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.
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). The optimum method for collecting helminths is to examine a host that has been dead less than 4-6 hr since most helminths will still be alive. A thorough necropsy should be conducted and all major organs examined. Organs are washed over a 106 μm sieve under running water and contents examined under a stereo microscope. All helminths are counted and a representative number are fixed (either in 70% ethanol, 10% buffered formalin, or alcohol-formalin-acetic acid). For species identification, helminths are either cleared in lactophenol (nematodes and small acanthocephalans) or stained (trematodes, cestodes, and large acanthocephalans) using Harris' hematoxylin or Semichon's carmine. Helminths are keyed to species by examining different structures (e.g. male spicules in nematodes or the rostellum in cestodes). The protocols outlined here can be applied to any vertebrate animal. They require some expertise on recognizing the different organs and being able to differentiate helminths from other tissue debris or gut contents. Collection, preservation, and staining are straightforward techniques that require minimal equipment and reagents. Taxonomic identification, especially to species, can be very time consuming and might require the submission of specimens to an expert or DNA analysis.
Vertebrates are parasitized by four major groups of helminths (worms). Two of the groups, trematodes, or flukes, and cestodes, or tapeworms, fall within the Phylum Platyhelminthes. The other two groups are the nematodes, or roundworms, (Nematoda) and the acanthocephalans, or thorny-headed worms (Acanthocephala). Many of these parasites have been documented as causes of morbidity and mortality in wild birds and mammals1,2.
Most helminths have complex life cycles involving more than one host1,2 . For instance, trematodes have one or two intermediate hosts (usually invertebrates) and a final host. All hosts need to be available for the cycle to be completed and thus for adult helminths to be present in the final hosts (which are the topic of this manuscript). So it is important to keep in mind that seasonal fluctuations can occur in the prevalence and intensity of some helminth species and long-term monitoring is desirable to capture the complete helminth of auna of a particular host.
The optimum method for collecting helminths is to examine a host that has been euthanized. This allows for the collection of helminths while they are still alive and for their proper relaxation and fixation. Material from hosts that have been dead more than 24 hr, or have been frozen and thawed for examination, is often inferior and difficult to identify. Trematodes and cestodes from frozen or formalinized hosts are often badly contracted and have lost structures crucial to identification like oral spines on trematodes or the hooks on the scolex of tapeworms. However, the reality is that most material available from vertebrate hosts these days is frozen or preserved.
Databases containing DNA sequence information for helminths is quickly growing and thus taxonomic identification is already possible for many species. Therefore, helminths should be preserved for potential DNA analyses as much as possible. However, DNA of good quality is not possible if specimens are fixed in formalin. Methods outlined here for collecting and killing helminths will yield material that is suitable for DNA extraction and genetic analyses.
Below, we describe detailed methods on how to necropsy vertebrates (amphibians, reptiles, birds and mammals; monogenean trematodes from fish are not included) for the collection of helminths, followed by procedures on how to preserve and process them for taxonomic identification.
Animals used in the following experiments were found dead as road kills.
1. Animal Necropsy and Screening of Major Organs for Helminth Collection
2. Preservation of Helminths
3. Taxonomic Identification of Helminths
4. Depository of Helminths in U.S. National Parasite Collection
Using the methods outlined above, a survey of the helminths of the masked shrew, Sorex cinereus, was conducted in Missoula County, Montana between 2007-2011. A total of 56 shrews were collected from pitfall traps and examined within 2 hr after death. Overall prevalence of infection was 96%; only 2 shrews were free of parasites. Fifteen species of helminths were identified, including 9 species of cestodes and 6 species of nematodes. One species of the cestode genus Staphylocystoides was previously undescribed. Organs found infected included the small intestine (9 cestodes, 3 nematodes), stomach (1 nematode), lungs (1 nematode), and urinary bladder (1 nematode). Total intensities of infection ranged from 7-234 worms per infected host. Species richness (the number of helminth species per infected host) ranged from 1-8 with a mean of 4.1 (Figure 2).
Figure 1. Representative drawings and photographs of helminth groups showing major anatomical features for each Phylu.: A) Trematoda; B) Cestoda; C) Acanthocephala; and D) Nematoda. For the photographs, information on the scientific name, organ and host are included. Trematodes and cestodes are hermaphrodites. Drawings are not necessarily on the same scale.Click here to view larger image.
Figure 2. Summary of preliminary helminth data from masked shrews, Sorex cinereus collected from Missoula County, Montana between 2007-2011.
Click here to view larger image.
It is extremely difficult to identify helminth parasites, even to genus, based on poor material. Cestodes and trematodes, in particular, tend to die and deteriorate fairly rapidly after the death of the host. The taxonomy of cestodes depends greatly on the number, size, and shape of rostellar hooks, which are often lost in frozen material. The same applies to certain trematodes with spines around the oral sucker such as echinostomes and heterophyids, which are also frequently lost. Because of their thick cuticles, nematodes and acanthocephalans are a little hardier, but still may be contracted or have the proboscis retracted. Therefore, the importance of obtaining the freshest possible material cannot be overemphasized. Optimally, live hosts should be captured, euthanized and examined immediately. Failing that, hosts should be examined within 3-4 hr after death, or frozen as quickly as possible. Removing intestinal tracts immediately after collecting hosts and flash-freezing them with liquid nitrogen has yielded excellent material. If even a few hosts can be examined freshly and the helminths properly identified, this may allow the sorting of species collected from subsequent frozen material. Helminths can move within a host after death, and this needs to be taken into account when describing location of parasites during necropsy.
The use of a sieve to screen intestinal contents and to wash blood from other organs is a significant improvement over earlier methods of necropsy. It greatly increases the chances of finding the smaller helminths and improves quantitative analysis. Killing cestodes and trematodes in hot water yields specimens that are better for staining than those previously obtained by relaxing in cold tap water in a refrigerator.
The critical step in the staining of helminths is in the destaining process in acid ethanol and this is directly related to the type of parasite and its size. If the specimen is not destained enough, organs may not be able to be differentiated and if it is destained too much, organs may not even be visible. There is a certain art to this process and the only way to learn it is through trial and error. Specimens need to be monitored carefully under the dissecting scope while destaining. When multiple specimens of a species are available to be stained, a few may be removed from the acid ethanol at intervals to achieve a graduated series and the best specimens selected after clearing. If specimens appear to be destained too much, the whole process may be repeated by placing them back in the stain overnight.
The future of helminth taxonomy will be heavily influenced by DNA studies. The DNA database for helminth species is rapidly expanding and allows specimens collected today to be identified in the future. Therefore, it is important to preserve (in formalin-free fixative) a subset of specimens in ethanol for DNA extraction. The methods outlined here for collecting and killing helminths will yield material that is suitable for DNA extraction and genetic analyses.
The authors have nothing to disclose.
The authors would like to thank Dr. Joe N. Caudell, Disease Biologist for the Indiana USDA APHIS Wildlife Services Program at Purdue University, for providing specimens used for the collection of helminths for the production of this video. Jennifer Serafin prepared all of the chemical solutions.
Fixing | |||
Alcohol-Formalin-Acetic Acid (AFA) | Fisher Scientific | A407-1 (ethanol) F75F-1GAL (formaldehyde) BP2401500 (glacial acetic acid) |
Ethanol 85% (85 ml) + formaldehyde 37% (10 ml) + glacial acetic acid (5 ml) |
Ethanol for killing and long-term preservation | Fisher Scientific | A407-1 | Ethanol 100% (70 ml) + distilled water (30 ml) |
Ethanol for fixing and DNA studies | Fisher Scientific | AC61511-0010 | Ethanol 80-90% (80-90 ml) + distilled water (20-10 ml), respectively |
Formalin Buffered | Fisher Scientific | SF100-4 | Formalin 10% (10 ml) + distilled water (90 ml) |
Glycerine-alcohol | Fisher Scientific | A407-1 (ethanol) AC15365-1000 (glycerine) |
Ethanol 70% (95 ml) + glycerine (5 ml) |
Clearing, staining and mounting | |||
Ethanol for dehydrating during staining | Fisher Scientific | A407-1 | Ethanol 80%, 95%, 100% (80 ml, 95 ml, 100 ml) + distilled water (20 ml, 5 ml, 0 ml) |
Lactophenol | Fisher Scientific | R400-27 | – |
Phenol | Fisher Scientific | A931I-1 (phenol) A407-1 (ethanol) |
Phenol 100% (80 ml) + ethanol 100% (20 ml) |
Harris’ Hematoxylin | Fisher Scientific | S25347 (hematoxylin) A407-1 (ethanol) A567-500 (ammonium aluminum sulfate) S25553 (sodium iodate) |
Hematoxylin (5 g) + ethanol 100% (50 ml) + ammonium aluminum sulfate (100 g) + sodium iodate (0.37 g) + distilled water (1,000 ml) |
Semichon’s acetic carmine | Fisher Scientific | BP2401500 (glacial acetic acid) S25236 (carmine) A407-1 (ethanol) |
Glacial acetic acid (100 ml) + carmine (1.5 g) + distilled water (100 ml). Heat in a boiling water bath for 15 min and cool. Filter to make stock solution. Mix 1:2 with 70% alcohol for staining. |
Xylene | Fisher Scientific | X4-4 | Use only under a chemical fume hood. |
Methyl salicylate | Fisher Scientific | S25437 | – |
Ethanol basic | Fisher Scientific | A407-1 (ethanol) S25159A (concentrated ammonium hydroxide) |
Ethanol 70% (100 ml) + ammonium hydroxide (0.1 ml) |
Ethanol acid | Fisher Scientific | A407-1 (ethanol) SA9233-100 (hydrochloric acid) |
Ethanol 70% (99.5 ml) + hydrochloric acid (0.5 ml) |
Canada balsam | Fisher Scientific | B10-100 | – |