Veterinary Parasites Laboratory Procedures

Rev. 10/06/2004

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Collection of Samples

  1. Feces
  2. Blood

Collection of Intestinal Parasites

1. Direct Fecal Smear

Very good technique for identification of motile parasites commonly used for diarrheic or mucoid fecal samples. Use feces that is as fresh as possible.

  1. Place a drop of saline solution on a clean glass slide
  2. Collect a rectal sample of feces using gloved finger (or touch gloved finger or toothpick to fresh sample already collected)
  3. Smear very thinly onto clean slide and cover with coverslip
  4. May use a stain such as Ziehl-Neelsen, carbol-fuchsin, or Giemsa (optional-not used often in private veterinary practices)
  5. Use less light on the microscope for examination due to clear color of most unstained parasites

Negative results are inconclusive, perform a concentration technique such as flotation on all samples that are negative direct smear.

Make sure that the sample on the slide is very thin (you should be able to read print through the sample), if the fecal layer is too thick, it becomes difficult to identify movement of parasites.

2. Flotation Fluids

Since parasite eggs will sink in water, salt or sugar solutions are used to concentrate and separate eggs from most fecal debris.

The most commonly used flotation fluids are magnesium sulfate (Epsom salts), sugar, sodium nitrate, and zinc sulfate. A specific gravity from 1.2 to 1.3 is best for floating most eggs.

Each solution has advantages and disadvantages. Magnesium sulfate is inexpensive, but if slides have to sit a while before they are read, the fluid will crystallize and eggs may be distorted. Sugar solution allows slides to be kept longer before reading, but is sticky and may be more expensive. Sodium nitrate can be purchased already in solution and therefore saves time used for mixing, but it is relatively expensive. Zinc sulfate is the best solution to use for the detection of Giardia cysts because the cysts do not become distorted as quickly with it.

To prepare flotation fluids:

3. Centrifugation with Magnesium Sulfate Flotation Fluid

Using a centrifuge reduces the number of eggs that rise slowly to the surface of a flotation setup. Thus reducing the number of false negative fecal examinations.

To be used for concentrating protozoan cysts, nematode, cestode and some arthropod eggs. (Not usually used for an egg which has an operculum)

Materials

Procedure

Note: for herbivores or other feces with a lot of debris (e.g. cat feces with adherent litter), strain feces by mixing with a little water in a paper cup, add MgSO4, mix, pour through gauze into another cup or centrifuge tube, then proceed as above.

4. Modified Wisconsin Procedure

(for cattle, horses, dogs, cats, and swine)

The modified Wisconsin procedure for egg counts by a flotation method is used to quantify the amount of parasites in an individual animal (cattle, horses, dogs, cats, swine)

  1. Weigh out a 2 g fecal sample (5 g for cows) in a paper cup
  2. Place 10-cc water in the paper cup with the fecal material.
  3. Stir very well with a spatula and mash the material until it is completely broken apart
  4. Pour the mixture through gauze or strainer ( while it is well mixed ) into another cup, stirring the material in the strainer while pouring.
  5. Press the material remaining in the strainer with the spatula until nearly dry
  6. Add a small amount of water to the paper cup just emptied and rinse into a mixture the material clinging to the sides and bottom, and then pour this mixture through the material in the strainer, stirring the material in the strainer while pouring.
  7. Press the material in the strainer until dry again, then discard.
  8. Stir the material in the cup that was under the strainer and immediately pour the contents of the cup into one 15-ml tube. If the tube is not too full, squirt water down the sides of the cup in sufficient amounts to remove the material clinging to it and finish filling up the tube.
  9. Centrifuge the tube at 1500 rpm for 10 minutes (not including the time required for acceleration and deceleration)
  10. Decant the tube, being careful tot to pour off the fine material at the top of the sediment
  11. Fill the tube ½ full of MgSO4, (sp.gr. 1.2-1.25) and mix the sediment ant the MgSO4 solution with an applicator stick, being careful to scrape the sides and bottom of the tube to insure the removal and mixing of all material.
  12. Finish filling the tube with flotation fluid
  13. With a medicine dropper, add MgSO4 to the tube until it is full enough so that a 22-mm square coverglass can be placed on the top. (there should neither be an air bubble under the cover slip, nor should the material overflow so that it runs down the side of the tube)
  14. Centrifuge at 1500 rpm for 10 minutes (not including time required for acceleration and deceleration)
  15. Remove the coverglass by lifting straight upward and place it on a glass slide. If properly done, there should be a good thickness of material under the coverglass.
  16. Count the all of the worm eggs under the entire coverglass using a low power (10x) objective.
  17. Record the results very carefully giving (a) the specimen number, (b) the date of collection, (c) the number of worm eggs of each type seen

The count is number of eggs per 2 (or 5) grams of feces.

5. Significance of Eggs Per Gram Counts

  1. The number of parasite eggs per gram feces is influenced by:

     

  2. Significant numbers of parasite eggs vary between host species and parasite types:

6. Fecal Culture

The following method is a technique for culturing eggs of nematodes of the order Strongylida to infective L 3 larvae and then recovering these larvae for identification. It involves a culture phase, which minimizes the time for development to the L 3 and one of the many modifications of the Baermann technique for recovery of the larvae. The Baermann technique takes advantage of the fact that the L 3 will migrate out of a fecal mass into a fluid medium but being unable to swim against gravity will then settle to the bottom of the medium container.

Materials:

  1. Examination gloves and specimen containers
  2. Lab balance
  3. Dried sterile sphagnum moss ("Nodampoff" sphagnum moss is a pre-sterilized seed-starting medium available through lawn and garden shops)
  4. 250 ml glass beakers
  5. wooden applicator sticks
  6. 100 x 25 mm disposable petri dishes with covers
  7. 150 x 25 disposable petri dishes with covers
  8. cheesecloth or gauze
  9. rubber bands
  10. Lugol’s iodine solution
  11. Centrifuge with centrifuge tubes
  12. Pasteur pipettes
  13. Microscope with microslides and coverslips

Procedure:

  1. Collect at least 10 gm feces rectally from each animal to be checked using clean gloves and clean specimen containers (this is important to prevent contamination with free-living nonparasitic nematodes)
  2. If the specimen is from cattle, swine or dog , weigh out a minimum of 10-g sphagnum moss for each specimen to be examined. Place the moss in a suitable container and mix with sufficient warm tap water to produce a thick "soup". Allow the moss to become thoroughly saturated and then squeeze out all excess water.
  3. Weigh out 5 g feces into a 250 ml beaker and thoroughly break up the sample with applicator sticks
  4. If the specimen is from cattle swine or dog , add 15 g saturated sphagnum moss to the beaker and mix thoroughly. Make sure that none of the feces is left adhering to the sides of the beaker. Horse, sheep and goat feces do not require the addition of moss
  5. Place the feces/moss mixture in a 100 x 25-mm petri dish and compress lightly . The particles of the material should be in uniform contact with each other but remain "fluffy" . Cover and incubate at room temperature for 14 days. (If an incubator is available, horse feces may be incubated for 7 days t 30 degrees C.) Do not be alarmed if mold forms on sample during incubation
  6. Following incubation, remove the cover of the dish and place 2 layers of cheesecloth or gauze over the top of the dish. Stretch the cloth and secure with a rubber band. Trim away excess cloth and replace cover. Invert the dish and tap bottom smartly to dislodge the sample onto the cloth.
  7. Place 2 wooden applicator sticks in a 150 x 25-mm disposable petri dish and add 100-ml warm tap water. Keeping the 100-mm petri dish inverted, remove its cover and place it cloth side down onto the applicator sticks in the 150-mm petri dish. Gently press on the bottom of the 100-mm dish until a small amount of air escapes from under the rim and then release. Cover and let stand (Baermannize) at room temperature for 18 to 24 hours.
  8. Following Baermannization, remove the 100-mm dish and the wooden applicator sticks and discard. Add 0.3 ml of Lugol’s iodine solution to the liquid in the 150-mm dish and agitate gently. Pour the mixture into a centrifuge tube and centrifuge at 1000 to 1500 rpm for 10 minutes. (If a centrifuge is not available, the liquid can be poured into an Imhoff settling cone or similar cone-shaped container and allowed to stand for 10 minutes.
  9. Transfer a drop of sediment from the bottom of one of the centrifuge tubes with a Pasteur pipette to a microslide. Add a coverslip and identify the L3 larvae present under the Microscope.
  10. Descriptions of the various larvae are provided in Georgi, Parasitology for Veterinarians , 6 th ed. (1995), Dunn, Veterinary Helminthology , 2 nd ed. (1978), or Levine, Nematode Parasites of Domestic Animals and Man , (1980)

Collection of Blood Parasites

1. Knott's Method

The modified Knott’s method is used for the concentration and identification of microfilaria.

Procedure

  1. Add 1-ml blood to 10 ml of 2% formalin and mix.
  2. Centrifuge for 5 minutes at 1000 to 1500 rpm.
  3. Pour off supernatant fluid. Note: The tube may be inverted on a paper towel to allow all the liquid to drain.
  4. Mix sediment with equal volume of 1:1000 aqueous methylene blue.
  5. Examine as wet mount.

Collection of Muscle Parasites

1. Squash Preparation

Used to identify Trichinella spp cysts within muscle:

  1. Collect a small amount of fresh muscle
  2. Place a small amount of tissue on a glass slide
  3. Cover with a second glass slide
  4. Press the two slides together using thumb and index finger
  5. While still holding slides together, tape both ends of the slides together (scotch tape works well for this)
  6. Trim away tissue not contained by the two slides
  7. Examine with a microscope using low power to identify larval cysts within the muscle


Copyright © 1997 all rights reserved.
Published by RM Corwin and Julie Nahm,
University of Missouri College of Veterinary Medicine.

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