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

The overall goal of this procedure is to perform a basic rodent necropsy. This is accomplished by first recording basic information about the animal. The second step of the procedure is to examine the viscera.

The third step is to collect samples such as organs or mesenteric lymph nodes. Ultimately, the health status of an animal can be determined through histopathology or other techniques. Generally, people new to performing a necropsy will struggle because rodent anatomies learned on the job.

These methods can help answer key questions about the health of animals in a research colony. Visual demonstration of these techniques is essential as it takes time to learn the locations of various tissues and how to identify normal from abnormal tissue. Before euthanizing the animal, collect information on its history and record its current characteristics such as physical condition, behavior, sex, age, weight, strain, and so forth.

This record is a vital age to the person who will be examining the tissues prior to euthanizing the animal. Be sure that appropriate personal protective equipment is used. Prepare a dissection, board forceps, scissors, labels for containers, fixative, media and collection vessels.

After euthanizing the animal, assess its bodily condition, check for skin and coat abnormalities, emaciation, and dehydration. Also search for evidence of artificial manipulations, implants or surgical scarring on the animal. Next, using a dissecting microscope, examine all of the external orifices in detail.

These include the ears, eyes, nose, mouth, anus, and genitalia. Now, lay the animal on its back on the dissection board, lift its skin with forceps and use scissors to cut the skin open from the anus to the chin. Next, reflect the skin to the sides and cut through the abdominal wall and rib cage.

Expose the thoracic viscera by making two cuts laterally up each side of the rib cage. Cut the diaphragm. Then remove the rib cage by cutting across the top of the sternum.

Now, examine the exposed thoracic and abdominal organs. Know any color changes, unusual odors, size, differences, and missing or dislocated organs. Examine the consistency of the organ surfaces and note any tissue masses or fluid filled pockets.

Also, note the presence of fluid in either cavity. Continue the examination with the gastrointestinal tract. Examine the mesentery for enlarged lymph nodes or tissue masses.

Describe the contents of the gastrointestinal tract and check carefully for thickened walls, masses, or hemorrhage. To examine the interior of the kidneys, use a blade to make a longitudinal section of the left kidney and a cross section of the right kidney off center of the midline. Check the parenchyma for any abnormalities.

Now inspect the urogenital system in males. Identify the seminal vesicles, the testes, the epididymus, the ureters, and the perusal glands in females. Identify the uterus, the ovaries, and the ureters.

Examine the urogenital system for blockages, fluid pockets, hemorrhage, or other abnormalities. After completing these examinations, isolate and excise the target organs for further examination. Typically, the heart, liver, kidneys, and spleen are collected as they're excised.

Submerge each organ in a fixative or follow the first steps of a protocol requiring these tissues such as freezing or immunohistochemistry to get adequate fixation. Generally, tissues are submerged in a preservative like 10%NBF at a ratio of 20 volumes fixative to one volume tissue. When working with rats, collect the bronchial aspirate before the nasal aspirate.

Begin by reflecting the skin in subcutaneous tissues away from the cervical area, thus exposing the salivary glands and the cervical musculature. Then remove those tissues to expose the trachea. Make a three millimeter incision into the trachea and insert a pipette loaded with one milliliter of sampling fluid into the tracheal lumen directed quarterly.

Slowly flush the sampling fluid through the trachea and into the lungs three times. Then withdraw the sampling fluid and remove the pipette from the trachea. Not all of the fluid will return into the pipette.

Therefore, repeat the process if more fluid is needed. For testing in mice. To take a nasal aspirate or bronchial aspirate sample, access the nasal pharyngeal ATU using sterilized scissors.

Sever the temporal mandibular joint and reflect the mandible away from the maxilla, exposing the nasal pharyngeal atu to collect the nasal aspirate. Insert a pipette loaded with one milliliter of sampling fluid into the mouse's nasal pharyngeal atu, or the rat's tracheal lumen. The pipette should be directed cran and pushed in.

So the tip contacts the nasal palate. Once there slowly inject the sampling fluid. As the fluid enters the nasal cavity, it will form menisci at the nasal orifice and should also be visible through the translucent oral palate.

However, if fluid exits through the mouth, reorient the pipette. Once fully injected, withdraw the sampling fluid from the nasal cavity into the pipette, and then remove the pipette. Begin by grasping the trachea with forceps and snip through it just above the forceps.

Next, gently tug the trachea upwards, snipping tissue connections until the entire set of thoracic tissues can be excised. Now, lay the lungs flat on the work surface, and loosely tie a piece of suture or string around the trachea. Do not tighten the knot, or you will not be able to pass the needle into the trachea.

Now, fill a syringe with fixative and insert the needle into the opening of the trachea while securing the trachea to the needle. With forceps, slowly fill the lungs with fixative until they're fully inflated. The volume of fixative will vary according to the animal's age, strain, and health.

If underinflated tissues will appear flat, if overinflated, foamy liquid will seep from the tissue once filled, tighten the suture material or string surrounding the trachea to prevent backflow of fixative out of the lungs. Then place the inflated lungs in fixative. Place the carcass on a clean dissection board in ventral recumbent and use scissors and forceps to remove the skin and muscle overlying the calver.

Then decapitate the carcass using small scissors. Insert the bottom blade into the foramen magnum and cut through the midline of the calver using an upward motion. Now, expose the brain with forceps by pulling open the calver.

If the pathologist prefers to cut sections of the brain while in the skull, immediately place the skull with exposed brain into 10%NBF. Otherwise, grasp the head around the nose with forceps or fingers and gently invert the skull so that gravity will help the tissue fall from the skull.Carefully. Slide curve forceps along the outer edge of the brain beginning at the olfactory lobes.

Continue sliding the forceps along the skull towards the cerebellum, while gently pinching nerves with the same forceps. When the brain releases place into fixative, to locate the mesenteric lymph nodes first, locate the large comm shaped secum of the intestine. The mesenteric lymph nodes are the yellow ovoid or spherical lumps of tissue in the mesentery adjacent to the cecum.

They're often slightly thicker and firmer in texture than the surrounding mesentery and fat. A dissecting scope may be useful to differentiate tissues at first, but with experience it will not be necessary. Using forceps and scissors, isolate the mesenteric lymph nodes, then transfer them to a storage tube to be used for culture, PCR or histopathology.

After completing the protocol, the following is available to a pathologist or scientist, a written report of objective and subjective condition at euthanasia. An objective description of the tissues prior to their fixation, all of the major abdominal and thoracic organs in fixative, the brain and mesenteric lymph nodes and samples of nasal and bronchials aspirate. Once mastered, an abbreviated necropsy can take less than 30 minutes.

While performing this procedure, it's important to remember to record your observations of the condition of the animal and its tissues. After watching this video, you should have a good understanding of how to perform a complete rodent necropsy, including collection of perfused lungs, mesenteric lymph nodes, and the brain.