Necropsy procedure

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Once the mouse has been euthanized it should be superficially disinfected by submersion in a dilute solution of a germicidal detergent such as Calgon Vestal Process NPD One Step Germicidal Detergent (ConvaTec, St. Louis, MO), or a solution of 95% ethanol. When necropsying a mouse with an abnormality of the hair coat, it is important to collect samples of the hair before the mouse is dipped in the disinfectant. The hair should be plucked manually using the thumb and forefinger. Do not use forceps, as this will damage the hair shaft. Plucking of the hair allows for examination of the whole hair shaft from root to tip. The hair should be stored in a clean Nunc cryopreservation tube (Nalge Nunc International, Denmark). When studying mouse mutants, standardization of collection techniques is as important with hair as with other organs. Hair samples should be collected from the same area on every mouse in a study. We pluck the hair from the left flank from shoulder to hip to get hairs from both the dorsal and ventral surfaces. Be sure to take any skin that will be collected from an area of the mouse that was not plucked to prevent artifactual changes in the hair follicle being examined. If the vibrissae are abnormal, these should be collected as well, from the same side as the hair is plucked. They should be stored in a separate container.

If a mouse has a skin abnormality, remove hair prior to disinfection. This is usually accomplished by shaving the mouse with electric hair clippers such as the Oster Finisher Trimmer (Cat. # 76059-030, Oster Professional Products, McMin-nville, TN). These clippers are easy to handle and have a small blade that is ideal for mice or other small mammals. If complete hair removal is desired, there are commercially available depilatory products (Nair, Carter-Wallace Inc., New York, NY; Neet, Reckitt & Coleman Inc., Wayne, NJ) that can be applied to the mouse after shaving. These products should be left on for 2 to 3 minutes, then rinsed off under warm running water, which will wash away the hair as well.

At this point, the mouse may be disinfected. The disinfectant also serves to wash off any loose hairs and mats down hair on an unshaven mouse for ease of examination. After letting some of the disinfectant drain from the mouse back into the container, place the mouse on one to two layers of absorbent paper towel on a cork board. The cork board should be approximately 14 cm x 21.5 cm in size, which is large enough for two mice, yet easy to move about during the necropsy. It should be at least 1.0 cm thick.

If skin is to be collected for characterization of a mutant, it should be done at this point. With the mouse ventral surface down on the board, gently grasp a fold of dorsal skin from the caudal region and make a small incision with the scissors. Carefully cut out a rectangular piece of skin along the dorsal midline from the thoracolumbar junction to the interscapular region and collect it in the desired medium. We often collect skin from one mouse for many different assays. Preservation methods are discussed in the fixatives section, but it is important to remember that presentation is just as important as your choice of fixative. Skin should be laid out flat on a piece of aluminum foil or mesh backing for fixation so that it may be cut neatly into thin strips for histology. Orient the skin sample cranial-caudally, and trim lengthwise on this axis to optimize the orientation of the hair follicles.

After collecting dorsal skin, collect a sample of ventral skin in a similar fashion. Fresh samples of both dorsal and ventral skin may be frozen in OCT for antigen expression studies. The skin from the head should also be collected, including the pinnae (ears), eyelids, and muzzle. All may be carefully peeled and trimmed from the skull as a unit and mounted flat on a piece of aluminum foil for fixation. Tail skin may be collected by severing the tail from the body and making an incision with a tip of a scissors down its length. Grasp the loose skin from the base of the tail, and strip the skin away from the bone and tendons. Mount the tail skin flat on a piece of foil as you did the other samples. Tail and head skin are often collected toward the end of the necropsy.

After skin has been collected, place the mouse ventral side up and pin each limb firmly to the board. The rear feet may be pinned between the gastrocnemius tendon and the bone, while the front feet may be pierced through the skin, between the metacarpal bones, in order to do the least damage to the tissues being collected. During the necropsy, the board may be rotated easily to adjust the position of the mouse, providing different angles of access for organ collection.

If the skin is not to be collected for a study, you will begin your necropsy by pinning the mouse down, ventral side up, as describe above. With either a #12 scalpel blade and #3 handle or a pair of sharp/sharp iris scissors, make a ventral midline longitudinal incision through the skin, from the external genitalia to the ramus of the mandible, then cut from the genitalia laterally toward the rear feet, along the medial surface of the rear legs. On female mice, this incision passes between the fourth and fifth nipples of each side. Grip the skin on either side of the incision and pull gently outward, or use light strokes of the scalpel, to separate the skin from the abdominal muscles. Reflect the skin far enough so that it does not interfere with the rest of the necropsy.

With the skin reflected back, collect the external lymph nodes. The peripheral lymph nodes are located on either side of the salivary gland (cervical lymph nodes), under each of the fore legs (axial lymph nodes), and on the inside of each rear leg (inguinal lymph nodes). The cervical lymph nodes are collected attached to the salivary gland. The inguinal lymph nodes can be located, if you look carefully, in the fat pad on the medial surface of the rear leg. They may be slightly darker in color than the fat and are usually no larger than 0.10 to 0.20 cm. The easiest way to collect them is to remove the fat pad with the lymph node embedded in it and trim away as much of the fat as possible before placing the lymph node in the fixative.

Next, using a clean, sterile set of instruments, grip the abdominal muscles at the inguinal region with forceps and lift firmly, making a small incision to let air into the abdomen. This will cause the abdominal wall to stretch further and the viscera to fall away so a more aggressive incision can be made without contaminating the tissues inside. Continue the cut through the abdominal muscles on each side, extending from the inguinal midline to the lateral thorax. This will neatly expose the viscera. If culturing from swabs within the abdomen is necessary, a third sterile set of instruments should be used at this time, with a new set used for each culture taken.

After any necessary abdominal samples have been taken for microbiological culture, tissue collection may begin. As a general rule, no more than fifteen minutes should pass between euthanization of the healthy mouse and collection of its tissues. This should allow adequate time for photography, culturing and other procedures. Throughout the necropsy, the visceral organs should be evaluated to determine if they are in their proper anatomic orientation. Some mutations such as situs inversus (iv) can cause the orientation to be reversed.19 All tissues and organs should be carefully checked for abnormalities and all observations should be noted. Gross photographs of any external or internal abnormalities are important to obtain in the study of a new mutant. See Chapter 6, Photography of Laboratory Mice, for details.

The intestines should be collected first because autolysis begins quickly in these organs. Grasp the cecum with forceps, lift it up and cut away the small intestine at the rotund sac, where the ileum meets the cecum. This is the smaller of the two intestines joining at the cecum. Drop the cecum and grasp the severed end of the ileum, pulling it gently as the small intestine begins to unravel. Carefully cut any mesenteric tissue that provides resistance. Be sure not to puncture or sever the intestine itself, as this will make it more difficult to inflate and prepare for histology. If the intestine does break at any point, simply lay the broken section on a moist portion of the paper towel, maintaining proper orientation, then continue the procedure. Toward the duodenojejunal flexure, the pancreas is firmly adherent to both the small and large intestine. Care must be taken to separate the two intestines and the pancreas, while preserving the integrity of all three. Continue freeing the small intestine until you reach the stomach, then release the tension on the intestine. Before separating these organs, open the small intestine just distal to where it joins the stomach. Gently compress the gall bladder. Bile should empty out of the common bile duct, indicating that the duct is patent. Now cut away the stomach from the small intestine at the pars pylorica. Lay the intestine down in a large loop around the mouse remembering which end is the duodenum and which is the ileum. Intes tinal rolls must be oriented the same each time to allow the pathologist to interpret them properly. It helps to develop habitual patterns of tissue collection to avoid errors.

For proper fixation, the intestine must be inflated with fixative before it is rolled or otherwise prepared for histological presentation. This procedure is simple and rapid but runs the risk of injury by squirting fixative into eyes accidentally. Be sure to take precautions, wear safety goggles, and use a fume hood if one is available. Use a 10 cc syringe with a 17 to 22 gauge needle. Fill the syringe with fixative and introduce the needle into either end of the intestine. Gently depress the plunger on the syringe and the intestine will slowly inflate. Making several injections along the length of the intestine is generally safer for the technician and is often necessary to provide proper inflation (Figure 5.4). An intestine that has been over inflated is much more difficult to roll. To make a second injection, simply pierce the wall of the intestine and clamp it firmly around the tip of the needle to prevent backflow of the fixative and proceed as before.10 Some laboratories prefer to open the entire intestine and remove digested food material and feces prior to rolling.20 This approach yields good mucosal fixation but may also cause damage to the mucosa.

FIGURE 5.4 Intestines are removed and inflated by injection with fixative. (Drawn by Ingrid K. Sundberg.)

Intestines can be rolled full length for histological presentation.1020 Each roll must be able to fit comfortably in a histological cassette. In an average sized adult mouse, this requires the small intestine to be cut into three equal pieces. With practice you will learn to judge the number of sections required for proper sized rolls.

Another approach to presenting intestines is similar to the way they are routinely collected in larger animals. Representative segments are cut and fixed by immersion. More precision comes by laying out the entire gastrointestinal tract and cutting segments out at specified distances from the anus or pyloris.

To create what are called "Swiss Rolls" for full-length histological presentation, roll the inflated intestine in concentric, centrifugal circles on a piece of unlined index card (Figure 5.5), letting as little fixative as possible flow out of the segment. If the

FIGURE 5.4 Intestines are removed and inflated by injection with fixative. (Drawn by Ingrid K. Sundberg.)

fixative drains, the intestine may flatten, making it more difficult to roll. We use large index cards cut into strips approximately the width of our histological cassettes to mount the tissues on. As mentioned previously, the orientation of the segments is important and must be agreed upon with the pathologist. It is commonly accepted to keep the end of the intestine proximal to the stomach toward the center of the roll. Once the intestines have been rolled onto the paper, let them stiffen for a few minutes before placing them into a jar of fixative, to prevent the roll from unwinding.

FIGURE 5.5 "Swiss roll" of intestines. (Drawn by Ingrid K. Sundberg.)

Intestinal rolls take practice to master, and one may wish to develop his or her own techniques for the best presentation. The important objectives to remember are (1) do not over inflate the intestine, (2) keep the proper orientation, (3) make the rolls smaller than the cassettes, and (4) try not to let the fixative drain while rolling the segments.

Before the colon and attached cecum may be dealt with, one must first remove the reproductive organs, including the preputial gland or the clitoral gland, and urinary bladder. In the male mouse, testes may be gently grasped by the inguinal fat pad, cut away from the other viscera and placed on a bit of cardstock. Orient them in such a way that the epididymis and testis are in the same plane and may be trimmed simultaneously for histological presentation. The preputial gland is a paired organ located subcutaneously between the penis and the rectal opening. It may be collected by grasping one edge of the gland with your forceps, cutting it away from the abdominal wall and placing it directly into fixative. The seminal vesicles, urinary bladder, and penis may be removed as a unit by gently grasping the apex of the seminal vesicles with forceps and lifting it away from the colon. Insert the tip of heavy duty scissors between the colon and the pelvis and cut the bone on both sides, then lift the reproductive tract further and cut away the connective tissue between it and the colon. Remove the reproductive tract and arrange it on a card, then place it in the fixative.

The female reproductive tract is removed in a similar fashion. The clitoral glands are normally not easily visible, but are located subcutaneously just in front of the vaginal opening. The best way to collect these glands is to cut a small square of 0.5

Swiss Roll Intestine

FIGURE 5.5 "Swiss roll" of intestines. (Drawn by Ingrid K. Sundberg.)

cm to 0.8 cm of abdominal muscle and overlying skin immediately anterior to the clitoris. This section will include the clitoral glands. Smooth this piece of tissue gently onto a piece of white cardboard and, using parallel pencil marks, indicate the area where the glands are expected to be. After placing the clitoral glands in fixative, grasp the fat pad of one ovary and cut it free from the mesentery, laying that ovary and uterine horn over on the other side of the colon. Then cut the pelvic bone as before, cutting the other half of the uterus and its ovary free from the surrounding fat and moving it out of the way before severing the contralateral pelvic bone. Remove the entire female reproductive tract as you did the male organs, then arrange it on a card before fixation to maintain orientation.

Now grasp the cecum with the forceps and pull it gently, cutting away the mesentery to free the colon. Trim the anus away from the surrounding skin and lay the structure down on your moist work surface. Inflate the colon with fixative as you did the small intestine (Figure 5.6). Insert the needle carefully into the anus and slowly depress the plunger of the syringe. Fecal matter in the colon can block the flow of the fixative. Gently express the feces and then inflate with fixative. The cecum should be inflated just slightly through the ileocecal junction, cut from the colon, then laid out on a flat surface to stiffen before dropping it into the fixative. The colon should be rolled similar to the rest of the intestine, with the same orientation. The colon is often stiff and more difficult to handle due to the presence of fecal pellets, and should be left out to harden for quite some time once it has been inflated and arranged properly on the card.

FIGURE 5.6 Inflation of the cecum with fixative. (Drawn by Ingrid K. Sundberg.)

The last of the gastrointestinal tract to collect is the stomach. Grasp this organ at the pylorus and gently pull it away from the liver until you can see the esophagus, which presents itself as a narrow whitish structure extending from the center of the stomach upward. Cut this carefully while maintaining your grip on the pylorus and pull the stomach away from the liver, trimming away any mesentery or bits of pancreas that may adhere to the gastric surface. Inject a small amount of fixative into the stomach through the pylorus, similar to the way the cecum was inflated, then place it in the jar with the other organs.

The Necropsy The Mouse

FIGURE 5.6 Inflation of the cecum with fixative. (Drawn by Ingrid K. Sundberg.)

The spleen and pancreas are connected and are often removed as a unit unless it is necessary to examine one or the other separately, as in the case of a mouse model for diabetes where special fixative (Bouin's solution) and stains (Aldehyde Fuchsin) are used to examine the beta cells of the pancreatic islets. Grasp the pancreas gently with the forceps and pull upward, cutting away any mesenteric tissue that adheres to the spleen or pancreas to free the structures, then place them directly into the fixative.

Kidneys may be removed by grasping the surrounding fat and pulling upward while cutting around the organs. The adrenal gland is a small white structure that lies within the perineal fat pad just anterior to the kidneys. It should be left within the fat that clings to the kidney and the two should be presented to histology as a unit. It is important for the pathologist to be able to distinguish between the right and left kidneys. When it comes time to trim the kidneys after fixation, the right kidney should be cut transversely, while the left is cut lengthwise (left/long) for histological presentation. A small transverse incision in the right kidney before fixation will allow you to distinguish between the two and improve fixation if left intact.

The liver is the last organ to be removed from the abdominal cavity. To accomplish this, it helps to enter the thorax so you may use the diaphragm as a handle to trim out the liver. Move the liver gently out of the way with the edge of a forceps. Then grasp the xiphoid process firmly with the forceps and pull upward to create a negative pressure within the thorax. Cautiously cut through the ribs and diaphragm of one side. This will allow the air to enter the thoracic cavity, and the lungs within will shrink away from the diaphragm. If a thoracic microbiological culture is to be taken, extend the cut through the diaphragm and rib cage to expose the lungs, then use sterile scissors and forceps to take a tissue sample before proceeding. After all samples for microbiological cultures have been taken, grasp the diaphragm with forceps and trim it completely away from the ribs, being cautious near the spinal column so as not to cut the liver. The liver may then be easily lifted away. Place the liver on a damp work surface and separate the right and left medial lobes as a unit, along with the gall bladder. This may be accomplished by folding these lobes back onto the work surface and trimming the ligaments that connect them to the remaining liver lobes. The lateral left lobe is the largest and should be separated from all others in a similar fashion. The remaining smaller lobes may be placed in the fixative together. When separating the liver lobes, take care to handle them gently. The liver is very fragile and is easily damaged.

The thoracic cut may now be extended through the rest of the ribs at the costochondral junction to just short of the internal thoracic vein and artery on both sides. Often the mediastinum between the area of the heart and thymus continues pulling on the excised part of the rib cage, so it must be carefully trimmed away to prevent the rib cage from falling back over the rest of your work.

At some point, before you cut into the cervical area, the salivary glands should be removed. These glands lie subcutaneously in the ventral cervical area. Gently pulling back the skin in this area will sufficiently expose the glands for removal. The tip of this lobed structure is narrower and lies closer to the thorax. Grasp it gently with your forceps and slowly pull upward, freeing it from the surrounding tissue with your scissors, then cut the glands horizontally across their base and place them in the fixative.

To remove the heart and lung, turn the cork board around so that the mouse's head is facing you. With your forceps, lift the lower jaw, (remember, the mouse is on its back) and cut through the hinge of the mandible with the scissors, separating the lower jaw from the rest of the skull. The epiglottis will now become apparent. Being careful not to puncture the trachea or the esophagus, continue your cut on either side of the neck until you reach the first rib. Using the tip of your scissors, sever the first rib and continue to dissect carefully through the remaining ribs, avoiding the trachea, esophagus, and lung. Repeat this on the other side. Once all the surrounding tissue has been removed, pull up gently on the mandible, trimming any remaining mesenteric tissue from the underside of the lungs. The entire structure, including the lungs, heart, trachea, epiglottis, lower jaw, and tongue will be removed as one. The lungs are another structure that must be inflated with fixative to ensure proper histological presentation. Use the same syringe and needle setup as you used for the intestines. Slip the tip of a needle into the trachea via the glottis, which is normally the most apparent hole at the base of the tongue. Clamp down around the needle with a pair of forceps and slowly depress the plunger of the syringe. If the lungs begin to expand and blanch, you have successfully found the trachea. If the fixative flows out in a puddle between the lungs, try again. Inflate the lungs very slowly, stopping when they are about the size that they would normally be on inhalation. Overinflating the lungs can damage the alveoli, often causing diagnostic difficulty.

Once the lungs have been inflated, cut through the trachea and esophagus to separate the mandible from the heart and lungs. The heart, lungs, and thymus should remain together as a unit for fixation. The tongue should be separated from the mandible by grasping the tip of the tongue with your forceps and cutting it off at the base. Place the tongue, mandible, heart, lung, and thymus into the fixative.

The next organ to be collected is the brain. To access the brain, cut the spinal cord at the base of the skull. Slowly pull and cut the skin away from the skull if the skin has not already been removed. Be sure to cut carefully around the eyes, leaving the eyelids attached to skin and avoiding damage to the eyes themselves. To focus on the eyes, you may wish to remove them from the skull. To remove the eye, sink a pair of curved forceps behind the orbit, grasp the optic nerve and pull outward until the eye has been freed from the socket and may now be fixed and embedded separately. However, histological presentation of the eye within the skull is often sufficient for viewing many abnormalities.

Cut any remaining vertebrae off the skull. You will see a mass of white matter protruding from the foramen magnum. This is the spinal medulla. Slip one blade of your scissors in between the neural tissue and the bone and make two small longitudinal cuts in the occipital bone (Figure 5.7), one on each side of the spinal medulla. Hold the skull between your thumb and forefinger and, with your forceps, grasp the edge of the occipital bone and pull upward to neatly break it off. Then, gently insert the tip of your scissors between the brain and the skull and make a cut in the skull along the sagittal suture (Figure 5.8). Continue breaking away the interparietal and the parietal bones in the same manner, being careful not to harm the delicate brain below. The frontal bone comes to a slight point at the intersection of the sagittal and coronal sutures. This area should be broken off as well, to allow the brain to be removed cleanly from the cranial cavity. There may be a thin reddish membrane around the brain, particularly in the area of the cerebrum. This is the meninges and must be removed carefully with forceps or it will cut into the brain as you try to remove the brain from the skull. Once you have removed the meninges, turn the skull upside down over the jar of fixative. Gently work the forceps between brain and bone and pull away the connective tissues, freeing the brain from the cranial cavity. The brain will fall into the fixative. Place the skull into the fixative as well.

FIGURE 5.7 Initial cuts in the occipital bone to access the brain. (Drawn by Ingrid K. Sundberg.)

FIGURE 5.8 Second set of cuts down the sagittal suture for exposure of the brain. (Drawn by Ingrid K. Sundberg.)

The spinal column may now be collected. Grasp the proximal end of the spine and lift it away from the skin, cutting away the fascia that hold the two together. Cut through the pelvis to sever the hind limbs from the distal vertebrae, then cut the tail away as well. Cut the ribs away from the spine, as close as possible, without damaging the vertebrae. Place the spinal column in fixative, making sure to keep it straight so it is oriented correctly when it is time to trim the tissues.

Trim away the front and rear limbs from the remaining skin and place them in fixative. If skin was not collected earlier in the necropsy, it may still be important to save it for your archives. We take the full pelt of the mouse and fold it in half

FIGURE 5.7 Initial cuts in the occipital bone to access the brain. (Drawn by Ingrid K. Sundberg.)

FIGURE 5.8 Second set of cuts down the sagittal suture for exposure of the brain. (Drawn by Ingrid K. Sundberg.)

over a strip of cardstock before placing it in fixative. This is a quick and easy way to save the skin in an orderly fashion for future use, if necessary.

In summary, when working up the effects of a new mutation, it is important to collect study sets of total tissues in a methodical, standardized fashion to avoid diagnostic discrepancies due to presentation. From these study sets, a more focused tissue collection protocol may be developed, concentrating on those tissues known to express abnormalities in your mutant. Always remember the importance of adopting standard criteria for tissue collection, agreed upon by the technicians, researchers, and the pathologist who will be reading the slides.

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