Name: helminth collection and identification from wildlife
Uploaded: 2013-12-14T20:15:00.000+00:00
Duration: 9 min 37 s
Description: Watch this Scientific Journal Video about Helminth Collection and Identification from Wildlife at JoVE.com

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.

Animals used in the following experiments were found dead as road kills.
1. Animal Necropsy and Screening of Major Organs for Helminth Collection
<ol>
	<li>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&#160;1).</li>
	<li>If working with birds, wash with tap water to dampen the feathers prior to dissecting.</li>
	<li>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.</li>
	<li>Open the thoracic cavity as above; for large animals, bone cutters or a small handheld-saw might be needed to cut through the ribs.</li>
	<li>If working with birds, examine air sacs in situ prior to removing any organs. Use a magnifying glass if needed.</li>
	<li>Examine both cavities for filarial roundworms and large trematodes.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Remove liver, gall bladder and duct, pancreas, spleen, kidneys and urinary bladder and examine as described in step 1.8.</li>
	<li>Remove the complete digestive system and place each section in glass dishes/trays.</li>
	<li>After organs have been removed, wash the body cavity with a hose or squirt bottle filled with 0.9% saline into a 106 &#956;m mesh sieve and examine the sieve contents under a stereo microscope for the presence of filarial worms or blood trematodes.</li>
	<li>Open each digestive tract section with scissors, scrape the mucosa and examine carefully for the presence of large parasites (&#62;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.</li>
	<li>Place each opened section under running water and wash all contents into a 106 &#956;m sieve as described in step 1.11.</li>
	<li>Open the heart and major blood vessels (pulmonary and aorta), trachea, and urinary and gall bladders and examine contents in the same way.</li>
	<li>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 &#956;m sieve as described in step 1.11.</li>
	<li>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.</li>
	<li>Label vials/jars by placing a small piece of a plain index card written with pencil (ink and ruled index cards will &#34;run&#34; in alcohol and stain specimens).</li>
</ol>
2. Preservation of Helminths
<ol>
	<li>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.</li>
	<li>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.
	<ol>
		<li>Drop the slide into a Petri dish containing AFA (alcohol-formalin-acetic acid, see Table 1) and let the cover slip float off.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>Relax live acanthocephalans by placing in a Petri dish for one to several hours in tap water in a refrigerator (~4 &#176;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.</li>
	<li>Treat dead cestodes and acanthocephalans the same as dead trematodes (see above).</li>
	<li>Kill live small (&#60;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.</li>
	<li>Kill (and clear) medium to large (&#62;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.</li>
	<li>Place dead nematodes directly in vial/jar filled with 70% ethanol and 5% glycerine.</li>
</ol>
3. Taxonomic Identification of Helminths
<ol>
	<li>Prepare Harris&#39; hematoxylin by dissolving hematoxylin in alcohol and by dissolving ammonium aluminum sulfate in water with heat. Mix both solutions and bring to a boil.
	<ol>
		<li>Add sodium iodate and boil for 2-3 min. Filter (541 filter paper) solution after it is back to room temperature.</li>
	</ol>
	</li>
	<li>Overstain trematodes, cestodes and acanthocephalans by leaving them in diluted (1:10) Harris&#39; hematoxylin or Semichon&#39;s carmine overnight (see Table 1).
	<ol>
		<li>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).</li>
		<li>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.</li>
		<li>Mount specimens on a microscope slide after adding a drop of Canada balsam and a cover slip. Let dry overnight.</li>
	</ol>
	</li>
	<li>Clear small to medium nematodes and acanthocephalans for 15-30 min in temporary mounts of lactophenol on a microscope slide with a cover slip.
	<ol>
		<li>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).</li>
	</ol>
	</li>
	<li>Use the following approaches for examination of special structures.
	<ol>
		<li>For cestodes, cut off the scolex and place on a microscope slide in lactophenol and &#34;squash&#34; with a cover slip for measuring and counting rostellar hooks (see Figure 1B).</li>
		<li>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).</li>
		<li>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).</li>
	</ol>
	</li>
	<li>Identify parasites to species using a combination of primary literature and standard taxonomic references such as:
	<ol>
		<li>Trematodes: Yamaguti (1958, 1971)3,4 and Gibson et al.5-7</li>
		<li>Acanthocephalans: Yamaguti8</li>
		<li>Cestodes: Yamaguti9, Schmidt10, and Khalil et al.11</li>
		<li>Nematodes: Yamaguti12 and Anderson et al.13</li>
	</ol>
	</li>
</ol>
4. Depository of Helminths in U.S. National Parasite Collection
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>

<ol>
	<li>Atkinson, C. T., Thomas, N. J., Hunter, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Parasitic Diseases of Wild Birds. , (2009).</li><li>Samuel, W. M., Pybus, M. J., Kocan, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Parasitic Diseases of Wild Mammals. , (2001).</li><li>Yamaguti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parts+1+and+2.+The+Digenetic+Trematodes+of+Vertebrates.">Parts 1 and 2. The Digenetic Trematodes of Vertebrates.</a> Systema Helminthum. I, (1958).</li><li>Yamaguti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Synopsis of Digenetic Trematodes of Vertebrates. I and II, (1971).</li><li>Gibson, D. I., Jones, A., Bray, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Keys to the Trematoda. 1, (2002).</li><li>Jones, A., Bray, R. A., Gibson, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Keys to the Trematoda. 2, (2005).</li><li>Bray, R. A., Gibson, D. I., Jones, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Keys to the Trematoda. 3, (2008).</li><li>Yamaguti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acanthocephala+of+Vertebrates.">Acanthocephala of Vertebrates.</a> Systema Helminthum. V, (1963).</li><li>Yamaguti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Cestodes+of+Vertebrates.">The Cestodes of Vertebrates.</a> Systema Helminthum. II, (1959).</li><li>D, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Keys+to+the+Cestode+Parasites+of+Vertebrates.">Keys to the Cestode Parasites of Vertebrates.</a> CRC Handbook of Tapeworm Identification. , (1986).</li><li>Khalil, L. F., Jones, A., Bray, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Keys to the Cestode Parasites of Vertebrates. , (1994).</li><li>Yamaguti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Nematodes+of+Vertebrates.+Parts+1+and+2.">The Nematodes of Vertebrates. Parts 1 and 2.</a> Systema Helminthum. III, (1961).</li><li>Anderson, R. C., Chabaud, A. G., Willmott, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Keys to the Nematode Parasites of Vertebrates: Archival Volume (Nos). 1-10. , (2009).</li></ol>

We have nothing to disclose

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, nemato...

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.

<table><tbody><tr><td>&lt;strong&gt;Fixing&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Alcohol-Formalin-Acetic Acid (AFA)</td><td>Fisher Scientific</td><td>A407-1 (ethanol)&lt;br/&gt; F75F-1GAL (formaldehyde)&lt;br/&gt; BP2401500 (glacial acetic acid)</td><td>Ethanol 85% (85 ml) + formaldehyde 37% (10 ml) + glacial acetic acid (5 ml)</td></tr><tr><td>Ethanol for killing and long-term preservation</td><td>Fisher Scientific</td><td>A407-1</td><td>Ethanol 100% (70 ml) + distilled water (30 ml)</td></tr><tr><td>Ethanol for fixing and DNA studies</td><td>Fisher Scientific</td><td>AC61511-0010</td><td>Ethanol 80-90% (80-90 ml) + distilled water (20-10 ml), respectively</td></tr><tr><td>Formalin Buffered</td><td>Fisher Scientific</td><td>SF100-4</td><td>Formalin 10% (10 ml) + distilled water (90 ml)</td></tr><tr><td>Glycerine-alcohol</td><td>Fisher Scientific</td><td>A407-1 (ethanol)&lt;br/&gt; AC15365-1000&lt;br/&gt; (glycerine)</td><td>Ethanol 70% (95 ml) + glycerine (5 ml)</td></tr><tr><td>&lt;strong&gt;Clearing, staining and mounting&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ethanol for dehydrating during staining</td><td>Fisher Scientific</td><td>A407-1</td><td>Ethanol 80%, 95%, 100% (80 ml, 95 ml, 100 ml) + distilled water (20 ml, 5 ml, 0 ml)</td></tr><tr><td>Lactophenol</td><td>Fisher Scientific</td><td>R400-27</td><td>-</td></tr><tr><td>Phenol</td><td>Fisher Scientific</td><td>A931I-1 (phenol)&lt;br/&gt; A407-1 (ethanol)</td><td>Phenol 100% (80 ml) + ethanol 100% (20 ml)</td></tr><tr><td>Harris&amp;rsquo; Hematoxylin</td><td>Fisher Scientific</td><td>S25347 (hematoxylin)&lt;br/&gt; A407-1 (ethanol)&lt;br/&gt; A567-500 (ammonium aluminum sulfate) S25553 (sodium iodate)</td><td>Hematoxylin (5 g) + ethanol 100% (50 ml) + ammonium aluminum sulfate (100 g) + sodium iodate (0.37 g) + distilled water (1,000 ml)&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Semichon&amp;rsquo;s acetic carmine</td><td>Fisher Scientific</td><td>BP2401500 (glacial acetic acid)&lt;br/&gt; S25236 (carmine)&lt;br/&gt; A407-1 (ethanol)</td><td>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.</td></tr><tr><td>Xylene</td><td>Fisher Scientific</td><td>X4-4</td><td>Use only under a chemical fume hood.</td></tr><tr><td>Methyl salicylate</td><td>Fisher Scientific</td><td>S25437</td><td>-</td></tr><tr><td>Ethanol basic</td><td>Fisher Scientific</td><td>A407-1 (ethanol)&lt;br/&gt; S25159A (concentrated ammonium hydroxide)</td><td>Ethanol 70% (100 ml) + ammonium hydroxide (0.1 ml)</td></tr><tr><td>Ethanol acid</td><td>Fisher Scientific</td><td>A407-1 (ethanol)&lt;br/&gt; SA9233-100 (hydrochloric acid)</td><td>Ethanol 70% (99.5 ml) +&amp;nbsp; hydrochloric acid (0.5 ml)</td></tr><tr><td>Canada balsam</td><td>Fisher Scientific</td><td>B10-100</td><td>-</td></tr></tbody></table>

helminth collection and identification from wildlife

Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are &#34;roundworms&#34; (nematodes), &#34;thorny-headed worms&#34; (acanthocephalans), &#34;flukes&#34; (trematodes), and &#34;tapeworms&#34; (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 &#956;m 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&#39; hematoxylin or Semichon&#39;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.

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 undesc...

Watch this Scientific Journal Video about Helminth Collection and Identification from Wildlife at JoVE.com

Helminth Collection and Identification from Wildlife

Wild animals are commonly parasitized by a wide range of helminths.&#160; The four major types of helminths are &#8220;roundworms&#8221; (nematodes), &#8220;thorny-headed worms&#8221; (acanthocephalans), &#8220;flukes&#8221; (trematodes), and &#8220;tapeworms&#8221; (cestodes).&#160; Here we describe how helminths are collected from a vertebrate animal and how they are preserved and taxonomically identified.&#160;

Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are &#34;roundworms&#34; (nematodes), ...

helminth-collection-and-identification-from-wildlife

Research

JoVE Journal

Environment

15.5K Views.  Purdue University. 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. 

Video: Helminth Collection and Identification from Wildlife

Soil Sampling and Isolation of Entomopathogenic Nematodes (Steinernematidae, Heterorhabditidae)

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to two families: Steinernematidae and Heterorhabditidae. Until now, more than 70 species have been described in the Steinernematidae and there are about 20 species in the Heterorhabditidae. The nematodes have a mutualistic partnership with Enterobacteriaceae bacteria and together they act as a potent insecticidal complex that kills a wide range of insect species.
Herein, we focus on the most common techniques considered for collecting EPN from soil. The second part of this presentation focuses on the insect-baiting technique, a widely used approach for the isolation of EPN from soil samples, and the modified White trap technique which is used for the recovery of these nematodes from infected insects. These methods and techniques are key steps for the successful establishment of EPN cultures in the laboratory and also form the basis for other bioassays that consider these nematodes as model organisms for research in other biological disciplines. The techniques shown in this presentation correspond to those performed and/or designed by members of S. P. Stock laboratory as well as those described by various authors.

Soil Sampling and Isolation of Entomopathogenic Nematodes Steinernematidae, Heterorhabditidae

Entomopathogenic nematodes are soil-inhabiting roundworms that parasitize a wide range of insects. We demonstrate sampling methods used for the isolation of these nematodes from the soil using two techniques: the insect baiting and the modified White trap, for the recovery of nematodes from soil samples and infected insect cadavers, respectively.

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to ...

Use of Chironomidae (Diptera) Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies

Rapid bioassessment protocols using benthic macroinvertebrate assemblages have been successfully used to assess human impacts on water quality. Unfortunately, traditional benthic larval sampling methods, such as the dip-net, can be time-consuming and expensive. An alternative protocol involves collection of Chironomidae surface-floating pupal exuviae (SFPE). Chironomidae is a species-rich family of flies (Diptera) whose immature stages typically occur in aquatic habitats. Adult chironomids emerge from the water, leaving their pupal skins, or exuviae, floating on the water&#8217;s surface. Exuviae often accumulate along banks or behind obstructions by action of the wind or water current, where they can be collected to assess chironomid diversity and richness. Chironomids can be used as important biological indicators, since some species are more tolerant to pollution than others. Therefore, the relative abundance and species composition of collected SFPE reflect changes in water quality. Here, methods associated with field collection, laboratory processing, slide mounting, and identification of chironomid SFPE are described in detail. Advantages of the SFPE method include minimal disturbance at a sampling area, efficient and economical sample collection and laboratory processing, ease of identification, applicability in nearly all aquatic environments, and a potentially more sensitive measure of ecosystem stress. Limitations include the inability to determine larval microhabitat use and inability to identify pupal exuviae to species if they have not been associated with adult males.

Use of Chironomidae Diptera Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies

Rapid bioassessment protocols using benthic macroinvertebrates are often used to monitor and assess water quality. An efficient protocol involves collections of Chironomidae surface-floating pupal exuviae (SFPE). Here, techniques for field collection, laboratory processing, slide mounting, and identification of Chironomidae SFPE are described.

Rapid bioassessment protocols using benthic macroinvertebrate assemblages have been successfully used to assess human impacts on water quality. ...

Diagnosis of Ecto- and Endoparasites in Laboratory Rats and Mice

Internal and external parasites remain a significant concern in laboratory rodent facilities, and many research facilities harbor some parasitized animals. Before embarking on an examination of animals for parasites, two things should be considered. One: what use will be made of the information collected, and two: which test is the most appropriate. Knowing that animals are parasitized may be something that the facility accepts, but there is often a need to treat animals and then to determine the efficacy of treatment. Parasites may be detected in animals through various techniques, including samples taken from live or euthanized animals. Historically, the tests with the greatest diagnostic sensitivity required euthanasia of the animal, although PCR has allowed high-sensitivity testing for several types of parasite. This article demonstrates procedures for the detection of endo- and ectoparasites in mice and rats. The same procedures are applicable to other rodents, although the species of parasites found will differ.

This article describes various procedures for screening rats and mice to detect endo- or ectoparasitism. Several diagnostic assays will be demonstrated, both those suitable for use on live animals and those used after euthanasia of the animal. Photographs to aid in identification of rat and mouse parasites will be included.

Internal and external parasites remain a significant concern in laboratory rodent facilities, and many research facilities harbor some parasitized ...

Immunology and Infection

Rapid Isolation of Wild Nematodes by Baermann Funnel

Beyond being robust experimental model organisms, Caenorhabditis elegans and its relatives are also real animals that live in nature. Studies of wild nematodes in their natural environments are valuable for understanding many aspects of biology, including the selective regimes in which distinctive genomic and phenotypic characters evolve, the genetic basis for complex trait variation, and the natural genetic diversity fundamental to all animal populations. This manuscript describes a simple and efficient method for extracting nematodes from their natural substrates, including rotting fruits, flowers, fungi, leaf litter, and soil. The Baermann funnel method, a classical nematology technique, selectively isolates active nematodes from their substrates. Because it recovers nearly all active worms from the sample, the Baermann funnel technique allows for the recovery of rare and slow-growing genotypes that co-occur with abundant and fast-growing genotypes, which might be missed in extraction methods that involve multiple generations of reproduction. The technique is also well suited to addressing metagenetic, population-genetic, and ecological questions. It captures the entire population in a sample simultaneously, allowing an unbiased view of the natural distribution of ages, sexes, and genotypes. The protocol allows for deployment at scale in the field, rapidly converting substrates into worm plates, and the authors have validated it through fieldwork on multiple continents.

This protocol outlines a method for efficiently extracting live nematodes from natural substrates in the field.

Beyond being robust experimental model organisms, Caenorhabditis elegans and its relatives are also real animals that live in nature. Studies of wild ...

Genetics

Cultivation of Heligmosomoides Polygyrus: An Immunomodulatory Nematode Parasite and its Secreted Products

Heligmosomoides polygyrus (formerly known as Nematospiroides dubius, and also referred to by some as H. bakeri) is a gastrointestinal helminth that employs multiple immunomodulatory mechanisms to establish chronic infection in mice and closely resembles prevalent human helminth infections. H. polygyrus has been studied extensively in the field of helminth-derived immune regulation and has been found to potently suppress experimental models of allergy and autoimmunity (both with active infection and isolated secreted products). The protocol described in this paper outlines management of the H. polygyrus life cycle for consistent production of L3 larvae, recovery of adult parasites, and collection of their excretory-secretory products (HES).

Cultivation of Heligmosomoides Polygyrus: An Immunomodulatory Nematode Parasite and its Secreted Products

Heligmosomoides polygyrus is a murine nematode with powerful immunomodulatory capabilities that closely resemble those of highly-prevalent human helminth infection. Here we describe a protocol for the long-term maintenance of the H. polygyrus lifecycle.

Heligmosomoides polygyrus (formerly known as Nematospiroides dubius, and also referred to by some as H. bakeri) is a gastrointestinal helminth that ...

Diagnostic Necropsy and Selected Tissue and Sample Collection in Rats and Mice

There are multiple sample types that may be collected from a euthanized animal in order to help diagnose or discover infectious agents in an animal colony. Proper collection of tissues for further histological processing can impact the quality of testing results. This article describes the conduct of a basic gross examination including identification of heart, liver, lungs, kidneys, and spleen, as well as how to collect those organs. Additionally four of the more difficult tissue/sample collection techniques are demonstrated. Lung collection and perfusion can be particularly challenging as the tissue needs to be properly inflated with a fixative in order for inside of the tissue to fix properly and to enable thorough histologic evaluation. This article demonstrates the step by step technique to remove the lung and inflate it with fixative in order to achieve optimal fixation of the tissue within 24 hours. Brain collection can be similarly challenging as the tissue is soft and easily damaged. This article demonstrates the step by step technique to expose and remove the brain from the skull with minimal damage to the tissue. The mesenteric lymph node is a good sample type in which to detect many common infectious agents as enteric viruses persist longer in the lymph node than they are shed in feces. This article demonstrates the step by step procedure for locating and aseptically removing the mesenteric lymph node. Finally, identification of infectious agents of the respiratory tract may be performed by bacterial culture or PCR testing of nasal and/or bronchial fluid aspirates taken at necropsy. This procedure describes obtaining and preparing the respiratory aspirate sample for bacterial culture and PCR testing.

This article describes the procedures for conducting a basic postmortem examination of a mouse or rat, and the collection of basic organs, as well as more challenging sample types from for histological, microbiological, and PCR evaluation.

There are multiple sample types that may be collected from a euthanized animal in order to help diagnose or discover infectious agents in an animal ...

Biology

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Caenorhabditis elegans (C. elegans)&#160;has proven to be an excellent model for studying host-microbe interactions and the microbiome, especially in the context of the intestines. Recently, ecological sampling of wild Caenorhabditis nematodes has discovered a diverse array of associated microbes, including bacteria, viruses, fungi, and microsporidia. Many of these microbes have interesting colonization or infection phenotypes that warrant further study, but they are often unculturable. This protocol presents a method to enrich the desired intestinal microbes in C. elegans and related nematodes and reduce the presence of the many contaminating microbes adhering to the cuticle. This protocol involves forcing animals into the dauer stage of development and using a series of antibiotic and detergent washes to remove external contamination. As dauer animals have physiological changes that protect nematodes from harsh environmental conditions, any intestinal microbes will be protected from these conditions. But, for enrichment to work, the microbe of interest must be maintained when animals develop into dauers. When the animals leave the dauer stage, they are singly propagated into individual lines. F1 populations are then selected for desired microbes or infection phenotypes and against visible contamination. These methods will allow researchers to enrich unculturable microbes in the intestinal lumen, which make up part of the natural microbiome of C. elegans and intracellular intestinal pathogens. These microbes can then be studied for colonization or infection phenotypes and their effects on the host fitness.

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Wild Caenorhabditis nematodes are associated with many microbes, often in the gut lumen or infecting the intestine. This protocol details a method to enrich unculturable microbes colonizing the intestine, taking advantage of the resistance of the dauer cuticle.

Caenorhabditis elegans (C. elegans)&#160;has proven to be an excellent model for studying host-microbe interactions and the microbiome, especially in the ...

Recognition of Zoonotic Roundworms by a Fecal Flotation Technique

A Simple Fecal Flotation Method for Diagnosing Zoonotic Nematodes Under Field and Laboratory Conditions

Diagnosis of canine parasites with zoonotic potential such as Toxocara canis and Ancylostoma caninum under field conditions is usually difficult due to limited access to a laboratory in rural and suburban areas in Mexico. This study aimed to detect T. canis and Ancylostoma spp. in fecal samples collected from dogs in Mexico from 2017 to 2021 under field conditions. Sample size calculation resulted in a target enrollment of 534 dogs across the country.
Samples were collected directly from the rectum or the ground after defecation. Samples were stored in individual, tightly sealed, plastic bags at 4 &#176;C. A saturated solution of sodium chloride (specific gravity [SpG] 1.20) was prepared both under field and laboratory conditions. Within 3 days of collection, 2-4 g of feces were tested for parasites using a flotation method by suspending each fecal sample in a saline solution. Feces were mixed with the flotation solution and crushed using a metal spoon.
Once a uniform consistency was achieved, the fecal sample was poured into a new plastic cup using a sieve and allowed to sit for 10-15 min. Three drops from the top of the mixture were collected using a sterilized inoculating loop. The slides were placed on the microscope and parasites were identified by trained parasitologists. Fecal samples from 1,055 dogs were screened microscopically. The number of positive samples for Ancylostoma spp.&#160;was 833 (78.95% frequency) and 222 (21.04%) for T. canis. These findings illustrate the importance of identifying zoonotic helminths in dogs living in urban and rural areas in Mexico using a coproparasitoscopic technique in the laboratory and under field conditions.

Author Spotlight: Coproparasitoscopic Examination of Dog Stools for Control and Prevention of Zoonotic Parasite Diseases

This work describes the use of a flotation method to identify Toxocara canis and Ancylostoma spp. detected in fecal samples collected from dogs in Mexico from 2017 to 2021 under field conditions.

Diagnosis of canine parasites with zoonotic potential such as Toxocara canis and Ancylostoma caninum under field conditions is usually difficult due to ...

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

Caenorhabditis elegans is one of the major model organisms in biology, but only recently have researchers focused on its natural ecology. The relative sparsity of information about C. elegans in its natural context comes from the challenges involved in the identification of the small nematode in nature. Despite these challenges, an increasing focus on the ecology of C. elegans has caused a wealth of new information regarding its life outside of the laboratory. The intensified search for C. elegans in nature has contributed to the discovery of many new Caenorhabditis species and revealed that congeneric nematodes frequently cohabitate in the wild, where they feed on microbial blooms associated with rotting plant material. The identification of new species has also revealed that the androdioecious mating system of males and self-fertilizing hermaphrodites has evolved three times independently within Caenorhabditis. The other two selfing species, C. briggsae and C. tropicalis, share the experimental advantages of C. elegans and have enabled comparative studies into the mechanistic basis of important traits, including self-fertilization. Despite these advances, much remains to be learned about the ecology and natural diversity of these important species. For example, we still lack functional information for many of their genes, which might only be attained through an understanding of their natural ecology. To facilitate ecological research of selfing Caenorhabditis nematodes, we developed a highly scalable method to collect nematodes from the wild. Our method makes use of mobile data collection platforms, cloud-based databases, and the R software environment to enhance researchers&#39; ability to collect nematodes from the wild, record associated ecological data, and identify wild nematodes using molecular barcodes.

A Highly Scalable Approach to Perform Ecological Surveys of Selfing Caenorhabditis Nematodes

This protocol can be used to perform large-scale ecological surveys of selfing Caenorhabditis nematodes. The primary advantage of this method is the efficient organization and analysis of ecological and molecular data associated with the nematodes collected from nature.

Caenorhabditis elegans is one of the major model organisms in biology, but only recently have researchers focused on its natural ecology. The relative ...

Collecting Hemolymph from Small Arthropods

A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods

Arthropods are known to transmit a variety of viruses of medical and agricultural importance through their hemolymph, which is essential for virus transmission. Hemolymph collection is the basic technology for studying virus-vector interactions. Here, we describe a novel and simple method for the quantitative collection of hemolymph from small arthropods using Laodelphax striatellus (the small brown planthopper, SBPH) as a research model, as this arthropod is the main vector of rice stripe virus (RSV). In this protocol, the process begins by gently pinching off one leg of the frozen arthropod with fine-tipped tweezers and pressing the hemolymph out of the wound. Then, a simple micropipette consisting of a capillary and a pipette bulb is used to collect the transudative hemolymph from the wound according to the principle of capillary forces. Finally, the collected hemolymph can be dissolved into a specific buffer for further study. This new method for collecting hemolymph from small arthropods is a useful and efficient tool for further research on arboviruses and vector-virus interactions.

Author Spotlight: A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods  

We describe a method to collect quantifiable hemolymph efficiently from small arthropods for subsequent analysis.

Helminth Collection and Identification from Wildlife

Podsumowanie

Przeglądaj więcej filmów

PLAYLIST