Helminth Collection and Identification from Wildlife

Animals used in the following experiments were found dead as road kills.

1. Animal Necropsy and Screening of Major Organs for Helminth Collection

1. Position the animal on a flat surface and examine all external openings including ears, oral cavity and eyes with a magnifying glass for the presence of nematodes (round) and trematodes (flat and not-segmented) (see Figure 1).
2. If working with birds, wash with tap water to dampen the feathers prior to dissecting.
3. Using a sharp blade or knife and with the aid of a pair of dissecting forceps, make an incision over the abdominal cavity being careful not to rupture any organs. For turtles with a hard carapace, use a handheld saw to cut through the bone at the bridge area between the pectoral and abdominal scutes.
4. Open the thoracic cavity as above; for large animals, bone cutters or a small handheld-saw might be needed to cut through the ribs.
5. If working with birds, examine air sacs in situ prior to removing any organs. Use a magnifying glass if needed.
6. Examine both cavities for filarial roundworms and large trematodes.
7. Prior to removing the gastrointestinal tract from the abdominal cavity, tie a thread close to the beginning of the esophagus and another one at the end of the large intestine (rectum).
8. Separate the following thoracic organs into Petri dishes (or glass trays if organs are too large) for examination under a stereo microscope (10-30X): heart and major blood vessels, trachea, and lungs.
9. Remove liver, gall bladder and duct, pancreas, spleen, kidneys and urinary bladder and examine as described in step 1.8.
10. Remove the complete digestive system and place each section in glass dishes/trays.
11. After organs have been removed, wash the body cavity with a hose or squirt bottle filled with 0.9% saline into a 106 μm mesh sieve and examine the sieve contents under a stereo microscope for the presence of filarial worms or blood trematodes.
12. Open each digestive tract section with scissors, scrape the mucosa and examine carefully for the presence of large parasites (>1 cm). Use a magnifying glass if needed. The probosci of acanthocephalans are often deeply imbedded in the wall of the intestine and may have to be carefully teased from the tissue with forceps and/or small scissors.
13. Place each opened section under running water and wash all contents into a 106 μm sieve as described in step 1.11.
14. Open the heart and major blood vessels (pulmonary and aorta), trachea, and urinary and gall bladders and examine contents in the same way.
15. Slice and tease apart solid organs such as lungs, liver, kidneys, spleen, and pancreas using a sharp blade and forceps. Wash them under running water into a 106 μm sieve as described in step 1.11.
16. Record the types of parasites found, the number of each type, and the organ(s) in which they are found. Fixation and staining methods are listed below.
17. Label vials/jars by placing a small piece of a plain index card written with pencil (ink and ruled index cards will "run" in alcohol and stain specimens).

2. Preservation of Helminths

1. For helminths that need to be preserved for later genetic studies, place them directly into 80-90% ethanol (without glycerine). The ethanol should be made by diluting 95% ethanol, preferably reagent grade (Table 1). Never allow these specimens to be exposed to formalin, as formalin degrades DNA to an unusable state.
2. Place live trematodes on a glass microscope slide in a drop of saline, place a glass cover slip and pass the slide over a flame without boiling the saline.
   1. Drop the slide into a Petri dish containing AFA (alcohol-formalin-acetic acid, see Table 1) and let the cover slip float off.
   2. Fix in AFA for one to several days, rinse with tap water, and transfer to 5 ml glass vials (or larger glass vial/jar depending on size and amount of parasites) filled with 70% ethanol for long-term storage.
3. Fix dead trematodes in AFA or 10% buffered formalin for 48 hr and then transfer to vial/jar filled with 70% ethanol for long-term storage.
4. Relax live cestodes in a Petri dish by pouring near-boiling water over them and then transfer to vial/jar filled with 70% ethanol for long-term storage.
5. Relax live acanthocephalans by placing in a Petri dish for one to several hours in tap water in a refrigerator (~4 °C) until the proboscis is fully everted (this may take up to 24 hr). Transfer to vial/jar filled with 70% ethanol or AFA for long-term storage.
6. Treat dead cestodes and acanthocephalans the same as dead trematodes (see above).
7. Kill live small (<5 mm, or fragile nematodes (e.g. trichostrongylids, capillarids) in hot (near-boiling) 70% ethanol and preserve in vial/jar filled with 70% ethanol with 5% glycerine (Table 1) added to preserve the worms in case of evaporation.
8. Kill (and clear) medium to large (>5 mm) nematodes (e.g. ascarids, spirurids, oxyurids, filarids) by placing in a Petri dish with cold glacial acetic acid (Table 1) and after 15 min transfer to vial/jar filled with 70% ethanol and 5% glycerine. The acetic acid can be used repeatedly in this way.
9. Place dead nematodes directly in vial/jar filled with 70% ethanol and 5% glycerine.

3. Taxonomic Identification of Helminths

1. Prepare Harris' hematoxylin by dissolving hematoxylin in alcohol and by dissolving ammonium aluminum sulfate in water with heat. Mix both solutions and bring to a boil.
   1. Add sodium iodate and boil for 2-3 min. Filter (541 filter paper) solution after it is back to room temperature.
2. Overstain trematodes, cestodes and acanthocephalans by leaving them in diluted (1:10) Harris' hematoxylin or Semichon's carmine overnight (see Table 1).
   1. De-stain using 70% acid ethanol until organs are visible (can take between a few seconds to over an hour and should be closely observed).
   2. Halt de-staining by transferring specimens to 70% basic ethanol for at least 30 min. Next, dehydrate them in a series of graduated ethanols (80%, 95%, 100%) and clear them in xylene or methyl salicylate.
   3. Mount specimens on a microscope slide after adding a drop of Canada balsam and a cover slip. Let dry overnight.
3. Clear small to medium nematodes and acanthocephalans for 15-30 min in temporary mounts of lactophenol on a microscope slide with a cover slip.
   1. Use 80% phenol for up to 60 min for large nematodes (ascarids and spirurids) and acanthocephalans. Examine under a light microscope (40-400X) for evaluation of smaller structures (male spicules and hooks).
4. Use the following approaches for examination of special structures.
   1. For cestodes, cut off the scolex and place on a microscope slide in lactophenol and "squash" with a cover slip for measuring and counting rostellar hooks (see Figure 1B).
   2. For nematodes, cut off the anterior end and mount en face on a microscope slide in a few drops of lactophenol (without coverslip) or in glycerine jelly for a permanent mount (with cover slip) and observe mouthparts under a light microscope (35-400X) (see Figure 1D).
   3. Cut off the tails of large nematodes and mount on a microscope slide in lactophenol with a cover slip to observe the ventral papillae and spicule morphology of males under a light microscope (see Figure 1D).
5. Identify parasites to species using a combination of primary literature and standard taxonomic references such as:
   1. Trematodes: Yamaguti (1958, 1971)^3,4 and Gibson et al.^5-7
   2. Acanthocephalans: Yamaguti⁸
   3. Cestodes: Yamaguti⁹, Schmidt¹⁰, and Khalil et al.¹¹
   4. Nematodes: Yamaguti¹² and Anderson et al.¹³

4. Depository of Helminths in U.S. National Parasite Collection

1. Deposit representative helminth specimens at the U.S. National Parasite Collection (USNPC). This is mandatory for most parasite journals. After submission, each helminth species is assigned an accession number for inclusion in manuscripts.
2. Include the following information when submitting specimens: 1) Scientific name of parasite; 2) host information (species, longitude/latitude coordinates of geographic location); 3) organ found in host; 4) name of who collected the specimens and date of collection; 5) type of fixative, stain, or clearing/mounting media used; 6) location of other collections (museum and accession numbers) if additional specimens are submitted elsewhere; and 7) information on publication and journal. These data should also be sent together with the specimens and under separate cover.
3. Package specimens in strong containers with shockproof absorbent material. Tape/Parafilm screw caps on glass vials and place each vial in a separate plastic bag. Wrap slides in paper and place in a slide box with shock-absorbing material between them.
4. Ship specimens and separate letter with information about specimens to: U.S. National Parasite Collection, USDA, ARS, ANRI, Bldg. 1180, BARC-East, 10300 Baltimore Avenue, Beltsville, MD 20705-2350.