasite (arrow) are present. Characteristics helpful in the recognition of pseudoparasites are lack of clear internal structure and discontinuities in the outer layer. (B) Pseudoparasite, probably a pollen grain (400×).


Fig. 1.12 Examples of pseudoparasites. (A) Insect hair from the feces of an insectivorous bird. Insect and plant hairs may be confused with worms but have no internal structure. (B) This artifact in ruminant feces appears to have structures resembling the hooks of a tapeworm embryo, but there is no distinct embryo and the outer layer is poorly defined with projections that are variable in size and shape (40×).


Fig. 1.13 Examples of pseudoparasites. (A) Plant hairs and other fibrous material can resemble nematode larvae. They can be present in a variety of shapes and colors, but can usually be easily differentiated from nematodes because they lack clear internal structures like a digestive tract. Also, while one end is tapered, the other end often looks as though it has been broken off another structure. (B) This photo shows Saccharomycopsis guttulatus, a nonpathogenic yeast common in rabbits and seen occasionally in dogs (400×).


Fig. 1.14 Examples of pseudoparasites. (A) Free‐living mites that contaminate animal feed can be found in fecal flotation procedures. Unlike many parasitic mites, free‐living species lack specialized structures on their legs (suckers etc.) for adhering to the host. (B) Eggs from free‐living mites will also float in flotation solution. They are usually very large (>100 μm). Developing legs of the mite can sometimes be seen inside the egg (arrow).


Fig. 1.15 Examples of pseudoparasites. Among the most common pseudoparasites found in feces are insect larvae, which may still be alive and moving when presented. Insect larvae may be ingested in food or, in the case of fly maggots, like the one shown here, eggs that are laid on the feces hatch rapidly in hot weather. Spiracles are present on the posterior (right) end of the segmented larva.


Fig. 1.16 Spurious parasites are parasite eggs or cysts from another host that are acquired through predation or coprophagy and have merely passed through the digestive tract of the animal being tested. (A) Tapeworm egg found in a fecal sample from a calf. Although the configuration of hooks inside this egg clearly identifies it as a tapeworm, it is most likely a rodent or bird tapeworm egg. (B) Spurious parasites are common in samples from dogs that ingest fecal material. Eggs of livestock strongylid species can be found in feces of manure‐eating dogs. Ruminant and equine strongylid eggs look like canine hookworm eggs but are larger and ruminant coccidia can usually be differentiated from dog and cat species based on size and shape.


Fig. 1.17 Examples of spurious parasites. (A) Large cyst of Monocystis, a protozoan parasite of earthworms found in the feces of a snake that feeds on earthworms (100×). (B) Individual Monocystis oocysts that have been freed from a large cyst like the one shown in Figure 1.17A. These individual Monocystis oocysts are common pseudoparasites (400×).

Photo B courtesy of Dr. Yoko Nagamori, College of Veterinary Medicine, Oklahoma State University.


Fig. 1.18 Examples of spurious parasites. (A) Adelina sp. oocyst in a canine fecal sample. The oocysts of this genus are coccidia of insects and oligochetes and contain eight sporocysts. (B) Feline fecal sample containing two eggs from a feline parasite (Toxocara) and a single egg of a spurious parasite, Trichosomoides, a rodent parasite that is present as a result of hunting activity.

Photos courtesy of Dr. Manigandan Lejeune, Animal Health Diagnostic Center, Cornell University.


Nematode larvae are passed in the feces of animals infected with various species of lungworms (Aelurostrongylus abstrusus, Angiostrongylus vasorum, Crenosoma vulpis, Dictyocaulus spp., Filaroides hirthi, Muellerius capillaris, Oslerus osleri, Protostrongylus spp., and others) or the intestinal threadworm, Strongyloides stercoralis. Accurate identification of nematode larvae detected on fecal flotation or Baermann tests tends to be a challenge for the veterinary laboratory diagnostician. In many cases where larvae are detected on fecal flotation, the damage due to the effects of high SPG flotation media obscures the larval morphology to the point that identification is not possible (Fig. 1.19). Therefore, the Baermann technique is the preferred method to recover first‐stage nematode larvae from feces except in the case of O. osleri or F. hirthi infection in dogs (Figs. 1.94, 1.95). The Baermann technique is effective in recovering larvae that are vigorous and able to move out of the fecal matter. The larvae present in feces of dogs infected with Oslerus and Filaroides are sluggish and unable to migrate out of the feces. Therefore, zinc sulfate centrifugal flotation is the recommended method for the detection of larvae in the feces of dogs infected with these lungworms.

A further complication in larval identification may occur when there is a loss of sample integrity due to improper collection. Fecal samples that are not collected immediately after deposit on the ground may be invaded by free‐living soil or plant parasitic nematodes. The challenge of sorting out these nematodes from the true parasitic ones is beyond the training and experience of most veterinary laboratory diagnosticians. In addition, hookworm, strongyle, or trichostrongyle eggs, if present in feces, can develop and hatch in a short time under warm conditions, resulting in the detection of larvae that will be difficult to distinguish from those parasites normally passed as larvae in the feces. In small‐animal practice where pet owners collect the fecal sample, the clients must be given guidance as to the requirements for a proper fecal sample. In the case of dogs, clients should be instructed to collect the fecal sample immediately after deposit and place it in an airtight, leakproof container. If submission to the veterinarian cannot occur within several hours of collection, the sample should be refrigerated at 4°C. In the case of cats, the litter pan should be cleaned and the next fecal sample observed in the pan should be collected and handled as above. Ruminant or horse samples should be collected from the rectum. If clients are collecting samples from the ground, they should be instructed to avoid collection of the portion of manure in direct contact with the soil. Lastly, a further complication in test evaluation can be the presence of spurious parasites acquired through predation or coprophagy.


Fig. 1.19 Larva detected on fecal flotation from a dog infected with lungworm. The larva is damaged due to the osmotic pressure of the high specific gravity flotation fluid. Loss of morphologic detail to this degree prevents specific identification.

The first question in the decision tree when evaluating nematode larvae is: parasite or free‐living? Parasite larvae range in size from 150 to 400 μm and their simple anatomy consists of a mouth opening leading to a buccal tube, esophagus, intestine, and anus. There may also be a discernable genital primordium. Free‐living/soil/plant nematodes often occur in multiple life stages (from egg to adult), and size measurements are highly variable. The presence of adult female (eggs in the uterus, vaginal opening—Fig. 1.20) or adult male (spicules—Fig. 1.21) worms or the presence of an oral stylet in the buccal tube (Fig. 1.22) indicates that the sample may have been invaded by free‐living nematodes. Unfortunately, Strongyloides spp. have a free‐living generation that will develop if the sample is incubated and therefore are also a possibility when adult stages are recovered in a fecal sample. Detection of adult worms in the sample is an indication that the animal should be resampled and a fresh fecal sample should be submitted.

Another source of potential confusion in identifying larvae in feces is particularly common in coprophagic dogs, and can also occur in dogs or cats that are allowed to hunt. First‐stage larvae present in feces or a prey animal will pass through the gastrointestinal tract intact. Depending on the timing of ingestion, the larvae may still be vigourously motile when recovered as a spurious parasite on Baermann examination. Reports of Aelurostrongylus abstrusus infection in the dog have all been based on detection of L1 in feces and are most likely false positive due to the ingestion of cat feces. Familiarity with common lungworms of other species will be helpful in recognizing the possibility of spurious parasitism.


Fig. 1.20 Vaginal opening (arrow) of a free‐living adult female nematode recovered from the feces of a dog. The feces were left on the ground long enough prior to collection to allow free‐living soil nematodes to invade the sample.


Fig. 1.21 Tail of an adult male free‐living nematode recovered from an improperly collected fecal sample of a dog. Note the chitinized spicules (arrow) at the cloacal opening.


Fig. 1.22 Anterior end of a plant parasitic nematode recovered from an improperly collected fecal sample of a dog. Note the oral stylet (arrow) in the buccal chamber. The stylet is a daggerlike structure used in the feeding process to pierce plant roots. No parasitic first‐stage larvae have this structure.

Detection of larvae on microscopic examination of a slide prepared from a Baermann test is facilitated by the eye‐catching vigorous motion of the larvae. However, once detected, a careful evaluation of the morphologic features is not possible in actively motile larvae. Therefore, it is necessary to kill them in a way that does not damage the morphology. Larvae are best killed by adding a drop of dilute Lugol’s iodine (the color of weak tea) to the edge of the coverslip. The iodine will be slowly drawn across the coverslip resulting in the death of the larvae. Alternatively, the larvae can be heat killed by passing the coverslip over the flame of a Bunsen burner to effect (several to many times). Larvae recovered on fecal flotation may or may not be already dead. Fecal flotation slides should be viewed as quickly as possible since the larval damage due to osmotic pressure will progressively worsen over time. Differentiation of the various parasitic nematode first‐stage larvae is based on overall size and the morphology of the esophagus and tail. First‐stage larvae of intestinal parasites (i.e., S. stercoralis or hookworm–strongyle–trichostrongyle eggs that have hatched) can be differentiated from the numerous lungworm larvae based on the presence of a distinct rhabditiform esophagus (Figs. 1.23A, 1.24, and 1.26). The rhabditiform esophagus consists of an anterior corpus that narrows to an isthmus and then ends in a muscular bulb. The rhabditiform esophagus is sharply delineated and well defined with an obvious sharp demarcation between the end of the esophagus and the beginning of the intestine. The overall length of the rhabditiform esophagus is less than 25% of the total length of the larvae (Fig. 1.24). In contrast, the esophagus of the lungworm larvae tends to be less well defined and longer, making up about 33%–50% of the total length of the larvae (Figs. 1.23B and 1.25). The relatively short buccal tube differentiates the larvae of Strongyloides (Fig. 1.23A) from those of hookworms–strongyles–trichostrongyles, which have a long buccal tube (Fig. 1.26).


Fig. 1.23 (A) Anterior end of a first‐stage larva of Strongyloides stercoralis recovered from the feces of a dog. This larva has been killed and stained with dilute iodine. The rhabditiform esophagus is well defined and obvious. Note the short buccal tube (arrow) and the rhabditiform esophagus made up of the corpus (C), isthmus (I), and muscular bulb (MB). Also note the distinct border demarcating the end of the esophagus and the start of the intestine (GI). (B) Anterior end of an iodine‐stained first‐stage larva of Crenosoma vulpis recovered from the feces of a dog. Note the poorly defined esophagus (E). It is difficult to discern the demarcation between the end of the esophagus and the start of the intestine (GI).

Differentiation of the various lungworm larvae is based on tail morphology. In dogs, larvae with a straight tail and lacking a rhabditiform esophagus are Crenosoma vulpis (see Figs. 1.25 and 1.92). Larvae that have a kinked S‐shaped tail but lack a dorsal spine are either Oslerus osleri or Filaroides hirthi (see Figs. 1.94 and 1.95). Larvae with a kinked tail and a dorsal spine are Angiostrongylus vasorum (see Fig. 1.93). In cats, larvae with a kinked tail and a dorsal spine are Aelurostrongylus abstrusus (see Figs. 1.90 and 1.91).


Fig. 1.24 Strongyloides stercoralis first‐stage larvae recovered from the feces of a dog (Lugol’s iodine stained and killed). Note the rhabditiform esophagus (R), prominent genital primordium (GP), and the straight tail (T). The rhabditiform esophagus makes up about 25% of the total length of the larvae.


Fig. 1.25 Crenosoma vulpis first‐stage larva recovered from the feces of a dog. Note the indistinct poorly defined esophagus (E). The esophagus makes up about 33%–50% of the total length of the larvae in metastrongyloid lungworms.

There should be only a single species of nematode lungworm larva, Dictyocaulus viviparus (see Fig. 1.151), recovered in properly collected fresh feces of cattle. The larvae have an abundance of visible food granules and a straight tail. The same situation occurs with the horse, although infection with Dictyocaulus arnfieldi (see Fig. 1.176, 1.177) is patent in donkeys, but only rarely in horses. In small ruminants, there are two larvae with straight tails, one with an abundance of visible food granules (Dictyocaulus filaria) (see Fig. 1.152) and the other without (Protostrongylus rufescens) (see Figs. 1.149 and 1.150). Another lungworm, Muellerius capillaris, produces larvae with a kinked tail and dorsal spine (see Figs. 1.146 and 1.147). Cystocaulus and Neostrongylus are small ruminant lungworms that are found in parts of Europe and Asia. The tails of their larvae have additional spines that can be used to differentiate them from Muellerius larvae.


Fig. 1.26 Anterior end of a first‐stage larva of a hookworm, Uncinaria stenocephala, recovered from an improperly collected fecal sample of a dog. This larva has been killed and stained with dilute iodine. As with Strongyloides, note the corpus (C), isthmus (I), and muscular bulb (B) of the rhabditiform esophagus and the intestine (GI). In contrast to Strongyloides, note the long buccal tube (BT).


Grazing animals are infected with a variety of species of strongylid nematodes, which produce eggs that are not easily differentiated. In veterinary practices, it is usually unnecessary to identify individual species because treatment and control are generally directed to the entire group of nematodes rather than to a single species. If identification of the strongylid genera present in an animal or group of animals is needed, the simplest method for identification is culture of eggs to the third larval stage. In ruminants, these larvae can then be identified to parasite genus. In horses, this technique can be used to differentiate large and small strongyle larvae and identify some genera specifically. Currently, researchers are developing protocols for identification of parasite genera using molecular techniques (polymerase chain reaction [PCR]), and these procedures are now becoming commercially available.

Fecal Culture

  1. Fresh feces from cattle or horses should be thoroughly mixed and moistened with water if dry. Feces should not be wet, only moist. Larvae do not survive well in very wet fecal material. If feces are very soft or liquid, peat moss or vermiculite can be added to create a more suitable consistency. Sheep and goat pellets can be cultured as they are, without breaking them up. Rectal fecal samples are preferred for culture to prevent contamination with free‐living nematodes.
  2. Place feces in a cup or jar in a layer several centimeters deep. The container should have a loose cover that does not prevent air circulation but will deter flies and reduce desiccation. The culture can be kept at room temperature for 10–20 days or at 27°C for 7 days. Daily stirring of the culture will inhibit mold growth and circulate oxygen for the developing larvae. Additional water can be added if feces begin to dry out.
  3. Following the culture period, harvest larvae with the Baermann test described previously. Alternative containers and methods for harvest of larvae can be found in Bowman (2014), Taylor et al. (2015), and other textbooks of veterinary parasitology.

Identification of Ruminant and Camelid Third‐Stage Larvae

To identify larvae, place a drop or two of liquid containing larvae from the Baermann procedure on a microscope slide. Add an equal amount of Lugol’s iodine. The iodine will kill and stain the larvae so that they can be examined closely.

Larvae recovered from ruminant fecal material can be most easily identified by a combination of morphology and size. The shape of the head and the shape of the tail and of the sheath extending beyond the tail at the posterior end of the larva are important characteristics, and both should be evaluated on each larva before an identification is made. The sheath is the retained cuticle of the second larval stage and provides larvae with increased protection from environmental conditions. Measurements of total larval and sheath length are helpful as well (Table 1.3), but sizes often overlap between genera and size characteristics can be affected by culture conditions and age of larvae. Consequently, measurements alone should not be used to identify larvae.

Table 1.3. Morphologic characteristics of infective third‐stage strongylid larvae of domestic ruminants

Sources: Bowman (2014) and Ministry of Agriculture, Fisheries and Food (1986).

Genus Overall length (μm) Anus to tip of sheath (μm) End of tail to tip of sheath (μm) Other characteristics

Head rounded; tail of sheath short; tail may have one or two tuberosities

Head squared; tail of sheath shorter in sheep

Head rounded; sheath tail medium length, offset

Head squared with two refractile oval bodies at anterior end of the esophagus; medium‐length sheath tail tapering to fine point

Broad, rounded head; intestine with eight cells; tail notched and lobed; long thin sheath tail

Small larva with rounded head; long thin sheath tail

Rounded head; long thin sheath tail; 16–24 triangular intestinal cells

Rounded head; long thin sheath tail; 24–32 rectangular intestinal cells
Sheep 710–789 175–220 110–150

Relative proportions of parasite genera in larval cultures cannot be used to predict numbers of adult worms in the gastrointestinal tract. For example, Haemonchus contortus is highly prolific and may dominate in small ruminant fecal cultures, even when adult parasites of other genera are present in substantial numbers. Figures 1.271.37 show morphologic characteristics of common third‐stage larvae of small ruminants and cattle.


Fig. 1.27 Third‐stage larvae of common small ruminant strongylid genera collected from fecal culture. This photo shows the relative size relationships among the larvae. The following photographs show the details and the anterior and posterior ends of the individual larvae. Nematodirus spp. larvae are usually not encountered in cultures and are not illustrated here. They are bigger in total length and have a longer tail sheath than other larvae. (A) Trichostrongylus, (B) Teladorsagia, (C) Oesophagostomum/Chabertia, (D) Haemonchus, and (E) Cooperia.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.28 Trichostrongylus larva from sheep, head (A) and tail sheath (B). The head of Trichostrongylus larvae is tapered and the tail sheath is short. The tail may end in one or two tuberosities.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.29 Ovine Teladorsagia head (A) and tail sheath (B). Teladorsagia can easily be confused with Trichostrongylus, but Teladorsagia is generally larger and the head is squared, not tapered. The sheath of the tail of Teladorsagia is short.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department ofAnimal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.30 Oesophagostomum/Chabertia head (A) and tail sheath (B). The larvae of these two genera are not easily distinguishable, but they are not difficult to differentiate from other genera. The tail sheath is long and filamentous, and the head is broad and rounded.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.31 Haemonchus head (A) and tail sheath (B). The larvae of Haemonchus have the most narrowly rounded head of the common larvae. The tail sheath is medium in length and often has a slight kink at the end of the tail.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.32 Cooperia from a sheep, head (A) and tail sheath (B). Cooperia third‐stage larvae are distinguished by a pair of refractile bodies (arrow) present in a squared head. These bodies are difficult to photograph but easy to appreciate under the microscope. The sheath of the tail is medium in length and tapering or finely pointed.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.33 Strongyloides papillosus is a nematode of ruminants that is unrelated to the important strongylid nematodes. Infective third‐stage larvae of Strongyloides may be present in larval cultures. They do not have a sheath and the esophagus is very long (E). Additionally, free‐living nematodes may be numerous in cultures contaminated with soil. For information on identifying free‐living nematodes, see the section on identifying larval nematodes in fecal samples.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.34 Third‐stage Cooperia larvae from cattle. Several species of Cooperia infect ruminants, and the length of the tail sheath is variable. Cooperia oncophora (A) produces larvae with a longer tail sheath than other species of the genus (B).

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.35 Tail sheath of Ostertagia (A) and Trichostrongylus (B). Both genera have a short tail sheath, but Ostertagia has a blunter head.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.

Additional information on identification of ruminant third‐stage larvae can be found at the website of the RVC/FAO Guide to Veterinary Diagnostic Parasitology: www.rvc.ac.uk/review/Parasitology/Index/Index.htm and in van Wyk and Mayhew (2013).


Fig. 1.36 Tail sheath of Haemonchus (A) and Oesophagostomum (B) from cattle. The tail sheaths of the parasites occurring in cattle are similar to those in sheep.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.


Fig. 1.37 Tail sheath of Bunostomum. Species of this ruminant hookworm infect both cattle and sheep. The third‐stage larva is smaller than those of other genera and has a thin tail sheath.

Photo courtesy of Dr. Tom Yazwinski and Mr. Chris Tucker, Department of Animal Science, University of Arkansas, Fayetteville, AR.

Identification of Third‐Stage Larvae of Equine Strongyles

Horses are infected with over 30 species of strongylid parasites, but only a few can be identified on the basis of the third‐stage larva. Most of the small strongyle species can only be identified as cyathostomin parasites from the infective larval stage (Fig. 1.38 and Table 1.4). The posterior portion of the sheath of horse strongyle larvae is very long and filamentous, making these larvae easily recognizable as infective parasite larvae. The number of intestinal cells in these larvae is variable and is useful in identification.


Fig. 1.38 Infective third‐stage larvae of both large and small equine strongyles have a very long filamentous extension of the sheath. (A) The larvae of small strongyles (cyathostomes) have eight intestinal cells, which can be easily counted in the larva shown here. (B) Larvae of large strongyle species have more than eight intestinal cells, like this Strongylus vulgaris larva with at least 28 cells.

Table 1.4. Morphologic characteristics of infective third‐stage strongylid larvae of horses

Source: Adapted from Ministry of Agriculture, Fisheries and Food (1986).

Genus Characteristics
Strongyloides Sheath absent; esophagus almost half the length of the body
Trichostrongylus axei Tail of sheath short, not filamentous
Most small strongyles (Cyathostominae) Long filamentous sheath; eight triangular intestinal cells
Gyalocephalus (small strongyle) Long filamentous sheath; 12 rectangular intestinal cells
Oesophagodontus (small strongyle) Large larva; long filamentous sheath; 16 triangular intestinal cells
Posteriostomum (small strongyle) Long filamentous sheath; 16 roughly rectangular intestinal cells
Strongylus equinus (large strongyle) Long, thin larva with filamentous sheath; 16 poorly defined rectangular intestinal cells
Triodontophorus (large strongyle) Medium‐length and broad larva with filamentous sheath; 18–20 well‐defined rectangular intestinal cells
Strongylus edentatus (large strongyle) Smaller larvae with filamentous sheath; 18–20 poorly defined and elongated intestinal cells
Strongylus vulgaris (large strongyle) Large larvae with filamentous sheath; short esophagus; 28–32 well‐defined, rectangular intestinal cells

Fecal Egg Count Reduction Test (FECRT)

One of the principal uses of quantitative egg counts is the evaluation of drug efficacy. Anthelmintic resistance in strongylid nematodes of horses, small ruminants, and cattle is rapidly increasing worldwide. The only technique for evaluating drug efficacy that can be conducted by veterinary practitioners in all host species is the FECRT. In this procedure, the percentage reduction in strongylid fecal egg counts following treatment is calculated to evaluate the efficacy of the product. This test has also been used to evaluate resistance to anthelmintics in equine Parascaris spp. infections.

The World Association for the Advancement of Veterinary Parasitology (WAAVP) is an international organization that assembles expert opinion and has issued recommendations for testing and evaluating antiparasiticide efficacy. Updated recommendations for evaluating drug resistance are expected in the near future. Some general principles for conducting FECRT are presented here. For additional details in conducting these tests several recent publications can be consulted: Kaplan R. M. 2020. Biology, epidemiology, diagnosis, and management of anthelmintic resistance in gastrointestinal nematodes of livestock. Vet. Clin. North Am. Food Anim. Pract. 36:17–30, and American Association of Equine Practitioners. Internal Parasite Control Guidelines, updated 2019; https://aaep.org/guidelines/parasite‐control‐guidelines.

Although there is some variation in protocols for conducting FECRT, the following general principles are presented followed by comments specific to host species.

Test Groups and Selection of Animals

An accurate FECRT requires adequate animal numbers. In cases where owners wish to test very small flocks or herds, it is important to be cautious in interpretation of results. For ruminants 15 animals are preferred, but at least 10 should be used for each drug to be tested. For horses, where herd sizes are often low, at least six horses per group is recommended. Where only one or two animals are available an impression of drug efficacy can be obtained following treatment, but it should not be considered a reliable or accurate FECRT. In very large herds, 10% of the total population is adequate.

All animals used in the FECRT should have adequate egg counts to allow reductions to be determined accurately. For sheep, lambs of 3–6 months of age and cattle less than 16 months are the best candidates for a FECRT. Goats of all ages can generally be used. It is best if animals used in the test are similar in age and management. Fecal egg counts in adult cattle usually are too low to be used in an FECRT. It will be helpful to perform the FECRT at the time of year when fecal egg counts are expected to be the highest, based on epidemiology of the parasites in a region.

As a general guideline, the modified McMaster test can be used when the average FEC is 500 EPG with a group size of 10 or 250 EPG with a group size of 20. If average egg counts fall below this level, an alternative test for quantifying parasite eggs must be used to provide an accurate picture of drug efficacy. Alternatives include decreasing the detection limit of the modified McMaster test by using larger sample aliquots (e.g., using larger counting chambers available from several companies,) or using other tests with a lower detection limit (mini‐FLOTAC, Stoll, or Wisconsin tests; see section on “Egg‐Counting Procedures [Quantitative Fecal Exams]”).

An alternative procedure for conducting a FECRT using composite fecal samples is described in Kaplan (2020).

Test Drugs and Collection of Posttreatment Samples

Once the test group or groups of animals have been established with a pretreatment fecal egg count, each animal should be individually weighed and treated with the test drug at the manufacturer’s recommended dose.

On farms where individual animals cannot be weighed, the fecal egg count reduction (FECR) can be approximated by treating all animals with the drug dose for the estimated heaviest animal in the group. The results, however, will not be entirely accurate because some animals will be receiving more than the recommended dose, which may be temporarily effective against worms resistant to the recommended dose. This will lead to an overestimation of drug efficacy.

The optimum time following treatment for collection of posttreatment samples varies with the test drug because of variable effects on larval stages and temporary sterilizing effects on adult parasites. For ruminants, the following intervals are recommended:

  • benzimidazoles, levamisole, or pyrantel 10–14 days,
  • ivermectin and other avermectins 14–17 days,
  • moxidectin 17–21 days.

For convenience in both ruminants and horses, a standard period of 14 days before collection of the posttreatment samples is often recommended.

Interpretation of Results

The % FECR is calculated using arithmetic group means in the following formula:


When small groups of animals are used, individuals with particularly high fecal egg counts can dramatically skew the FECR when group means are used in the calculation. In these cases (e.g., in small groups of horses), it is best to calculate individual FECR using the above formula and then average the individual FECR to obtain a group average.

The FECRT can also be conducted by comparing mean posttreatment EPG in a group of untreated animals with that of a group of treated animals. This procedure is used less often than the comparison of pre‐ and posttreatment samples of individuals in the same group of animals because more animals are needed and pretreatment egg counts must be similar in the two groups. The formula for calculating FECR is altered accordingly when treatment and control groups are used:


If no resistance is present, and the test is administered correctly, anthelmintics in ruminants can reliably decrease fecal egg counts by >95%. For horses, reductions in fecal egg counts when no resistance is present should be >95% for benzimidazoles, >98% for ivermectin and moxidectin, and >90% for pyrantel.

The FECR obtained when testing a group of animals gives an indication of drug efficacy, but is not equivalent to the proportion of the adult worm population that is removed by treatment. For example, if the percentage reduction is 10%, a large proportion of worms in the animals are probably resistant. If the reduction is 80%, the proportion of resistant worms is much smaller, but still significant. To further evaluate which worm genera are resistant in ruminants, the nematode eggs present in both pre‐ and posttreatment fecal samples can be identified by larval culture (see earlier) or PCR. In horses, drug resistance is most often present in small strongyles (cyathostomins), which cannot be readily differentiated to species based on morphology.


Tapeworm segments and adult gastrointestinal nematodes may occasionally be passed in feces and presented for identification by concerned owners. Segments of common tapeworms can usually be readily identified to the level of genus by shape and identification of eggs in the segments (see Figs. 1.99, 1.102, 1.105, and 1.154 for photographs of common tapeworm segments), but nematode parasites may be more difficult to identify. When preserving nematodes for further identification, it is helpful to place them first in tap water and refrigerate the container for several hours. This will relax the worms and make them easier to examine. After relaxation, the worms can be placed in 70% ethanol or 10% buffered formalin. Ethanol, but not formalin, allows later identification with molecular assays. While not optimum preservatives for all helminths, these chemicals are readily available to most veterinarians.

The most common nematodes presented by pet owners are the ascarids (roundworms; see Fig. 1.72). These large, stout‐bodied worms are common in feces and vomitus of kittens and puppies. Horse owners might also see equine ascarids that are up to 50 cm (about 20 in.) in length. Smaller nematodes present in equine manure may be the large and small strongyles or pinworms (Oxyuris). Larval or adult horse strongyles may be red or cream in color and no more than about 2–4 cm in length (see Fig. 1.168). Oxyuris, the equine pinworm, can reach 15 cm, and the females have distinctive long, thin tails (see Fig. 1.174). Nematodes are most likely to be seen in diarrheic feces or following treatment.

Specific identification of adult nematodes is usually based on morphologic variations of the outer layer, or cuticle, of the worms. Microscopic examination of the mouthparts and accessory sexual structures may be required. To enhance visualization of these structures, the worm can be mounted in a clearing solution, which dissolves the soft tissue, leaving only the cuticle. If the worm is large, the areas of diagnostic importance (usually the anterior and posterior ends) can be cut off and mounted in a few drops of the clearing solution. Procedures for making Hoyer’s and lactophenol solutions are given below, but they are not usually prepared in veterinary practices because they require either controlled or hazardous substances. Both are commercially available.

Depending on the parasite species, accurate worm identification may require the evaluation of subtle morphologic characteristics that will be unfamiliar to most practicing veterinarians. When specific parasite identification is needed, worms should be submitted to a parasitologist for examination.

Hoyer’s Solution

Hoyer’s solution also provides a permanent mounting medium for specimens, although the clearing process will continue until eventually internal structures will no longer be visible:

  • 30 g gum arabic;
  • 16 mL glycerol;
  • 200 g chloral hydrate;
  • 50 mL distilled water.

Dissolve the gum arabic in water with gentle heat. Add the chloral hydrate, then the glycerol.


  • 20 mL glycerin, pure;
  • 10 mL lactic acid;
  • 10 mL phenol crystals, melted;
  • 10 mL distilled water.

Combine all ingredients.


The following photographs in this chapter illustrate the diagnostic stages found in feces of a wide variety of both common and some uncommon parasites of major domestic species. Because an appreciation of relative sizes of parasite eggs, cysts, and oocysts, is very helpful in identification, a line drawing precedes sections showing parasites of common mammalian hosts (Figs. 1.39, 1.40, 1.121, 1.122, 1.162, 1.180). Generally, photographs of eggs and cysts were taken using the high‐dry (40×) objective, although some photographs using the 10× objective are included to show relative sizes of eggs and cysts.

The figures in which each parasite appears are listed after the name. They may include figures in other sections where more than one parasite is illustrated.

An effort has been made to minimize taxonomic information while still permitting an appreciation of the larger groups to which each individual species belongs. For more specific taxonomic information, a textbook of veterinary parasitology should be consulted.

Also at the beginning of the sections for dogs and cats, ruminants, horses, and swine are tables showing common U.S. label approved products for treatment of a number of parasitic infections. Label dose and withdrawal information should always be consulted before treatment of animals with parasiticides.


Fig. 1.39 Parasites found in canine feces. Figure courtesy of Dr. Bert Stromberg and Mr. Gary Averbeck, College of Veterinary Medicine, University of Minnesota, Minneapolis, MN.

*Differentiate from other larvae that could be present in canine feces, including Filaroides, Oslerus and Angiostrongylus.

**Spirocerca lupi, which is common in many parts of the world, has a larvated egg similar in appearance to Physlaoptera, but more elongated.

***Oocysts of Hammondia heydorni are similar in appearance to Neospora.


Fig. 1.40 Figure courtesy of Dr. Bert Stromberg and Mr. Gary Averbeck, College of Veterinary Medicine, University of Minnesota, Minneapolis, MN.

*Oocysts of Hammondia hammondi and Besnoitia spp. are similar in appearance to Toxoplasma oocysts.

Table 1.5. Representative treatments for selected parasites of dogs

Parasite Effective treatments Dose and administration route
Cystoisospora spp. aPonazuril, atoltrazuril 10–30 mg/kg orally q 24 h × 1–3 d
bSulfadimethoxine Administer according to label directions
Giardia sp. aFebantel 30 mg/kg q 24 h × c3 d (combined with praziquantel and pyrantel); avoid in pregnant animals
aFenbendazole 50 mg/kg orally q 24 h × c3 d
Ancylostoma spp.
Toxascaris leonina Toxocara canis
Uncinaria stenocephala
Febantel, fenbendazole, dmilbemycin oxime, transdermal moxidectin, pyrantel pamoate Administer according to label directions
Trichuris vulpis Febantel, fenbendazole, milbemycin oxime, transdermal moxidectin Administer according to label directions
Eucoleus spp. aMilbemycin oxime, atransdermal moxidectin Extralabel use of label dose of transdermal moxidectin or elevated dose (2 mg/kg) of milbemycin oxime
Physaloptera spp. aPyrantel pamoate 20 mg/kg, repeat q 14 d until clinical signs resolve
Spirocerca lupi a , eDoramectin 0.4 mg/kg subcutaneously q 7 d × 12 weeks
Strongyloides stercoralis aFenbendazole
a , eIvermectin
50 mg/kg fenbendazole orally q 24 h × 5 d, repeat in 4 weeks; 0.2 mg/kg ivermectin subcutaneously, repeat in 2 weeks
Dipylidium caninum
Taenia spp.
Echinococcus spp.
fEpsiprantel, gfenbendazole, praziquantel Administer according to label directions
Mesocestoides spp.
Alaria spp.
aPraziquantel Effective against intestinal stages when administered according to label directions
Diphyllobothrium latum
Spirometra spp.
aPraziquantel Administer elevated dose (25 mg/kg) for 2 consecutive days
Heterobilharzia americana
Paragonimus kellicotti
50 mg/kg orally for 10–14 days
25 mg/kg every 8 hours for 2–3 days
Nanophyetus salmincola aPraziquantel 20–30 mg/kg once

a Extralabel use supported by published data.

b Label‐approved for treating dogs with bacterial enteritis associated with coccidiosis.

c Longer courses of treatment may be necessary in some patients.

d Monthly products are label‐approved against Ancylostoma spp., Toxocara canis, and Toxascaris leonina, but not Uncinaria stenocephala.

e Extralabel use of high‐dose cattle products can be fatal in dogs; establish MDR1 status prior to treatment.

f Not label‐approved against Echinococcus spp.

g Only effective against Taenia spp., not D. caninum or Echinococcus spp.

Additional information on parasite treatments can be found in Chapter 7.

Table 1.6. Representative treatments for selected parasites of cats

Parasite Effective treatments Dose, route, and regimen
Cystoisospora spp. aPonazuril, atoltrazuril 10–30 mg/kg orally q 24 h × 1–3 d
aSulfadimethoxine Administer according to label directions
Tritrichomonas blagburni aRonidazole 30–50 mg/kg orally q 12 h × 14 d; use with caution due to safety concerns
Giardia sp. aFebantel 30 mg/kg × b3 d (combined with praziquantel and pyrantel); avoid in pregnant animals
aFenbendazole 50 mg/kg orally q 24 h × b3 d
Ancylostoma spp.
Toxocara cati
Emodepside, eprinomectin, civermectin, milbemycin oxime, pyrantel pamoate, transdermal moxidectin, selamectin Administer according to label directions
Strongyloides spp. aFenbendazole
a , dIvermectin
50 mg/kg fenbendazole orally q 24 h × 5 d, repeat in 4 weeks; 0.2 mg/kg ivermectin subcutaneously, repeat in 2 weeks
Aelurostrongylus abstrusus aTransdermal moxidectin Administer according to label directions
Dipylidium caninum
Taenia spp.
Echinococcus spp.
eEpsiprantel, praziquantel Administer according to label directions
Mesocestoides spp.
Alaria spp.
aPraziquantel Effective against intestinal stages when administered according to label directions
Diphyllobothrium latum
Spirometra spp.
aPraziquantel Administer elevated dose (25 mg/kg) for 2 consecutive days
Paragonimus kellicotti aFenbendazole
50 mg/kg orally for 10–14 days
25 mg/kg every 8 hours for 2–3 days
Platynosomum concinnum aPraziquantel 25 mg/kg every 8 hours for 2–3 days

a Extralabel use supported by published data.

b Longer courses of treatment may be necessary in some patients.

c Monthly product is only label‐approved against Ancylostoma spp., not Toxocara cati.

d Extralabel use of high‐dose cattle products can be fatal in cats; use with particular caution in young or debilitated patients.

e Not label‐approved against Echinococcus spp.

Additional information on parasite treatments can be found in Chapter 7.

Protozoan Parasites

Parasite: Cystoisospora (Isospora) spp. (Figs. 1.4–1.6, 1.411.45, 1.47, 1.70, 1.78, 1.101)

Common name:Coccidia.

Taxonomy: Protozoa (coccidia). Several host‐specific species are found in the dog (C. canis, C. ohioensis, C. neorivolta, C. burrowsi) and cat (C. felis, C. rivolta).

Geographic Distribution: Worldwide.

Location in Host: Small intestine, cecum, and colon.

Life Cycle: Cats and dogs are infected by ingestion of sporulated oocysts or infected transport hosts (often rodents, but also including rabbits, ruminants, birds, and other prey animals). Following development in the final host, oocysts are passed in feces and undergo sporulation in the environment.

Laboratory Diagnosis: Oocysts are detected by fecal flotation examination. Oocysts have smooth, clear cyst walls, are elliptical in shape, and contain a single, round cell (sporoblast) when freshly passed. The oocysts of C. ohioensis, C. burrowsi, and C. neorivolta are not morphologically distinguishable and are referred to as the C. ohioensis complex.

Size: C. canis, C. felis 38–51 × 27–39 μm

Other Cystoisospora spp. 17–27 × 15–24 μm

Clinical Importance: These are the organisms typically referred to as “coccidia” of dogs and cats, although other parasites also fall into this taxonomic group. Oocysts can be found in the feces of many clinically normal young dogs and cats. Clinical coccidiosis most often occurs in puppies and kittens, often in association with weaning, change of owner, or other stress factors. Signs include diarrhea, abdominal pain, anorexia, and weight loss. In severe cases, bloody diarrhea and anemia may occur. Respiratory and neurologic signs have also been reported in some animals. Clinical disease has been difficult to reproduce in experimental infections.


Fig. 1.41 Dog and cat coccidia species produce oocysts of different sizes. This figure shows C. canis (larger oocysts) and an oocyst of the C. ohioensis complex (smaller oocyst).

Photo courtesy of Dr. David Lindsay, Virginia‐Maryland College of Veterinary Medicine, Virginia Tech, Blacksburg, VA.


Fig. 1.42 Cystoisospora oocysts usually require a minimum of 1–2 days to become infective for the next host (sporulated). In warm conditions, oocysts undergo the first cell division soon after being passed in the feces. In the two‐cell stage, they may be mistaken for sporulated oocysts. In this fecal sample, a sporulated oocyst (arrow) is adjacent to one in the two‐cell stage.


Fig. 1.43 A sporulated Cystoisospora oocyst contains two sporocysts, each containing four sporozoites. The two sporocysts can be seen in this oocyst, although only two of the four sporozoites can be visualized in each sporocyst. A large, round residual body is also present in each sporocyst.


Fig. 1.44 Cystoisospora oocyst and two iodine‐stained Giardia cysts (arrows) in a canine fecal sample.

Photo courtesy of Dr. Robert Ridley, College of Veterinary Medicine, Kansas State University, Manhattan, KS.

Parasite: Toxoplasma gondii, Neospora caninum (Fig. 1.47)

Taxonomy: Protozoa (coccidia).

Geographic Distribution: Worldwide.

Location in Host: Intestine and other tissues of cats and other felids (Toxoplasma) and dogs and other canids (Neospora).

Life Cycle: Toxoplasma is transmitted to cats by ingestion of cysts containing bradyzoites in tissues of intermediate hosts. Prenatal and transmammary transmission as well as direct transmission through ingestion of sporulated oocysts can also occur. Transmission of Neospora in dogs appears to be similar to Toxoplasma transmission.

Laboratory Diagnosis: Oocysts are detected in feces by centrifugal or simple flotation techniques. However, very few oocysts of Neospora appear to be produced in infected dogs. Immunodiagnostic tests are available to identify current and past exposure to Toxoplasma in cats but are usually not positive until after fecal passage of oocysts has ceased. Dogs can also be tested for antibody to Neospora. The small, spherical‐shaped oocysts of the two genera are morphologically identical, have a clear smooth cyst wall, and contain a single round sporoblast. Some other coccidia genera, including Hammondia, produce similar oocysts, which precludes definitive identification of Toxoplasma or Neospora on the basis of oocyst presence alone.

Size:11–14 × 9–11 μm

Clinical Importance: Toxoplasma infections in cats are generally well tolerated. Clinical disease (ocular, respiratory, etc.) can occur in cats, especially young or immunosuppressed animals. Toxoplasmosis is an important zoonotic disease with especially serious consequences in pregnant women and the immunosuppressed. Congenital Neospora infection can result in severe central nervous system disease in dogs. Neospora infection is also an important cause of abortion in the bovine intermediate host.


Fig. 1.45 Cystoisospora canis oocysts in this canine fecal sample (arrows) are similar to the larger Toxascaris leonina egg (also Fig. 1.65), but the oocysts are smaller and lack the membranous appearance of the inside of the shell seen in Toxascaris eggs.


Fig. 1.46 Eimeria spp. oocysts are sometimes seen in dog and cat feces. Eimeria does not infect these hosts, but oocysts consumed as a result of predation or coprophagy will pass unharmed through the gastrointestinal tract and may be misidentified as Cystoisospora. Many (but not all) Eimeria oocysts have a knob at one end called the micropyle cap (arrow), whereas Cystoisospora spp. lack a cap. If this cap is present, an oocyst in dog or cat feces can be identified as a “spurious parasite.”


Fig. 1.47 Neospora and Toxoplasma oocysts are similar to common Cystoisospora spp., but they are smaller. The oocysts of these two coccidia genera are similar in appearance and also cannot be distinguished from oocysts of Hammondia, another coccidia genus of small animals. This photo of a feline fecal sample also shows an oocyst of C. rivolta (arrow).

Parasite: Sarcocystis spp. (Figs. 1.48 and 1.49)

Taxonomy: Protozoa (coccidia). A number of species infect dogs or cats, each with a specific intermediate host.

Geographic Distribution: Worldwide.

Location in Host: Small intestine of dogs and cats.

Life Cycle: Cat and dog definitive hosts are infected by ingesting intermediate host tissue containing sarcocysts. Sexual reproduction in dogs or cats leads to formation of oocysts that sporulate while still in the intestinal tract.

Laboratory Diagnosis: Oocysts form within the gastrointestinal tract of dogs and cats. The oocyst wall breaks down in the gut, and small, ellipsoidal sporulated sporocysts are released in the feces. They are detected by centrifugal or simple flotation techniques.

Size:7–22 × 3–15 μm

Clinical Importance: Sarcocystis is generally nonpathogenic in the definitive host, although some species can cause severe disease in the intermediate host (cattle, sheep, pigs, horses).

Parasite: Cryptosporidium spp. (Fig. 1.50)

Taxonomy: Protozoa (coccidia). Cryptosporidium felis and C. canis appear to be the primary species infecting cats and dogs, respectively.

Geographic Distribution: Worldwide.

Location in Host: Small intestine.

Life Cycle: These parasites have a direct life cycle. Cats and dogs are infected following ingestion of oocysts, which are infective as soon as they are passed in the feces. Following asexual and sexual multiplication of the organism in the intestine, oocysts are produced and exit the host in the feces.

Laboratory Diagnosis: Small oocysts in the feces are detected by use of acid‐fast or other stains of fecal smears, Sheather’s sugar flotation test, fecal antigen tests, or molecular diagnostic procedures. Oocysts of C. parvum and C. canis are morphologically indistinguishable, while C. felis oocysts are smaller than those of the other two species.

Size: C. felis 3.5–5 μm in diameter

C. parvum, C. canis 7 × 5 μm

Clinical Importance: Cryptosporidiosis has been reported as an uncommon cause of chronic diarrhea in cats. Affected cats are often immunosuppressed by other causes. Although implicated in rare instances, Cryptosporidium infections in dogs and cats do not appear to be a significant source of zoonotic exposure for humans.


Fig. 1.48 Sarcocystis sporocysts are smaller than typical coccidia oocysts and have a smooth, clear cyst wall. Each sporocyst contains four banana‐shaped sporozoites.

Photo courtesy of Dr. Robert Ridley, College of Veterinary Medicine, Kansas State University, Manhattan, KS.


Fig. 1.49 Sarcocystis

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