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>

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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 border="0">
	<tbody>
		<tr>
			<td colspan="4">Fixing</td>
		</tr>
		<tr>
			<td>Alcohol-Formalin-Acetic Acid (AFA)</td>
			<td>Fisher Scientific</td>
			<td>A407-1 (ethanol) 
			F75F-1GAL (formaldehyde) 
			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&#160;</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) 
			AC15365-1000 
			(glycerine)</td>
			<td>Ethanol 70% (95 ml) + glycerine (5 ml)</td>
		</tr>
		<tr>
			<td colspan="4">Clearing, staining and mounting</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) 
			A407-1 (ethanol)</td>
			<td>Phenol 100% (80 ml) + ethanol 100% (20 ml)</td>
		</tr>
		<tr>
			<td>Harris&#8217; Hematoxylin</td>
			<td>Fisher Scientific</td>
			<td>S25347 (hematoxylin) 
			A407-1 (ethanol) 
			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)&#160;&#160;</td>
		</tr>
		<tr>
			<td>Semichon&#8217;s acetic carmine</td>
			<td>Fisher Scientific</td>
			<td>BP2401500 (glacial acetic acid) 
			S25236 (carmine) 
			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) 
			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) 
			SA9233-100 (hydrochloric acid)</td>
			<td>Ethanol 70% (99.5 ml) +&#160; 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), ...

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15.4K visualizações.  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. 

Vídeo: Helminth Collection and Identification from Wildlife

Using Pharmacological Manipulation and High-precision Radio Telemetry to Study the Spatial Cognition in Free-ranging Animals

An animal&#39;s ability to perceive and learn about its environment plays a key role in many behavioral processes, including navigation, migration, dispersal and foraging. However, the understanding of the role of cognition in the development of navigation strategies and the mechanisms underlying these strategies is limited by the methodological difficulties involved in monitoring, manipulating the cognition of, and tracking wild animals. This study describes a protocol for addressing the role of cognition in navigation that combines pharmacological manipulation of behavior with high-precision radio telemetry. The approach uses scopolamine, a muscarinic acetylcholine receptor antagonist, to manipulate cognitive spatial abilities. Treated animals are then monitored with high frequency and high spatial resolution via remote triangulation. This protocol was applied within a population of Eastern painted turtles (Chrysemys picta) that has inhabited seasonally ephemeral water sources for ~100 years, moving between far-off sources using precise (&#177; 3.5 m), complex (i.e., non-linear with high tortuosity that traverse multiple habitats), and predictable routes learned before 4 years of age. This study showed that the processes used by these turtles are consistent with spatial memory formation and recall. Together, these results are consistent with a role of spatial cognition in complex navigation and highlight the integration of ecological and pharmacological techniques in the study of cognition and navigation.

This paper describes a novel protocol that combines the pharmacological manipulation of memory and radio telemetry to document and quantify the role of cognition in navigation.

An animal&#39;s ability to perceive and learn about its environment plays a key role in many behavioral processes, including navigation, migration, ...

Automated, High-resolution Mobile Collection System for the Nitrogen Isotopic Analysis of NOx

Nitrogen oxides (NOx = NO + NO2) are a family of atmospheric trace gases that have great impact on the environment. NOx concentrations directly influence the oxidizing capacity of the atmosphere through interactions with ozone and hydroxyl radicals. The main sink of NOx is the formation and deposition of nitric acid, a component of acid rain and a bioavailable nutrient. NOx is emitted from a mixture of natural and anthropogenic sources, which vary in space and time. The collocation of multiple sources and the short lifetime of NOx make it challenging to quantitatively constrain the influence of different emission sources and their impacts on the environment. Nitrogen isotopes of NOx have been suggested to vary amongst different sources, representing a potentially powerful tool to understand the sources and transport of NOx. However, previous methods of collecting atmospheric NOx integrate over long (week to month) time spans and are not validated for the efficient collection of NOx in relevant, diverse field conditions. We report on a new, highly efficient field-based system that collects atmospheric NOx for isotope analysis at a time resolution between 30 min and 2 hr. This method collects gaseous NOx in solution as nitrate with 100% efficiency under a variety of conditions. Protocols are presented for collecting air in urban settings under both stationary and mobile conditions. We detail the advantages and limitations of the method and demonstrate its application in the field. Data from several deployments are shown to 1) evaluate field-based collection efficiency by comparisons with in situ NOx concentration measurements, 2) test the stability of stored solutions before processing, 3) quantify in situ reproducibility in a variety of urban settings, and 4) demonstrate the range of N isotopes of NOx detected in ambient urban air and on heavily traveled roadways.

Automated, High-resolution Mobile Collection System for the Nitrogen Isotopic Analysis of NOx

Previous work suggests that the nitrogen isotopic composition of atmospheric nitrogen oxides might distinguish the influence of different sources in the environment. We report on an automated, mobile, field-based method for the high collection efficiency of atmospheric NOx for N isotopic analysis at an hourly time resolution.

Nitrogen oxides (NOx = NO + NO2) are a family of atmospheric trace gases that have great impact on the environment. NOx concentrations directly influence ...

Protocol for Assessing the Relative Effects of Environment and Genetics on Antler and Body Growth for a Long-lived Cervid

Cervid phenotype can be placed into one of two categories: efficiency, which promotes survival over extravagant morphometric growth, and luxury, which promotes growth of large weaponry and body size. Populations of the same species display each phenotype depending on environmental conditions. Although antler and body size of male white-tailed deer (Odocoileus virginianus) varies by physiographic region in Mississippi, USA and is strongly correlated with regional variation in nutritional quality, the effects of population-level genetics from native stocks and previous re-stocking efforts cannot be disregarded. This protocol describes how we designed a controlled study, where other factors that influence phenotype, such as age and nutrition, are controlled. We brought wild-caught pregnant females and six-month-old fawns from three distinct physiographic regions in Mississippi, USA to the Mississippi State University Rusty Dawkins Memorial Deer Unit. Deer from the same region were bred to produce a second generation of offspring, allowing us to assess generational responses and maternal effects. All deer ate the same high-quality (20% crude protein deer pellet) diet ad libitum. We uniquely marked each neonate and recorded body mass, hind foot, and total body length. Each subsequent fall, we sedated individuals via remote injection and sampled the same morphometrics plus antlers of adults. We found that all morphometrics increased in size from first to second generation, with full compensation of antler size (regional variation no longer present) and partial compensation of body mass (some evidence of regional variation) evident in the second generation. Second generation males that originated from our poorest quality soil region displayed about a 40% increase in antler size and about a 25% increase in body mass when compared to their wild harvested counterparts. Our results suggest phenotypic variation of wild male white-tailed deer in Mississippi are more related to differences in nutritional quality than population-level genetics.

Phenotypic differences among cervid populations may be related to population-level genetics or nutrition; discerning which is difficult in the wild. This protocol describes how we designed a controlled study where nutritional variation was eliminated. We found that phenotypic variation of male white-tailed deer was more limited by nutrition than genetics.

Cervid phenotype can be placed into one of two categories: efficiency, which promotes survival over extravagant morphometric growth, and luxury, which ...

Collection and Extraction of Occupational Air Samples for Analysis of Fungal DNA

Traditional methods of identifying fungal exposures in occupational environments, such as culture and microscopy-based approaches, have several limitations that have resulted in the exclusion of many species. Advances in the field over the last two decades have led occupational health researchers to turn to molecular-based approaches for identifying fungal hazards. These methods have resulted in the detection of many species within indoor and occupational environments that have not been detected using traditional methods. This protocol details an approach for determining fungal diversity within air samples through genomic DNA extraction, amplification, sequencing, and taxonomic identification of fungal internal transcribed spacer (ITS) regions. ITS sequencing results in the detection of many fungal species that are either not detected or difficult to identify to species level using culture or microscopy. While these methods do not provide quantitative measures of fungal burden, they offer a new approach to hazard identification and can be used to determine overall species richness and diversity within an occupational environment.

Determining the fungal diversity within an environment is a method utilized in occupational health studies to identify health hazards. This protocol describes DNA extraction from occupational air samples for amplification and sequencing of fungal ITS regions. This approach detects many fungal species that can be overlooked by traditional assessment methods.

Traditional methods of identifying fungal exposures in occupational environments, such as culture and microscopy-based approaches, have several ...

A Video Surveillance System to Monitor Breeding Colonies of Common Terns (Sterna Hirundo)

Many waterbird populations have faced declines over the last century, including the common tern (Sterna hirundo), a waterbird species with a widespread breeding distribution, that has been recently listed as endangered in some habitats of its range. Waterbird monitoring programs exist to track populations through time; however, some of the more intensive approaches require entering colonies and can be disruptive to nesting populations. This paper describes a protocol that utilizes a minimally invasive surveillance system to continuously monitor common tern nesting behavior in typical ground-nesting colonies. The video monitoring system utilizes wireless cameras focused on individual nests as well as over the colony as a whole, and allows for observation without entering the colony. The video system is powered with several 12 V car batteries that are continuously recharged using solar panels. Footage is recorded using a digital video recorder (DVR) connected to a hard drive, which can be replaced when full. The DVR may be placed outside of the colony to reduce disturbance. In this study, 3,624 h of footage recorded over 63 days in weather conditions ranging from 12.8 &#176;C to 35.0 &#176;C produced 3,006 h (83%) of usable behavioral data. The types of data retrieved from the recorded video can vary; we used it to detect external disturbances and measure nesting behavior during incubation. Although the protocol detailed here was designed for ground-nesting waterbirds, the principal system could easily be modified to accommodate alternative scenarios, such as colonial arboreal nesting species, making it widely applicable to a variety of research needs.

A Video Surveillance System to Monitor Breeding Colonies of Common Terns Sterna Hirundo

This paper describes a protocol that uses a remote video monitoring surveillance system to continuously monitor breeding colonies of ground-nesting waterbirds. The system includes five cameras monitoring individual nests and one camera monitoring the colony as a whole, and is powered by car batteries that are recharged via solar panels.

Many waterbird populations have faced declines over the last century, including the common tern (Sterna hirundo), a waterbird species with a widespread ...

Detection of Viruses from Bioaerosols Using Anion Exchange Resin

This protocol demonstrates a customized bioaerosol sampling method for viruses. In this system, anion exchange resin is coupled with liquid impingement-based air sampling devices for efficacious concentration of negatively-charged viruses from bioaerosols. Thus, the resin serves as an additional concentration step in the bioaerosol sampling workflow. Nucleic acid extraction of the viral particles is then performed directly from the anion exchange resin, with the resulting sample suitable for molecular analyses. Further, this protocol describes a custom-built bioaerosol chamber capable of generating virus-laden bioaerosols under a variety of environmental conditions and allowing for continuous monitoring of environmental variables such as temperature, humidity, wind speed, and aerosol mass concentration. The main advantage of using this protocol is increased sensitivity of viral detection, as assessed via direct comparison to an unmodified conventional liquid impinger. Other advantages include the potential to concentrate diverse negatively-charged viruses, the low cost of anion exchange resin (~$0.14 per sample), and ease of use. Disadvantages include the inability of this protocol to assess infectivity of resin-adsorbed viral particles, and potentially the need for the optimization of the liquid sampling buffer used within the impinger.

An anion exchange resin-based method, adapted to liquid impingement-based bioaerosol sampling of viruses is demonstrated. When coupled with downstream molecular detection, the method allows for facile and sensitive detection of viruses from bioaerosols.

This protocol demonstrates a customized bioaerosol sampling method for viruses. In this system, anion exchange resin is coupled with liquid ...

A Loop-mediated Isothermal Amplification (LAMP) Assay for Rapid Identification of Bemisia tabaci

The whitefly Bemisia tabaci (Gennadius) is an invasive pest of considerable importance, affecting the production of vegetable and ornamental crops in many countries around the world. Severe yield losses are caused by direct feeding, and even more importantly, also by the transmission of more than 100 harmful plant pathogenic viruses. As for other invasive pests, increased international trade facilitates the dispersal of B. tabaci to areas beyond its native range. Inspections of plant import products at points of entry such as seaports and airports are, therefore, seen as an important prevention measure. However, this last line of defense against pest invasions is only effective if rapid identification methods for suspicious insect specimens are readily available. Because the morphological differentiation between the regulated B. tabaci and close relatives without quarantine status is difficult for non-taxonomists, a rapid molecular identification assay based on the loop-mediated isothermal amplification (LAMP) technology has been developed. This publication reports the detailed protocol of the novel assay describing rapid DNA extraction, set-up of the LAMP reaction, as well as interpretation of its read-out, which allows identifying B. tabaci specimens within one hour. Compared to existing protocols for the detection of specific B. tabaci biotypes, the developed method targets the whole B. tabaci species complex in one assay. Moreover the assay is designed to be applied on-site by plant health inspectors with minimal laboratory training directly at points of entry. Thorough validation performed under laboratory and on-site conditions demonstrates that the reported LAMP assay is a rapid and reliable identification tool, improving the management of B. tabaci.

A Loop-mediated Isothermal Amplification LAMP Assay for Rapid Identification of Bemisia tabaci

This paper reports the protocol for a rapid identification assay for Bemisia tabaci based on loop-mediated isothermal amplification (LAMP) technology. The protocol requires minimal laboratory training and can, therefore, be implemented on-site at points of entry for plant imports such as seaports and airports.

The whitefly Bemisia tabaci (Gennadius) is an invasive pest of considerable importance, affecting the production of vegetable and ornamental crops in many ...

Data Collection on Marine Litter Ingestion in Sea Turtles and Thresholds for Good Environmental Status

The following protocol is intended to respond to the requirements set by the European Union&#39;s Marine Strategy Framework Directives (MSFD) for the D10C3 Criteria reported in the Commission Decision (EU), related to the amount of litter ingested by marine animals. Standardized methodologies for extracting litter items ingested from dead sea turtles along with guidelines on data analysis are provided. The protocol starts with the collection of dead sea turtles and classification of samples according to the decomposition status. Turtle necropsy must be performed in authorized centers and the protocol described here explains the best procedure for gastrointestinal (GI) tract isolation. The three parts of the GI (esophagus, stomach, intestine) should be separated, opened lengthways and contents filtered using a 1 mm mesh sieve. The article describes the classification and quantification of ingested litter, classifying GI contents into seven different categories of marine litter and two categories of natural remains. The quantity of ingested litter should be reported as total dry mass (weight in grams, with two decimal places) and abundance (number of items). The protocol proposes two possible scenarios to achieve the Good Environmental Status (GES). First: &#34;There should be less than X% of sea turtles having Y g or more plastic in the GI in samples of 50-100 dead turtles from each sub-region&#34;, where Y is the average weight of plastic ingested and X% is the percentage of sea turtles with more weight (in grams) of plastic than Y. The second one, which considers the food remain versus plastic as a proxy of individual health, is: &#34;There should be less than X% of sea turtles having more weight of plastic (in grams) than food remains in the GI in samples of 50-100 dead turtles from each sub-region&#34;.

The protocol focuses on the collection of sea turtle samples, describing all the steps from the animal recovery and necropsy to the classification and quantification of ingested marine litter. Moreover, the representative results show how to use the collected data to elaborate the possible thresholds for Good Environmental Status.

The following protocol is intended to respond to the requirements set by the European Union&#39;s Marine Strategy Framework Directives (MSFD) for the ...

A Method for Quantifying Foliage-Dwelling Arthropods

Terrestrial arthropods play an important role in our environment. Quantifying arthropods in a way that allows for a precise index or estimates of density requires a method with high detection probability and a known sampling area. While most described methods provide a qualitative or semi-quantitative estimate adequate for describing species presence, richness, and diversity, few provide an adequately consistent detection probability and known or consistent sampling areas to provide an index or estimate with adequate precision to detect differences in abundance across environmental, spatial, or temporal variables. We describe how to quantify leaf-dwelling arthropods by sealing the leaves and end of branches in a bag, clipping and freezing the bagged material, and rinsing the previously frozen material in water to separate arthropods from the substrate and quantify them. As we demonstrate, this method can be used at a landscape scale to quantify leaf-dwelling arthropods with adequate precision to test for and describe how spatial, temporal, environmental, and ecological variables influence arthropod richness and abundance. This method allowed us to detect differences in density, richness, and diversity of leaf-dwelling arthropods among 5 genera of trees commonly found in southeastern deciduous forests.

We describe how to quantify leaf dwelling arthropods by sealing the leaves and end of branches in a bag, clipping and freezing the bagged material, and rinsing the previously frozen material in water to separate arthropods from the substrate for quantification.

Terrestrial arthropods play an important role in our environment. Quantifying arthropods in a way that allows for a precise index or estimates of density ...

Quantifying Corticolous Arthropods Using Sticky Traps

Terrestrial arthropods play an important role in our environment. Quantifying arthropods in a way that allows for a precise index or estimate of density requires a method with high detection probability and a consistent sampling area. We used manufactured sticky traps to compare abundance, total length (a surrogate for biomass), richness, and Shannon diversity of corticolous arthropods among the boles of 5 tree species. Efficacy of this method was adequate to detect variation in corticolous arthropods among tree species and provide a standard error of the mean that was &#60;20% of the mean for all estimates with sample sizes from 7 to 15 individual trees of each species. Our results indicate, even with these moderate sample sizes, the level of precision of arthropod community metrics produced with this approach is adequate to address most ecological questions regarding temporal and spatial variation in corticolous arthropods. Results from this method differ from other quantitative approaches such as chemical knockdown, visual inspection, and funnel traps in that they provide an indication of corticolous arthropod activity over a relatively long-term, better including temporary bole residents, flying arthropods that temporarily land on the tree bole and crawling arthropods that use the tree bole as a travel route from the ground to higher forest foliage. Furthermore, we believe that commercially manufactured sticky traps provide more precise estimates and are logistically simpler than the previously described method of directly applying a sticky material to tree bark or applying a sticky material to tape or other type of backing and applying that to the tree bark.

We describe a semi-quantitative approach of measuring characteristics of corticolous (bark-dwelling) arthropod communities. We placed commercially manufactured sticky traps on tree boles to estimate abundance, total length (a surrogate to biomass), richness, and Shannon diversity for comparison among tree species.

Terrestrial arthropods play an important role in our environment. Quantifying arthropods in a way that allows for a precise index or estimate of density ...