Helminth Collection and Identification from Wildlife
December 14th, 2013
Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are “roundworms” (nematodes), “thorny-headed worms” (acanthocephalans), “flukes” (trematodes), and “tapeworms” (cestodes). Here we describe how helminths are collected from a vertebrate animal and how they are preserved and taxonomically identified.
The overall goal of this procedure is to collect, preserve, and identify helmets from wildlife. This is accomplished by first necropsy, a wild animal. Next helmets are collected from different organs, then helmets are preserved using a variety of techniques for later identification.
Finally, the helmets are cleared, stained, and mounted, and the species are identified. Ultimately, results can be obtained that show specific helmet structures through clearing and or staining techniques that can greatly aid in species identification. Though this method can be applied for the identification of helmets from mammals, birds, reptiles, and amphibians.
It can also be applied for other organisms like fishes. To begin position the animal on a flat surface and use a magnifying glass to examine all external openings, including ears, oral cavity, and eyes for the presence of nematodes, CSEs, toes, and trematodes. Using a sharp blade or knife and a pair of dissecting forceps.
Make an incision over the abdominal cavity being careful not to rupture any organs. Then with bone cutters or a small handheld saw if necessary, open the thoracic cavity. Examine both cavities for fal, roundworm, and large trematodes.
Then tie a thread close to the beginning of the esophagus and another one at the end of the large intestine for examination. Under a stereo microscope, separate the heart and major blood vessels, trachea and lungs into individual Petri dishes. Remove the liver, gallbladder, and duct pancreas, spleen, kidneys, and urinary bladder, and examine them under the stereo microscope.
Then remove the complete digestive system and place each section in a glass dish. After the organs have been removed, use a hose or squirt bottle filled with 0.9%saline to wash the body cavity, allowing it to filter through a 106 micrometer mesh sieve. Backwash the contents of the sieve into a Petri dish and examine the contents under a stereo microscope For Fal worms or blood trematodes using scissors, open each digestive tract section.
Scrape the mucosa and examine it carefully for the presence of large parasites. The psci of encanto cephas are often deeply embedded in the wall of the intestine. Therefore, if necessary, use forceps and or small scissors to carefully tease them from the tissue.
Place each open section under running water and wash the contents into a 106 micrometer sieve. Then open the heart and major blood vessels, trachea and urinary and gallbladders, and examine contents In the same way, use a sharp blade and forceps to slice and tease apart solid organs such as the liver, lungs, kidney, spleen, and pancreas, and wash and examine the contents. Record the types of parasites.
Found the number of each type and the organs in which they are found. Label vials or jars by placing inside a small piece of a plain index card labeled in pencil to preserve helmets for later genetic studies. First place them directly into ethanol.
After fixing the sample for one to several days, transfer it to a five milliliter glass vial filled with 70%ethanol for long-term storage. To preserve Trematodes, place live specimens on a glass microscope, slide in a drop of saline, apply a glass cover slip, and pass the slide over a flame without boiling the saline. Then drop the slide into a Petri dish containing alcohol, formin acetic acid or a FA and let the cover slip float off.
Fix dead trematodes in a FA or 10%buffered formin for 48 hours before transferring them to a jar filled with 70%ethanol for long-term storage. To fix estos, relax live animals in a Petri dish by pouring near boiling water over them, and then transfer to a jar filled with 70%ethanol for long-term storage. Relax live encanto cephas by placing them in a Petri dish of tap water in the refrigerator for one to several hours until the proboscis is fully averted.
Transfer them to a jar filled with 70%ethanol or a FA for long-term storage. Kill small or fragile nematodes and hot 70%ethanol and preserve in a jar filled with 70%ethanol with 5%glycerin to kill. Medium to large nematodes.
Place them in cold glacial acetic acid, and after 15 minutes, transfer to a jar filled with 70%ethanol and 5%glycerin. Prepare Harris Hematin by dissolving hematin and alcohol and by dissolving ammonium aluminum sulfate in water overheat. Mix both solutions and bring to a boil.
Then add sodium iodate and boil for two to three minutes. When the solution is cooled, use 5 41 filter paper to filter the solution to overt stain Trematodes estos and decant Cephas Place in Harris Hematin or semicon Carmine overnight to detain the samples. Place in 70%acid ethanol until the organs are visible.
To halt the des staining, transfer the specimens to 70%basic ethanol for at least 30 minutes. Dehydrate the samples in an ethanol series and use xylene or methyl salicylate to clear them to mount the specimens on a microscope slide, add a drop of Canada Bossum and a cover slip. Let dry overnight clear small to medium nematodes and encanto cephas by placing them in mounts of lact phenol on a microscope slide and covering with a cover slip for 15 to 30 minutes For large nematodes and encanto Cephas.
Place in 80%phenol for up to 60 minutes. Examine under a light microscope to verify the clearing of smaller structures for examining the culex atos. Cut it off and place it on amount of lacto phenol.
On a microscope slide. Use a cover slip to squash the culex to enable measuring and counting the rose stellar hooks. Cut off the anterior end and mount on Foss on a microscope.
Slide under a few drops of lact phenol or in a glycerin jelly for permanent mount. Observe mouth parts under a light microscope. After cutting off the tails of large nematodes and mounting in lacto phenol, observe the ventral paoli and spic morphology of males under a light microscope using the methods described in this video.
A survey of the helmets of the mask shrew Sox scenarios was conducted in Missoula County, Montana. Between 2007 and 2011. A total of 56 shrews were collected from pitfall traps and examined within two hours after death.
Overall prevalence of infection was 96%Only two shrews were free of parasites. 15 species of helmets were identified including nine species of estos and six species of nematodes. One species of the Esto genus staphylo authorities was previously undescribed.
Organs found infected included the small intestine, stomach, lungs, and urinary bladder. Total intensities of infection ranged from seven to 234 worms per infected host species richness or the number of helmet species per infected host range from one to eight with a mean of 4.1 Once mastered. This technique can be done in one to two Rs on a small mammal like a squirrel, and four to five Rs on a larger mammal like a raccoon.
After watching this video, you should have a very good understanding of how to collect, preserve, and identify helmets from wildlife.
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Overview
This article describes a procedure for collecting, preserving, and identifying helminths from wild animals. The process involves necropsy, organ sampling, and various preservation techniques to aid in taxonomic identification.
Key Study Components
Area of Science
- Parasitology
- Wildlife Biology
- Helminthology
Background
- Helminths are a diverse group of parasitic worms.
- They include nematodes, acanthocephalans, trematodes, and cestodes.
- Understanding helminth diversity is crucial for wildlife health.
- Identification aids in ecological and evolutionary studies.
Purpose of Study
- To provide a standardized method for helminth collection.
- To enhance identification accuracy through preservation techniques.
- To contribute to the understanding of helminth-host interactions.
Methods Used
- Necropsy of wild animals to access internal organs.
- Collection of helminths from various organs.
- Preservation using different techniques for later analysis.
- Clearing, staining, and mounting of specimens for identification.
Main Results
- Successful collection and preservation of helminths from multiple species.
- Identification of specific structures aiding in species determination.
- Demonstrated applicability across mammals, birds, reptiles, and amphibians.
Conclusions
- The methods outlined are effective for helminth research.
- Results contribute to the broader understanding of parasitism in wildlife.
- Future studies can build on these techniques for further insights.
