Name: diagnostic necropsy and selected tissue and sample collection in rats and mice
Uploaded: 2011-08-07T13:00:00.000+00:00
Duration: 9 min 13 s
Description: Watch this Scientific Journal Video about Diagnostic Necropsy and Selected Tissue and Sample Collection in Rats and Mice at JoVE.com

1. Preparation for the necropsy
<ol>
 <li>Important parts of any gross postmortem examination (gross necropsy) are the history of the animal and the description of the findings.</li>
 <li>A veterinary pathologist reading your histopathology slides may not have seen the animal and is relying on you for the background information.</li>
 <li>Describe exactly what you see before you euthanize the animal. For example, "female mouse; one of five in the cage; C57BL/6N; head tilted to the right" or "brown male rat; strain unknown; animal is scratching; there is patchy hair loss on the dorsum, near the tail and head". </li>
 <li>Use clear, descriptive, objective language. Modifiers such as "mild", "moderate", and "severe" may be useful if the cutoffs between levels are clearly demarcated. Describing things in terms of food or household objects is generally not recommended.</li>
 <li>Weighing and measuring animals, findings, or organs is often useful. A "large spleen" to you might be a normal spleen to another observer. Saying that the spleen measures 3cm x 1.5cm provides more objective information.</li>
 <li>Photodocumentation can be invaluable.</li>
 <li>Observe all relevant occupational health and personal protective equipment required for working with live animals, potentially infectious carcasses, or chemicals such as formalin.</li>
</ol>
2. Gross postmortem location and examination of heart, lungs, liver, kidneys, and spleen
<ol>
 <li>Gather the necessary supplies prior to euthanasia of the animal. Minimally, this should include a dissection board or similar work surface, forceps, scissors, labels for containers, fixative, and any media or collection tubes/cups that may potentially be needed. </li>
 <li>Briefly assess the condition, behavior, and movement of the animal. Observe and record breathing patterns (e.g., rapid, shallow) as well as ambulatory ability and gait (e.g., limping, circling, tremors). </li>
 <li>Euthanize the animal according to standard procedures at your institution, always adhering to the AVMA guidelines. </li>
 <li>Assess the animal's body condition for skin and coat abnormalities, emaciation, or dehydration. Make note of any artificial manipulations, implants, or surgical scarring. </li>
 <li>Examine all external orifices (ears, eyes, nose, anus, genital openings, and oral cavity). Use of a dissecting scope is recommended for closer observation.</li>
 <li>Lay the euthanized mouse or rat carcass in dorsal recumbency on a clean dissection board or similar work surface.</li>
 <li>Using scissors, incise the skin the full length of the ventrum from the anus to the chin, reflecting the skin and incising the abdominal wall, exposing the abdominal viscera, salivary and preputial/clitoral glands, and cervical and axillary lymph nodes. Cut the rib cage to expose and examine the thoracic viscera by making 2 cuts laterally up each side of the ribcage, then one across, at the top of the sternum, to open a space wide enough to thoroughly examine all the lobes of the lung. </li>
 <li>Examine the appearance of the musculoskeletal structure. </li>
 <li>Evaluate all organs for abnormalities. Specifically find and identify the heart and lungs in the thoracic cavity. Specifically find and identify the liver, kidneys and spleen in the abdominal cavity. Note any color changes, size differences, and missing or mislocated organs. Note the consistency of surfaces, any additional tissue (e.g., masses), fluid pockets, or the presence of fluid in the abdominal/thoracic cavities.</li>
 <li>Observe the gastrointestinal tract for contents, or lack of contents, paying particular attention for thickened walls, masses, and/or hemorrhage. Incise the kidneys (left-longitudinal section, right-cross section, on midline, but off-center) with a scalpel or razor blade to check parenchyma for any abnormality. Check the mesentery for enlarged lymph nodes and/or masses. </li>
 <li>Examine the urogenital system, looking for blockages, fluid pockets, hemorrhage or other abnormalities. </li>
</ol>
3. Postmortem collection of heart, liver, kidneys, and spleen for histopathology
<ol>
 <li>Gather appropriately sized, labeled container(s) filled with a suitable amount of 10% neutral buffered formalin (NBF). Adjust the amount of 10% NBF in order to obtain a 20:1 ratio of fixative to tissue.</li>
 <li>Place the mouse or rat carcass in dorsal recumbency on a clean dissection board or similar work surface and expose the tissue of interest. </li>
 <li>Remove the tissue from the carcass using forceps and scissors.</li>
 <li>Tissue should be closely trimmed to remove fat and unnecessary connective tissue. Tissue should be clean of blood; use normal (or physiologic) saline) to rinse as necessary. Never use distilled or tap water to rinse tissues.</li>
 <li>Place the tissue into the container of 10%NBF.</li>
</ol>
4. Postmortem collection and perfusion of lung tissue
<ol>
 <li>Gather appropriately sized, labeled container(s) filled with a suitable amount of 10% NBF. Adjust amount of 10% NBF in order to obtain a 20:1 ratio of fixative to tissue.</li>
 <li>Place the mouse or rat carcass in dorsal recumbency on a clean dissection board or similar work surface. </li>
 <li>Expose the trachea, heart, and lungs.</li>
 <li>Using scissors and forceps remove the skin and muscle overlying the ventral thoracic and cervical regions. </li>
 <li>Using scissors and forceps, remove the ribcage exposing the heart and lungs by making 2 cuts laterally up each side of the ribcage, then one across near the clavicle to open a space wide enough to thoroughly examine all the lobes of the lung.</li>
 <li>Cut away the neck muscles extending from the sternum and ribs to the jaw, including those overlying the trachea.</li>
 <li>Insert scissors beneath the anterior edge of the rib cage and make 2 cuts, one on either side, to remove the section of bone overlying the trachea.</li>
 <li>Grasp the trachea near the jaw with forceps and cut completely through the trachea with scissors placed above the forceps</li>
 <li>Gently tug the trachea upwards using the forceps, snipping ventral tissue connections with scissors until the entire set of thoracic tissues (trachea, lungs, and heart; this is sometimes called the "pluck") has been removed from the body. </li>
 <li>Lay the lungs flat on the work surface.</li>
 <li>Loosely tie a piece of suture material or kitchen twine around the trachea taking care not to pull tight.</li>
 <li>Fill a syringe with fixative and attach a needle that is small enough to enter the trachea. For mice, a 1ml or 3ml syringe with a 26 gauge needle works well. For rats, a 5ml syringe with an 18 gauge syringe works well. </li>
 <li>Insert the needle into the opening of the trachea and use forceps to hold the trachea surrounding the needle. Begin slowly filling the lungs with fixative.</li>
 <li>Fill the lungs until fully inflated. Do not over- or underinflate. The amount of fixative needed to fully inflate the lungs varies by age, strain, and health of the animal.
 <ol>
 <li>Over-inflation is detected by fluid seeping and foaming from the lung tissue. </li>
 <li>Underinflation is detected by lungs appearing flat and not full in all areas. </li>
 </ol>
 
 </li>
 <li>Remove the needle from the trachea.</li>
 <li>Tighten the suture material or string surrounding the trachea to prevent backflow of fixative out of the lungs.</li>
 <li>Place the inflated lungs into fixative using an approximate 20:1 fixative to tissue ratio.</li>
</ol>
5. Postmortem collection of brain 
<ol>
 <li>Gather appropriately sized, labeled container(s) filled with a suitable amount of 10% NBF. Adjust amount of 10% NBF in order to obtain a 20:1 ratio of fixative to tissue.</li>
 <li>Place the euthanized mouse or rat carcass in ventral recumbency on a clean dissection board or similar work surface. </li>
 <li>Using scissors and forceps, remove the skin and muscle overlying the calvaria.</li>
 <li>Using scissors remove the head completely from the carcass. </li>
 <li>Using small scissors insert the bottom blade into the foramen magnum, the opening where the skull opens into the spinal canal, and keeping scissor tips pointed upwards, begin cutting directly up and through the midline of the calvaria.</li>
 <li>Using forceps, reflect back both halves of the calvaria exposing the brain.
 <ol>
<li>When possible, place the exposed brain into fixative while still in the skull. This will allow the tissue to become firm prior to removal from the skull, if this is required. Many pathologists prefer that sections are cut from the brain while still in the skull.</li>
</ol>
 </li>
 <li>Gently invert the skull so that gravity will help the tissue to fall from the skull. </li>
 <li>Using curved forceps, carefully slide the forceps along the outer edge of the brain and under the brain starting at the olfactory lobes, moving under the cerebrum and towards the cerebellum. Gently pinch with the forceps any connective tissue or nerves that inhibit the brain from falling from the skull. </li>
 <li>Place the brain into fixative using an approximate 20:1 ratio of tissue to fixative. </li>
</ol>
6. Postmortem collection of the mesenteric lymph nodes (MLN)
<ol>
 <li>Tissue collection for PCR analysis should be done using aseptic technique. Flame sterilized, autoclaved or equivalently sterilized instruments should be used.</li>
 <li>Assemble sterile Eppendorf tubes and sterile scissors and forceps.</li>
 <li>Place the euthanized mouse or rat carcass in dorsal recumbency on a clean dissection board or similar work surface. </li>
 <li>Using sterilized scissors and forceps incise the ventral abdominal wall from the genital area to the base of the ribcage, removing both skin and muscle and exposing the intestines.</li>
 <li>The MLN are located in the abdominal cavity in the mesenteric tissue along the colon, immediately adjacent to the cecum. </li>
 <li>To locate the MLN, first locate the cecum which is the large, comma shaped section of the intestine. The colon extends from the cecum and it can often be identified by the presence of fecal pellets. The MLN are located in the mesentery along the colon adjacent to cecum. It can be identified as a yellow, ovoid or spherical small lump of tissue in the white mesenteric tissue and is often slightly thicker and firmer in texture than the surrounding mesentery and fat. Use anatomy text books as necessary for orientation.</li>
 <li>Using aseptic technique and sterile instruments remove the MLN and place into an Eppendorf tube labeled with identifying information. </li>
</ol>
7. Postmortem collection of respiratory aspirate
<ol>
 <li>Assemble needed supplies - sterile pipette, sterile scissors and forceps, sterile solutions to be flushed through the respiratory tract, and a clean dissecting board or similar work surface.</li>
 <li>Lay the euthanized mouse or rat carcass in dorsal recumbency on the dissecting board.</li>
 <li>For bronchial aspirate in rats, access the respiratory tract by way of the trachea. For nasal aspirate in rats, access either through the trachea or through the nasopharyngeal meatus. For bronchial or nasal aspirates in mice, access the respiratory tract through the nasopharyngeal meatus.</li>
 <li>If both nasal and bronchial aspirates are needed, perform the bronchial aspirate first. Perform nasal aspirate using a new sterile pipette.</li>
 <li>Tracheal access (recommended method for rats):
 <ol>
 <li>Reflect the skin away from the cervical area to expose subcutaneous tissues.</li>
 <li>Remove the salivary glands and cervical musculature to expose the trachea.</li>
 <li>Using sterile instruments, incise the trachea to allow access to the lumen. Maintain asepsis throughout collection (Go to step 7.7). </li>
 </ol>
 </li>
 <li>Nasopharyngeal meatus access (nasal aspirate and bronchial aspirate, recommended method for mice due to the smaller size of the trachea):
 <ol>
 <li>This procedure may be performed for both the nasal aspirate and bronchial aspirate. </li>
 <li>Using flame-sterilized or autoclaved instruments, sever the temporomandibular (jaw) joint and reflect the mandible away from the maxilla, exposing the nasopharyngeal meatus. Maintain asepsis throughout collection (Go to step 7.7).</li>
 </ol>
 </li>
 <li>Draw approximately 1ml of sampling fluid into a sterile pipette. This may be normal saline, phosphate buffered saline, or trypticase soy broth. (Go to step 7.8 or 7.9, depending on which aspirate you are collecting.)</li>
 <li>Bronchial aspirate:
 <ol>
<li>Aseptically insert pipette into tracheal lumen, directed caudally, and slowly inject the sampling fluid into the bronchi and the lung. Withdraw the sampling fluid from the bronchi and lungs into the pipette and remove the pipette from the trachea. Not all of the fluid will return into the pipette. Repeat if more fluid is needed for testing.</li>
</ol>
 </li>
 <li>Nasal aspirate:
 <ol>
 <li>Aseptically insert pipette into the nasopharyngeal meatus (mice) or tracheal lumen (rats), directed cranially, and slowly inject the sampling fluid into the nasal cavity.</li>
 <li>Ensure that the nasal cavity is reached by contact of the nasal palate with the pipette tip, or by observation of fluid forced into the cavity, seen as menisci forming at the nasal orifice (nares) or as fluid visible through the translucent oral palate. Fluid should not be seen exiting through the mouth. If so, re-orient the pipette.</li>
 <li>Withdraw the sampling fluid from the nasal cavity into the pipette and remove the pipette from the meatus or trachea.</li>
 </ol>
 </li>
 <li>Aseptically transfer sample to appropriate media or container for testing. </li>
</ol>
8. Representative results
<img src="/files/ftp_upload/2966/2966fig1.jpg" alt="Rodent anatomy diagram; labeled organs include thymus, heart, lungs, liver, intestines." data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1. Abdominal and thoracic organs in the mouse. These organs are typically visible when an animal is first opened (none of the organs have been moved to expose other organs also present in the abdominal cavity). a) thymus, b) heart, c) lungs, d) diaphragm, e) liver, f) small intestines, g) cecum, h) urinary bladder.
<img src="/files/ftp_upload/2966/2966fig2.jpg" alt="Mouse anatomy dissection, labeled organs: liver, intestines, stomach, spleen, kidneys, colon, uterus." data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2. Abdominal and retroperitoneal organs in the mouse. If the intestines and liver are lifted and moved (or removed), these organs can be seen. a) liver (for reference), b) intestines (for reference), c) stomach, d) spleen, e) kidneys, f) descending colon, g) uterus.
<img src="/files/ftp_upload/2966/2966fig3.jpg" alt="Rat anatomy dissection diagram showing seminal vesicles, urinary bladder, and testis for study." data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3. Male reproductive organs. These can be larger in larger, sexually mature males. a) seminal vesicles and coagulating glands, b) testis (pushed into the abdominal cavity from the scrotum through the inguinal rings, which remain open in mice and rats), c) urinary bladder, d) preputial glands, e) epididymis. 
<img src="/files/ftp_upload/2966/2966fig4.jpg" alt="Rat dissection showing uterus, ovary, urinary bladder, clitoral gland; anatomical study diagram." data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4. Female reproductive organs. a) non-gravid uterus (mice and rats have a bicornuate uterus) b) ovary, buried in an ovarian fat pad, c) urinary bladder, d) clitoral glands (analogous to the male preputial glands).

<ol>
	<li>AVMA. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> AVMA Guidelines on Euthanasia. , (2007).</li><li>Feldman, D. B., Seely, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Necropsy Guide: Rodents and the rabbit. , (1988).</li><li>King, J. M., Dodd, D. C., Roth, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Necropsy Book. , (2006).</li><li>Kittel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice--Part+2.+A+joint+publication+of+the+RITA+and+NACAD+groups.">Revised guides for organ sampling and trimming in rats and mice--Part 2. A joint publication of the RITA and NACAD groups.</a> Exp Toxicol Pathol. 55, 413-431 (2004).</li><li>Komarek, V., Fox, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+1.">Chapter 1.</a> The Mouse in Biomedical Research. , 1-22 (2007).</li><li>Morawietz, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice--Part+3.+A+joint+publication+of+the+RITA+and+NACAD+groups.">Revised guides for organ sampling and trimming in rats and mice--Part 3. A joint publication of the RITA and NACAD groups.</a> Exp Toxicol Pathol. 55, 433-449 (2004).</li><li>Popesko, P., Raijtov&#225;, V., Hor&#225;k, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Colour Atlas of Anatomy of Small Laboratory Animals. 2, 91-106 (2002).</li><li>Ruehl-Fehlert, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+guides+for+organ+sampling+and+trimming+in+rats+and+mice--part+1.">Revised guides for organ sampling and trimming in rats and mice--part 1.</a> Exp Toxicol Pathol. 55, 91-106 (2003).</li></ol>

The authors are all employees of Charles River's laboratory rodent diagnostic lab, where these services are offered commercially.

Data collection at the end of a study may require a postmortem examination of animals. Describe well what is seen and remember to examine all tissues. These procedures are designed primarily to optimize necropsy and sample collection for infectious disease surveillance, but most are easily adapted to focused investigations of suspected disease incidents or outbreaks in which perhaps only a subset of the procedures would be employed. Many of the procedures are also relevant to necropsies more commonly performed at the ...

<table><tbody><tr><td>Cutting board</td><td>Thermo Fisher Scientific, Inc.</td><td>Cat #36114</td><td></td></tr><tr><td>Small scissors</td><td>Roboz Surgical Instruments Co.</td><td>RS-5910, G204</td><td>23mm blades, 3.5&quot; length, straight</td></tr><tr><td>Medium scissors</td><td>Roboz Surgical Instruments Co.</td><td>RS-6808, G207</td><td>5&quot;</td></tr><tr><td>Large scissors</td><td>Roboz Surgical Instruments Co.</td><td>RS-6826, G65&quot;</td><td>6.25&quot;</td></tr><tr><td>Forceps-Curved</td><td>Roboz Surgical Instruments Co.</td><td>RS-8254 (M1/21004)</td><td>4.5&quot;, serrated, slight curve</td></tr><tr><td>Forceps-Microdissecting</td><td>Roboz Surgical Instruments Co.</td><td>RS-5238</td><td>Hudson-(EWALD)</td></tr><tr><td>Forceps-Tissue Forceps</td><td>Roboz Surgical Instruments Co.</td><td>RS--8160</td><td>Rat tooth</td></tr><tr><td>Flame sterilizer</td><td>Oxford Labware</td><td>Bacti-Cinerator #5889-001007</td><td></td></tr><tr><td>Scalpel</td><td>Cancer Diagnostics, Inc.</td><td>Finger Scalpels #60, Cat#FS0060</td><td></td></tr><tr><td>Pipettes</td><td>VWR international</td><td>Pasteur Pipet 5 3/4&quot;, Cat#14672-400</td><td></td></tr><tr><td>Autoclave bags</td><td>Propper Manufacturing Company</td><td>Sterilizer Bag Paper Pouches Cat#021002(3100923)E09110</td><td>For Pasteur pipettes</td></tr><tr><td>Pipette bulb</td><td>VWR international</td><td>Cat#56310-240</td><td></td></tr><tr><td>Eppendorf tubes</td><td>Sarstedt Ltd</td><td>SafeSeal microtube 2mL, Ref#72.695</td><td></td></tr><tr><td>Eppendorf tubes</td><td>Argos</td><td>5mL microtube Cat#T20765-C</td><td></td></tr><tr><td>Syringe </td><td>BD Biosciences</td><td>1ml-309602 3 ml-300910 5ml-309603 10mL Cat#309604 20mL Cat#309661</td><td></td></tr><tr><td>Needles</td><td>BD Biosciences</td><td>26G-309625 18G-305195</td><td></td></tr><tr><td>Neutral buffered formalin</td><td>VWR international</td><td>20L 10% NBF, Cat#16004-128</td><td></td></tr><tr><td>Saline</td><td>Thermo Fisher Scientific, Inc.</td><td>Buffered Blood Bank Saline, Cat#23-309-178</td><td></td></tr><tr><td>Trypticase soy broth</td><td>BD Biosciences</td><td>TSB: Dehydrated, Cat#211825</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>Sigma-Aldrich</td><td>HA Buffer, Cat#P3813-10PAK</td><td></td></tr><tr><td>Formalin cups</td><td>VWR international</td><td>4oz Cat#36318-852 8oz Cat#36318-860 16oz Cat#36318-858</td><td></td></tr><tr><td>Large formalin cups </td><td>OakRidge Products</td><td>32oz container Cat#0432-1100</td><td></td></tr><tr><td>Extra large formalin cup</td><td>VWR international</td><td>HDPE Multipurpose 160oz container Cat#89038-282</td><td></td></tr><tr><td>Suture material </td><td>Henry Schein</td><td>Braided silk surgical suture, Ref#100-5000, M766750</td><td>For mouse lung inflation</td></tr><tr><td>Twine</td><td>Staples</td><td>Cat#QUA-46173</td><td>For rat lung inflation</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Diagnostic Necropsy and Selected Tissue and Sample Collection in Rats and Mice at JoVE.com

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

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 ...

diagnostic-necropsy-selected-tissue-sample-collection-rats

Research

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Biology

75.1K Views.  Charles River. 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.

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

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

Mouse Adipose Tissue Collection and Processing for RNA Analysis

Compared to other tissues, white adipose tissue has a considerably less RNA and protein content for downstream applications such as real-time PCR and Western Blot, since it mostly contains lipids. RNA isolation from adipose tissue samples is also challenging as extra steps are required to avoid these lipids. Here, we present a procedure to collect three anatomically different white adipose tissues from mice, to process these samples and perform RNA isolation. We further describe the synthesis of cDNA and gene expression experiments using real-time PCR. The hereby described protocol allows the reduction of contamination from the animal's hair and blood on fat pads as well as cross-contamination between different fat pads during tissue collection. It has also been optimized to ensure adequate quantity and quality of the RNA extracted. This protocol can be widely applied to any mouse model where adipose tissue samples are required for routine experiments such as real-time PCR but is not intended for isolation from primary adipocytes cell culture.

The purpose of this paper is to present a step-by-step procedure to collect different white adipose tissues from mice, process the fat samples and extract RNA.

Compared to other tissues, white adipose tissue has a considerably less RNA and protein content for downstream applications such as real-time PCR and ...

Biochemistry

Immunohistochemical Analysis in the Rat Central Nervous System and Peripheral Lymph Node Tissue Sections

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based multicolor labeling is challenging, especially when utilized for assessing interrelations between target proteins in the tissue with a high fat content such as the central nervous system (CNS).
Our protocol represents a refinement of the standard immunolabeling technique particularly adjusted for detection of both structural and soluble proteins in the rat CNS and peripheral lymph nodes (LN) affected by neuroinflammation. Nonetheless, with or without further modifications, our protocol could likely be used for detection of other related protein targets, even in other organs and species than here presented.

We here present an optimized, detailed protocol for double immunostaining in formalin-fixed, paraffin-embedded rat central nervous system (CNS) and peripheral lymph node (LN) tissue sections.

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based ...

Neuroscience

Metabolomic Analysis of Rat Brain by High Resolution Nuclear Magnetic Resonance Spectroscopy of Tissue Extracts

Studies of gene expression on the RNA and protein levels have long been used to explore biological processes underlying disease. More recently, genomics and proteomics have been complemented by comprehensive quantitative analysis of the metabolite pool present in biological systems. This strategy, termed metabolomics, strives to provide a global characterization of the small-molecule complement involved in metabolism. While the genome and the proteome define the tasks cells can perform, the metabolome is part of the actual phenotype. Among the methods currently used in metabolomics, spectroscopic techniques are of special interest because they allow one to simultaneously analyze a large number of metabolites without prior selection for specific biochemical pathways, thus enabling a broad unbiased approach. Here, an optimized experimental protocol for metabolomic analysis by high-resolution NMR spectroscopy is presented, which is the method of choice for efficient quantification of tissue metabolites. Important strengths of this method are (i) the use of crude extracts, without the need to purify the sample and/or separate metabolites; (ii) the intrinsically quantitative nature of NMR, permitting quantitation of all metabolites represented by an NMR spectrum with one reference compound only; and (iii) the nondestructive nature of NMR enabling repeated use of the same sample for multiple measurements. The dynamic range of metabolite concentrations that can be covered is considerable due to the linear response of NMR signals, although metabolites occurring at extremely low concentrations may be difficult to detect. For the least abundant compounds, the highly sensitive mass spectrometry method may be advantageous although this technique requires more intricate sample preparation and quantification procedures than NMR spectroscopy. We present here an NMR protocol adjusted to rat brain analysis; however, the same protocol can be applied to other tissues with minor modifications.

The neurochemistry of mammalian brain is changed in many neurological and systemic diseases. Characteristic profiles of cerebral metabolites can be efficiently obtained based on crude extracts of brain tissue. To this end, high-resolution NMR spectroscopy is employed, enabling detailed quantitative analysis of metabolite concentrations (metabolomics).

Studies of gene expression on the RNA and protein levels have long been used to explore biological processes underlying disease. More recently, genomics ...

Using Fluorescence Activated Cell Sorting to Examine Cell-Type-Specific Gene Expression in Rat Brain Tissue

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant cell type in the brain, neurons are the most widely studied of these cell types given their direct role in impacting behaviors. Other cell types in the brain also impact neuronal function and behavior via the signaling molecules they produce. Neuroscientists must understand the interactions between the cell types in the brain to better understand how these interactions impact neural function and disease. To date, the most common method of analyzing protein or gene expression utilizes the homogenization of whole tissue samples, usually with blood, and without regard for cell type. This approach is an informative approach for examining general changes in gene or protein expression that may influence neural function and behavior; however, this method of analysis does not lend itself to a greater understanding of cell-type-specific gene expression and the effect of cell-to-cell communication on neural function. Analysis of behavioral epigenetics has been an area of growing focus which examines how modifications of the deoxyribonucleic acid (DNA) structure impact long-term gene expression and behavior; however, this information may only be relevant if analyzed in a cell-type-specific manner given the differential lineage and thus epigenetic markers that may be present on certain genes of individual neural cell types. The Fluorescence Activated Cell Sorting (FACS) technique described below provides a simple and effective way to isolate individual neural cells for the subsequent analysis of gene expression, protein expression, or epigenetic modifications of DNA. This technique can also be modified to isolate more specific neural cell types in the brain for subsequent cell-type-specific analysis.

The goal of this protocol is to use the fluorescence activated cell sorting (FACS) technique to sort specific types of neural cells for subsequent analysis of cell-type-specific gene expression, epigenetic markers, and or protein expression.

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant ...

Rabies Necropsy Techniques in Large and Small Animals

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the entire state, with the exception of New York City. The Rabies laboratory necropsies a variety of animals ranging in size from bats to bovids. Most of these specimens are animals exhibiting neurological signs, however, less than 10% actually test positive for rabies; implying trauma, lesions or other infectious agents as the cause of these symptoms. Due to the risk of aerosolizing undiagnosed infectious agents, the Rabies Laboratory does not use power tools or saws. Three necropsy techniques will be presented for animals whose skulls are impenetrable with scissors. The laboratory has implemented these techniques to decrease potential exposure to infectious agents, eliminate unnecessary manipulation of the specimen and reduce processing time. The advantages of a preferred technique opposed to another is subject to the trained individual processing the specimen.

The goal of this protocol is to demonstrate safe necropsy techniques in small and large animals to obtain satisfactory tissue samples for rabies testing.

The New York State Department of Health (NYSDOH) Rabies Laboratory receives between 6,000 to 9,000 specimens annually and performs rabies testing for the ...

Standardized 5/6 Nephrectomy in Rats for Renal Insufficiency and Malperfusion

5/6 Nephrectomy Using Sharp Bipolectomy Via Midline Laparotomy in Rats

Chronic kidney disease (CKD) affects over 10% of the global population, amounting to more than 800 million individuals worldwide. The advances in the treatment of CKD have had a significant impact on patient outcomes. While in the past, CKD was often considered a death sentence, with many patients succumbing to the complications of end-stage renal disease, it is now increasingly being managed as a chronic condition with the availability of dialysis and kidney transplantation, as well as new pharmaceutical developments such as SGLT2 inhibitors or nonsteroidal mineralocorticoid receptor antagonists. 
Yet, there continues to be a growing demand for further exploration into the pathophysiological processes and potential therapeutic interventions. Reliable biological models play a crucial role in facilitating this research. Given the multifaceted nature of renal disease, which encompasses not only cell biology but also vascular microanatomy and endocrine signaling, an appropriate model must possess a level of biological complexity that only an animal model can offer, rendering rodents an obvious choice.
This manuscript, therefore, provides an intricate, systematic protocol for surgically reducing renal parenchyma through midline laparotomy and combined total and partial nephrectomy in rats for survival as well as non-survival applications. It emphasizes the critical role of precise surgical techniques in ensuring consistent and reliable outcomes. Prime examples of potential applications for this model include biomolecular and pharmaceutical studies as well as the development of innovative intraoperative imaging modalities, such as hyperspectral imaging, to objectively visualize and differentiate renal malperfusion.

This manuscript introduces a standardized protocol for a 5/6 nephrectomy by sharp bipolectomy via midline laparotomy in a rat model, aiming to induce a state of renal insufficiency via renal parenchyma reduction with a great degree of methodical precision and low risk of technical error.

Chronic kidney disease (CKD) affects over 10% of the global population, amounting to more than 800 million individuals worldwide. The advances in the ...

Medicine

Diagnostic Necropsy and Tissue Harvest

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
Many animal experiments rely on final data collection time points that are gathered from the harvesting and testing of organs and tissues. The use of appropriate methods for the collection of organs and tissues can impact the quality of the samples and the analysis of the data that is gleaned for the testing of the tissues. The method of euthanasia of the animal can also impact the quality of the samples. This manuscript will outline proper necropsy techniques for rats.

Diagnostic Necropsy and Tissue Harvest in Rodents

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
Many animal experiments rely on final data ...

Education

JoVE Science Education

Basic Biology

Lab Animal Research

Sterile Tissue Harvest

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
In 1959 The 3 R&#39;s were introduced by W.M.S. Russell and R.L. Burch in their book The Principles of Humane Experimental Technique. The 3 R&#39;s are replacement, reduction, and refinement of the use of animals in research.1 The use of cell lines and tissue cultures that originated from research animals is a replacement technique, as it allows for many experiments to be conducted in vitro. Harvesting tissues and organs for use in cell and tissue cultures requires aseptic technique to avoid contamination of the tissues. Sterile harvest is also necessary for protein and RNA analysis and metabolic profiling of tissues. This manuscript will discuss the process of sterile organ harvest in rats and mice.

Sterile Tissue Harvest in Rodents

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN
In 1959 The 3 R&#39;s were introduced by W.M.S. ...

A Focus-Forming Assay for Quantifying Virus Titers in the Organs of an Infected Mouse

Source: Brien, J. D. et al., <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/59632">Isolation and Quantification of Zika Virus from Multiple Organs in a Mouse.</a> J. Vis. Exp.&#160;(2019)
The video demonstrates a procedure for quantifying the Zika virus in multiple mouse organs via a focus-forming assay. The process involves infecting mammalian cell monolayers with virus-infected tissue homogenates, followed by the use of methyl cellulose to prevent viral spread. Subsequently, immunostaining is employed to enable precise quantification and measurement of organ-specific infectivity.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Organ ...