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="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="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="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="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 border="1">
		<tbody>
				<tr>
						<th>Reagent name</th>
						<th>Company</th>
						<th>Catalog number</th>
						<th>Comments</th>
				</tr>
				<tr>
						<td>Cutting board</td>
						<td>Thermo Electron Corp	Cat #36114</td>
						<td>Cat #36114</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Small scissors</td>
						<td>ROBOZ</td>
						<td>RS-5910, G204</td>
						<td>23mm blades, 3.5" length, straight</td>
				</tr>
				<tr>
						<td>Medium scissors	</td>
						<td>ROBOZ</td>
						<td>RS-6808, G207</td>
						<td>5"</td>
				</tr>
				<tr>
						<td>Large scissors</td>
						<td>ROBOZ</td>
						<td>RS-6826, G65"</td>
						<td>6.25"</td>
				</tr>
				<tr>
						<td>Forceps-Curved</td>
						<td>ROBOZ</td>
						<td>RS-8254 (M1/21004)</td>
						<td>4.5", serrated, slight curve</td>
				</tr>
				<tr>
						<td>Forceps-Microdissecting</td>
						<td>ROBOZ</td>
						<td>RS-5238</td>
						<td>Hudson-(EWALD)</td>
				</tr>
				<tr>
						<td>Forceps-Tissue Forceps</td>
						<td>ROBOZ</td>
						<td>RS--8160</td>
						<td>Rat tooth</td>
				</tr>
				<tr>
						<td>Flame sterilizer</td>
						<td>Oxford Labware Products</td>
						<td>Bacti-Cinerator #5889-001007</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Scalpel</td>
						<td>Cancer Diagnostics, Inc.</td>
						<td>Finger Scalpels #60, Cat#FS0060</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Pipettes</td>
						<td>VWR</td>
						<td>Pasteur Pipet 5 3/4", Cat#14672-400</td>
						<td>&nbsp;</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</td>
						<td>Cat#56310-240</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Eppendorf tubes	</td>
						<td>Sarstedt</td>
						<td>SafeSeal microtube 2mL, Ref#72.695</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Eppendorf tubes	</td>
						<td>Argos</td>
						<td>5mL microtube Cat#T20765-C</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Syringe </td>
						<td>BD (Beckton Dickenson)</td>
						<td>1ml-309602 3 ml-300910 5ml-309603 10mL Cat#309604 20mL Cat#309661</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Needles</td>
						<td>BD</td>
						<td>26G-309625 18G-305195 </td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Neutral buffered formalin</td>
						<td>VWR</td>
						<td>20L 10% NBF, Cat#16004-128</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Saline</td>
						<td>ThermoScientific</td>
						<td>Buffered Blood Bank Saline, Cat#23-309-178</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Trypticase soy broth</td>
						<td>Becton Dickenson</td>
						<td>TSB: Dehydrated, Cat#211825</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Phosphate buffered saline</td>
						<td>Sigma Aldrich</td>
						<td>HA Buffer, Cat#P3813-10PAK</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Formalin cups</td>
						<td>VWR</td>
						<td>4oz Cat#36318-852 8oz Cat#36318-860 16oz Cat#36318-858</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Large formalin cups </td>
						<td>OakRidge Products</td>
						<td>32oz container Cat#0432-1100</td>
						<td>&nbsp;</td>
				</tr>
				<tr>
						<td>Extra large formalin cup</td>
						<td>VWR</td>
						<td>HDPE Multipurpose 160oz container Cat#89038-282</td>
						<td>&nbsp;</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 ...

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74.2K 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

Using Micro-Electro-Mechanical Systems (MEMS) to Develop Diagnostic Tools

Using Micro-Electro-Mechanical Systems MEMS to Develop Diagnostic Tools

Manual Restraint and Common Compound Administration Routes in Mice and Rats

Being able to safely and effectively restrain mice and rats is an important part of conducting research. Working confidently and humanely with mice and rats requires a basic competency in handling and restraint methods. This article will present the basic principles required to safely handle animals. One-handed, two-handed, and restraint with specially designed restraint objects will be illustrated. Often, another part of the research or testing use of animals is the effective administration of compounds to mice and rats. Although there are a large number of possible administration routes (limited only by the size and organs of the animal), most are not used regularly in research. This video will illustrate several of the more common routes, including intravenous, intramuscular, subcutaneous, and oral gavage. The goal of this article is to expose a viewer unfamiliar with these techniques to basic restraint and substance administration routes. This video does not replace required hands-on training at your facility, but is meant to augment and supplement that training.

Working safely and humanely with research rodents requires a core competency in handling and restraint methods. This article will present the basic principles required to safely handle and effectively administer compounds to mice and rats.

Being able to safely and effectively restrain mice and rats is an important part of conducting research.  Working confidently and humanely with mice and ...

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies showed that microinjection of germ cells from fertile donor mice into the seminiferous tubules of infertile recipient mice results in donor-derived spermatogenesis and sperm production by the recipient animal2. The use of donor males carrying the bacterial &beta;-galactosidase gene allowed identification of donor-derived spermatogenesis and transmission of the donor haplotype to the offspring by recipient animals1. Surprisingly, after transplantation into the lumen of the seminiferous tubules, transplanted germ cells were able to move from the luminal compartment to the basement membrane where spermatogonia are located3. It is generally accepted that only SSCs are able to colonize the niche and re-establish spermatogenesis in the recipient testis. Therefore, germ cell transplantation provides a functional approach to study the stem cell niche in the testis and to characterize putative spermatogonial stem cells. To date, germ cell transplantation is used to elucidate basic stem cell biology, to produce transgenic animals through genetic manipulation of germ cells prior to transplantation4,5, to study Sertoli cell-germ cell interaction6,7, SSC homing and colonization3,8, as well as SSC self-renewal and differentiation9,10.
Germ cell transplantation is also feasible in large species11. In these, the main applications are preservation of fertility, dissemination of elite genetics in animal populations, and generation of transgenic animals as the study of spermatogenesis and SSC biology with this technique is logistically more difficult and expensive than in rodents. Transplantation of germ cells from large species into the seminiferous tubules of mice results in colonization of donor cells and spermatogonial expansion, but not in their full differentiation presumably due to incompatibility of the recipient somatic cell compartment with the germ cells from phylogenetically distant species12. An alternative approach is transplantation of germ cells from large species together with their surrounding somatic compartment. We first reported in 2002, that small fragments of testis tissue from immature males transplanted under the dorsal skin of immunodeficient mice are able to survive and undergo full development with the production of fertilization competent sperm13. Since then testis tissue xenografting has been shown to be successful in many species and emerged as a valuable alternative to study testis development and spermatogenesis of large animals in mice14.

Protocols for germ cell transplantation and testis tissue xenografting are described. Germ cell transplantation results in donor-derived spermatogenesis in recipient testes and represents a functional reconstitution assay for identification of spermatogonial stem cells (SSCs). Testis tissue xenografting reproduces testis development and spermatogenesis of various donor species in recipient mice.

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies ...

Ovariectomy and 17&beta;-estradiol Replacement in Rats and Mice: A Visual Demonstration

Estrogens are a family of female sexual hormones with an exceptionally wide spectrum of effects. When rats and mice are used in estrogen research they are commonly ovariectomized in order to ablate the rapidly cycling hormone production, replacing the 17&beta;-estradiol exogenously. There is, however, lack of consensus regarding how the hormone should be administered to obtain physiological serum concentrations. This is crucial since the 17&beta;-estradiol level/administration method profoundly influences the experimental results1-3. We have in a series of studies characterized the different modes of 17&beta;-estradiol administration, finding that subcutaneous silastic capsules and per-oral nut-cream Nutella are superior to commercially available slow-release pellets (produced by the company Innovative Research of America) and daily injections in terms of producing physiological serum concentrations of 17&beta;-estradiol4-6. Amongst the advantages of the nut-cream method, that previously has been used for buprenorphine administration7, is that when used for estrogen administration it resembles peroral hormone replacement therapy and is non-invasive. The subcutaneous silastic capsules are convenient and produce the most stable serum concentrations. This video article contains step-by-step demonstrations of ovariectomy and 17&beta;-estradiol hormone replacement by silastic capsules and peroral Nutella in rats and mice, followed by a discussion of important aspects of the administration procedures.

Ovariectomy and 17&amp;#946;-estradiol Replacement in Rats and Mice: A Visual Demonstration

Ovariectomy and subsequent replacement of 17&beta;-estradiol by means of silastic capsules and peroral Nutella are demonstrated in rats and mice.

Estrogens are a family of female sexual hormones with an exceptionally wide spectrum of effects. When rats and mice are used in estrogen research they are ...

Proteomic Sample Preparation from Formalin Fixed and Paraffin Embedded Tissue

Preserved clinical material is a unique source for proteomic investigation of human disorders. Here we describe an optimized protocol allowing large scale quantitative analysis of formalin fixed and paraffin embedded (FFPE) tissue. The procedure comprises four distinct steps. The first one is the preparation of sections from the FFPE material and microdissection of cells of interest. In the second step the isolated cells are lysed and processed using 'filter aided sample preparation' (FASP) technique. In this step, proteins are depleted from reagents used for the sample lysis and are digested in two-steps using endoproteinase LysC and trypsin. 
After each digestion, the peptides are collected in separate fractions and their content is determined using a highly sensitive fluorescence measurement. Finally, the peptides are fractionated on 'pipette-tip' microcolumns. The LysC-peptides are separated into 4 fractions whereas the tryptic peptides are separated into 2 fractions. In this way prepared samples allow analysis of proteomes from minute amounts of material to a depth of 10,000 proteins. Thus, the described workflow is a powerful technique for studying diseases in a system-wide-fashion as well as for identification of potential biomarkers and drug targets.

Archival formalin fixed and paraffin embedded (FFPE) clinical samples are valuable material for investigation of diseases. Here we demonstrate a sample preparation workflow allowing in-depth proteomic analysis of microdissected FFPE tissue.

Preserved clinical material is a unique source for proteomic investigation of human disorders. Here we describe an optimized protocol allowing large scale ...

Milk Collection Methods for Mice and Reeves' Muntjac Deer

Animal models are commonly used throughout research laboratories to accomplish what would normally be considered impractical in a pathogen&#8217;s native host. Milk collection from animals allows scientists the opportunity to study many aspects of reproduction including vertical transmission, passive immunity, mammary gland biology, and lactation. Obtaining adequate volumes of milk for these studies is a challenging task, especially from small animal models. Here we illustrate an inexpensive and facile method for milk collection in mice and Reeves&#8217; muntjac deer that does not require specialized equipment or extensive training. This particular method requires two researchers: one to express the milk and to stabilize the animal, and one to collect the milk in an appropriate container from either a Muntjac or mouse model. The mouse model also requires the use of a P-200 pipetman and corresponding pipette tips. While this method is low cost and relatively easy to perform, researchers should be advised that anesthetizing the animal is required for optimal milk collection.

Milk Collection Methods for Mice and Reeves&#039; Muntjac Deer

Milk collection from animal models facilitates various research avenues: understanding passive immunity, identifying pathogens responsible for vertical transmission and, through the use of transgenic mice, even commercial production of proteins found in human breast milk. Here we illustrate a simple method for milk collection in mice and Reeves&#8217; muntjac deer.

Animal models are commonly used throughout research laboratories to accomplish what would normally be considered impractical in a pathogen&#8217;s native ...

Dissection of the Auditory Bulla in Postnatal Mice: Isolation of the Middle Ear Bones and Histological Analysis

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called the auditory bulla houses the ossicles, whereas the auditory capsule encloses the inner ear, namely the cochlea and semicircular canals. Murine ossicles are essential for hearing and thus of great interest to researchers in the field of otolaryngology, but their metabolism, development, and evolution are highly relevant to other fields. Altered bone metabolism can affect hearing function in adult mice, and various gene-deficient mice show changes in morphogenesis of auditory ossicles in utero. Although murine auditory ossicles are tiny, their manipulation is feasible if one understands their anatomical orientation and 3D structure. Here, we describe how to dissect the auditory bulla and capsule of postnatal mice and then isolate individual ossicles by removing part of the bulla. We also discuss how to embed the bulla and capsule in different orientations to generate paraffin or frozen sections suitable for preparation of longitudinal, horizontal, or frontal sections of the malleus. Finally, we enumerate anatomical differences between mouse and human auditory ossicles. These methods would be useful in analyzing pathological, developmental and evolutionary aspects of auditory ossicles and the middle ear in mice.

We present a protocol to isolate the auditory bulla, capsule, and ossicles from postnatal mice for whole mount and histological analysis.

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called ...

Cystometric and External Urethral Sphincter Measurements in Awake Rats with Implanted Catheter and Electrodes Allowing for Repeated Measurements

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually performed as terminal analysis under urethane anesthesia. It is well known that anesthetic drugs can influence bladder function. Hence, the aim of this technique is to perform cystometric measurements of the urinary bladder and external urethral sphincter in lightly restrained awake rats. For this purpose, a bladder catheter is implanted into the bladder dome. Subsequently, two electrodes are implanted bilateral to the external urethral sphincter and a ground electrode is sutured to a non-responsive skeletal muscle. The bladder catheter and the three electrodes are finally tunneled subcutaneously to the neck region and affixed to a harness. With this technique, the lower urinary tract can be measured at multiple time points in the same animal to assess lower urinary tract function. The main application of this technique is the follow-up of simultaneous urinary bladder and external urethral sphincter function in awake healthy rats and after induction of a disease or injury. Moreover, subsequent lower urinary tract monitoring can be performed during evaluation of the disease/injury and to monitor treatment efficacy.

This protocol first describes the surgical procedure of the permanent implantation of a urinary bladder catheter combined with external urethral sphincter electrodes, and second, the measurement of the function of the urinary bladder and external urethral sphincter in implanted awake animals.

Lower urinary tract function is mainly assessed by means of cystometric bladder function analysis in rodents. Conventional cystometries are usually ...

Determination of Reproductive Competence by Confirming Pubertal Onset and Performing a Fertility Assay in Mice and Rats

Assessment of reproductive competence is critical for understanding the impact of a treatment or genetic manipulation on the reproductive axis, also termed the hypothalamic-pituitary-gonadal axis. The reproductive axis is a key integrator of environmental and internal input adapting fertility to favorable conditions for reproduction. Prior to embarking upon a fertility study in mice and rats, sexual maturity is evaluated to exclude the possibility that the observed reproductive phenotypes are caused by delayed or absent pubertal onset. This protocol describes a non-invasive approach to assess pubertal onset in males through the determination of preputial separation, and in females through vaginal opening and first estrus. After the confirmation of the completion of puberty and the achievement of sexual maturity, a fertility study can be initiated. The procedure describes the optimal breeding conditions for mice and rats, how to set up a fertility study, and what parameters to evaluate and determine if the treatment or gene deletion has an impact on fertility.

Many treatments and genetic mutations impact the timing of sexual maturity and fertility. This protocol describes a non-invasive method to evaluate pubertal onset in mice and rats prior to setting up a fertility study in sexually mature animals.

Assessment of reproductive competence is critical for understanding the impact of a treatment or genetic manipulation on the reproductive axis, also ...

Tissue Collection of Bats for -Omics Analyses and Primary Cell Culture

As high-throughput sequencing technologies advance, standardized methods for high quality tissue acquisition and preservation allow for the extension of these methods to non-model organisms. A series of protocols to optimize tissue collection from bats has been developed for a series of high-throughput sequencing approaches. Outlined here are protocols for the capture of bats, desired demographics to be collected for each bat, and optimized methods to minimize stress on a bat during tissue collection. Specifically outlined are methods for collecting and treating tissue to obtain (i) DNA for high molecular weight genomic analyses, (ii) RNA for tissue-specific transcriptomes, and (iii) proteins for proteomic-level analyses. Lastly, also outlined is a method to avoid lethal sampling by creating viable primary cell cultures from wing clips. A central motivation of these methods is to maximize the amount of potential molecular and morphological data for each bat and suggest optimal ways to preserve tissues so they retain their value as new methods develop in the future. This standardization has become particularly important as initiatives to sequence chromosome-level, error-free genomes of species across the world have emerged, in which multiple scientific parties are spearheading the sequencing of different taxonomic groups. The protocols outlined herein define the ideal tissue collection and tissue preservation methods for Bat1K, the consortium that is sequencing the genomes of every species of bat.

This is a protocol for the optimal tissue preparation for genomic, transcriptomic, and proteomic analyses of bats caught in the wild. It includes protocols for bat capture and dissection, tissue preservation, and cell culturing of bat tissue.

As high-throughput sequencing technologies advance, standardized methods for high quality tissue acquisition and preservation allow for the extension of ...