EGG‐COUNTING PROCEDURES (QUANTITATIVE FECAL EXAMS)

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.

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.

SIMPLE SEDIMENTATION TEST

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.

CENTRIFUGAL SEDIMENTATION TEST

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.

QUALITY CONTROL FOR FECAL EXAM PROCEDURES

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.

USE OF THE MICROSCOPE

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.

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).

PSEUDOPARASITES AND SPURIOUS PARASITES

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. 1.10–1.15). “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. 1.16‐1.18). 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.

IDENTIFICATION OF NEMATODE LARVAE RECOVERED WITH FECAL FLOTATION OR BAERMANN PROCEDURES

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.

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.

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).

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).

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.

TECHNIQUES FOR EVALUATION OF STRONGYLID NEMATODES IN GRAZING ANIMALS

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.

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.

Genus	Overall length (μm)	Anus to tip of sheath (μm)	End of tail to tip of sheath (μm)	Other characteristics
Trichostrongylus				Head rounded; tail of sheath short; tail may have one or two tuberosities
Sheep Cattle	622–796 619–762	76–118 83–107	21–40 25–39
Ostertagia				Head squared; tail of sheath shorter in sheep
Sheep Cattle	797–910 784–928	92–130 126–170	30–60 55–75
Haemonchus				Head rounded; sheath tail medium length, offset
Sheep Cattle	650–751 749–866	119–146 158–193	65–78 87–119
Cooperia				Head squared with two refractile oval bodies at anterior end of the esophagus; medium‐length sheath tail tapering to fine point
Sheep Cattle	711–924 666–976	97–150 109–190	35–82 47–111
Nematodirus				Broad, rounded head; intestine with eight cells; tail notched and lobed; long thin sheath tail
Sheep Cattle	922–1118 1095–1142	310–350 296–347	250–290 207–266
Bunostomum				Small larva with rounded head; long thin sheath tail
Sheep Cattle	514–678 500–583	153–183 129–158	85–115 59–83
Oesophagostomum				Rounded head; long thin sheath tail; 16–24 triangular intestinal cells
Sheep Cattle	771–923 726–857	193–235 209–257	125–160 134–182
Chabertia				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 1.27–1.37 show morphologic characteristics of common third‐stage larvae of small ruminants and cattle.

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).

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.

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.

IDENTIFICATION OF ADULT WORMS

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.

Lactophenol

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

Combine all ingredients.