Fecal Samples

Collection and Handling of Fecal Samples

Preferably, fecal samples should be collected from the rectum. If material is collected from the ground it should be from the top of a freshly passed deposit. Avoid deposit areas in contact with the ground. Care must be taken with samples collected from the ground to avoid doing fecal exams on neighbourhood or stray animals. It is advisable to collect only those samples which can be positively identified as relevant to the animal in question

  1. A minimum sample size is 5 g. Preferably, submit a “golf-ball” size sample in a plastic bag or a container that is airtight, watertight, and suitably robust. Gloves are not suitable primary containers.
  2. All containers must be clearly and completely labelled: name, species of animal, date of collection.
  3. Store the specimens in a refrigerator until shipping. DO NOT FREEZE FECAL SAMPLES. Samples should be at the laboratory within 12 hours if possible.
  4. In cases involving a herd problem, ideally, every animal in the herd should be sampled. However for a reliable evaluation of a herd, sample 10-25% of individuals. When samples of individuals cannot be identified, such as in a feed lot situation, take random samples and clearly label them as such.
  6. Most parasites (excepting some protozoans) will still be detectable and easily identifiable in fecal samples examined 2 to 3 days after collection, if the samples have been refrigerated in the meantime. If more than 2 to 3 days may elapse between collection and examination (or the samples cannot be refrigerated), mix equal parts of 5% formalin and feces. This will prevent parasite development, especially the hatching of eggs. This procedure should not be used if the diagnostic technique depends on living parasites, such as the Baermann technique.

Feces, Direct Smears

These can be made by transferring a small sample of feces to a glass slide. Mix in a small amount of saline. Drop on a cover slip and examine directly under the microscope. This technique will reveal heavy infections of eggs and cysts (such as coccidiosis in poultry). It may also detect helminth eggs and larvae or protozoan trophozoites which typically do not float and therefore are not detected with standard fecal flotation techniques.

Feces, Visual Examination

Search macroscopically for large gravid segments of cestodes (e.g. Dipylidium caninum in dog and cat feces) or whole adult helminths (e.g. Parascaris equorum in horse feces).

Feces, Protozoal Diarrhea

Some developmental stages of these organisms (e.g. Giardia species, Trophozoites) are too fragile to withstand transport by courier. Testing ideally should be done at the practitioner’s own laboratory, by examining saline wet-mounts of fresh warm feces. Delivery shortly after passage to the diagnostic laboratory is an alternative.

Feces, Baermann Technique

This technique separates first-stage larvae from feces. Organisms detected can include lungworm in sheep (Mullerius capillaries), cattle (Dictyocaulus viviparous), dogs (Crenosoma vulpis) and cats (Aelurostrongylus abstrusus). This method can also detect Angiostrongylus vasorum and Stronglyoides stercoralis in dogs. Fresh feces should be collected to avoid confusion with free-living nematodes. Cattle feces should be collected from the rectum. Indicate on the submission form the requirement for a Baermann technique to be done.

An important note: The Baermann technique may also be used to separate lungworm larvae from lung tissue (e.g. Muellerius capillaris in sheep and goats).

Feces, Flotation

This technique will separate from feces various species of helminth eggs (e.g. Ascaris suum in pigs), and protozoan cysts (e.g. Eimeria spp. oocysts in sheep, Giardia canis cysts in dogs). Some helminth eggs (trematode, operculate cestode, various nematode), nematode larvae (Dictyocaulus spp., Muellarius capillaris, Strongyloides stercoralis) and protozoan trophozoites (Giardia canis) generally are not detected with this technique. If infection with any of these parasites is suspected a direct smear, sedimentation or Baermann technique should be requested. Indicate on the form the need for a fecal flotation to be done.

Grading System

A high egg count may indicate a high number of parasites but a low number of eggs does not necessarily indicate a low number of parasites. A grading (non-quantitative) system is used as follows:

  • 1-100 eggs on a slide is graded one plus + 101-300 eggs on a slide is graded two plus ++
  • 301-greater on a slide is graded three plus +++

Blood Parasites

Canine Heartworm (Dirofilaria immitis)

  1. Microfilaria I.D. (Knotts Test for Microfilaria Detection)The Knotts Test is used to detect and identify circulating microfilaria of Dirofilaria immitis. It is the only microfilarial test that allows differentiation between D. immitis and the non- pathogenic Dipetalonema reconditum. It can detect infected dogs as early as 6 months post- infection. Draw off at least 1 ml of venous blood (preferably 2 – 3 ml) into a vial containing either heparin or an EDTA (lavender top). Mix the blood and anticoagulant. Store sample in the refrigerator. Submit the sample to the diagnostic laboratory along with a complete history and time and date the sample was drawn. Due to the danger of a potentially fatal drug induced shock reaction in microfilaremic dogs given diethylcarbamazine (DEC), a microfilaria test is advised on all dogs prior to the use of DEC as a heartworm preventative. Be aware, however, that approximately 25-33% of the heartworm-infected dogs will not be detected with this test.
  2. Heartworm Antigen TestThis test is an ELISA serologic test that detects circulating adult worm antigen. Submit at least 1 ml of either serum or plasma (EDTA or heparin). The test detects infected dogs as early as 6.5 months post-infection (most reliably at 8 months post infection). If only one heartworm test is to be done (Knott’s vs. Antigen Test), the antigen test is preferable to the microfilaria test. In rare instances, the antigen test can give a false negative result in a microfilaremic dog. Therefore, doing both tests offers the greatest diagnostic power to detect infection. Routine testing of dogs should begin in the month of April each year.This test can also detect microfilaria in cats. If the animal is infected with only a few parasites, however, there could be insufficient antigen present to cause a detectable reaction. In these cases, the Heartworm antibody test is recommended and is a referred test.

Skin Scrapings, Mange Mites

  1. This technique is used for lesions with minimal epidermal hyperplasia and lesions caused by deeply burrowing mites (e.g. Sarcoptes, Notoedres) or in hair follicles (e.g. Demodex). Dip a scalpel blade in mineral oil or glycerine. Using the blade, scrape the periphery of the lesions at right angles to the skin until pinpoint hemorrhaging occurs. The material collected on the scalpel blade should be pink in color. Put the scalpel blade with the oil and detritus into a sealed container, such as a small ointment jar or stoppered test tube.Note: Do not use glycerine or mineral oil on samples destined for bacteriology or mycology. Separate samples should be collected for these procedures.
  2. The following technique is used for lesions with marked epidermal hyperplasia and exfoliation and lesions caused by lice and superficially dwelling mites (e.g. Chorioptes). Scrape the dried exudate and debris into a small specimen jar.
  3. Ear mites (e.g. Otodectes) can be found easily with an otoscope. They can be removed from the external ear with a cotton swab. Place the swab in a container and submit to the laboratory.
  4. Poultry mites (e.g. Dermanyssus gallinae) do not remain on the host in daylight. The bird’s environment must be examined. Search in bird nests, roosts, and nearby cracks and crevices in housing structures. Collect and contain specimens and submit to the laboratory for identification.
  5. Some surface feeding mites (e.g. Cheyletiella) in dogs and other hosts can be collected by vigorously brushing the host over a plastic sheet. Mites and debris will accumulate on the sheet and can be transferred to a container.


  1. Large slow moving ectoparasites, such as lice, keds, ticks and possibly fleas, can be collected with forceps or fingers. Gentle, steady traction, grasping the tick by its mouthparts as close to the skin as possible, can remove ticks with their mouthparts intact. Contain the specimen in a jar along with a paper towel soaked in water. Submit to the laboratory for identification.
  2. Winged bloodsucking insects can be collected using a simple suction tube or vacuum cleaner fitted with an in-line filter or a chloroform tube placed over the parasite. Lice and occasionally fleas may be caught by this method.
  3. Various parasites such as fleas and lice can be collected from recently killed or moribund small animals by placing the animal in a closed plastic or paper bag. The ectoparasite will leave the host and can be collected in the bag.
  4. Examination of hairs or feathers can reveal nits (louse eggs) or bot fly eggs (e.g. Hypoderma, Gastrophilus).
  5. Preserving large ectoparasites (e.g. ticks, fleas, keds, lice, dipterans) is needed for shipping. Specimens should be submitted in vials or sealed plastic bags with paper towels soaked in 70% ethanol as a preservative – 10% formalin should not be used. For long-term storage utilize 70% ethanol / 5% glycerine.
  6. To preserve small ectoparasites such as mites and mallophagan lice, 70% isopropyl alcohol is used. Specimens placed in 70% ethanol usually become too brittle for processing and identification. Specimens should be shipped in small vials filled with preservative. It should be noted where specimens were collected on (or in) the host. Lots from different hosts should not be mixed.


  1. The following techniques for preserving large endoparasites for shipping. Specimens of cestodes and flukes should be fixed in buffered 10% formalin or in 70% ethanol. If flatworms are alive, they should be allowed to relax in tap water or saline prior to killing in hot (65oC) water or formalin. The relaxation is necessary as contracted specimens are usually impossible to identify. Nematodes can be handled in a similar manner. After fixation in formalin, transfer to 70% ethanol / 5% glycerine for storage. It is important to be consistent in the use of fixation techniques as different methods can modify the morphological attributes of some helminths.
  2. Lungworm larvae (e.g. Muellerius capillaris in sheep) in lung tissue, can be separated by the Baermann technique. Lung tissue must be submitted as soon as possible after collection because the lungworm larvae must be alive for this technique to work. For this same reason, the sample cannot be placed in preservative. Indicate on the submission form the requirement for a Baermann technique to be done.
  3. Gut sections and scrapings may be submitted and are potentially useful to detect situations like nematodes infestations in ruminants. Gut sections with helminths should be fixed in 10% buffered formalin or 70% ethanol. Fixation of fresh material is desirable. Note on the submission form where in the alimentary tract the sections and/or scrapings were sampled. Each section of gut or each scraping should be submitted in individual containers to prevent mixing and confusion.
  4. Eggs of pinworms (e.g. Oxyuris equi in horses) can be collected by placing scotch tape on the anal region and examining under a microscope.

Toxoplasma gondii – canine, feline, caprine, ovine
Neospora caninum – canine, bovine

Specimen Collection and Preparation

Collect a specimen of whole blood in a red-topped tube. Following clot retraction, centrifuge, aspirate the serum by pipette or syringe and place into a separate red-topped tube or plastic screw capped shipping vial. Ensure that all red blood cells are removed to prevent hemolysis during shipping. Usually about 0.3 ml of serum can be obtained from each l ml of clotted blood. Label the separated sample as serum and submit fresh if delivery within 48 hours is possible or frozen if longer transit time is likely. Serum can ALWAYS be frozen and stored before shipment. Do not freeze and thaw sera more than once. The tests require 1 ml of serum.

Do not use plasma; all testing with the kit must be performed on sera. Reactivity of the reagents with plasma is not clearly defined.

In keeping with good laboratory technique, do not use contaminated, grossly hemolyzed, lipemic or turbid specimens, which may indicate that the specimen has been exposed to deleterious conditions and/or substances.

Aquatic Samples

  1. Myxobollus cerebrallis (whirling disease samples)For population screening cranial cartilage and gill arches from a minimum of 60 fish should be submitted. Samples once collected should be frozen and shipped frozen to the Diagnostic Laboratory for analysis. Refer to suggested protocols at this website: (http://www.afs-fhs.org/bluebook/inspection-index.php )
  2. Other Fish Parasites: Refer to protocols provided at this website: ( http://www.afs-fhs.org/bluebook/inspection-index.php)