1. Place the animal with the ventral side up on a clean dissection board and pin it. Avoid holding the animal in one hand to keep your hands clean and to cut the organs accurately.
2. Using soaked cotton, clean the ventrum to avoid the introduction of animal hair into the body that may infect the organs.
3. Hold the skin with dissecting forceps and raise it above the abdomen.
4. Make an incision through the body wall muscles by putting the scissor just anterior to the genital opening and continue cutting on one side of the midline, ventral to the thoracic cavity.

Note: blunt-ended scissors (either two blunt-end scissors or one blunt, one sharp point scissors) are recommended to prevent organ damage.

5. Keep the fur away from the body by pulling the skin from both sides of the thoracic cavity.
6. Isolate the larger parts of the lungs and store them in the cryogenic tubes quickly.
7. Clean the forceps and scissors using a solution containing bleach, water, and ethanol.
8. Find the spleen by moving the stomach to the left (the spleen has a triangular shape).
9. Pull the spleen out by holding it with the forceps and get rid of the white tissue with the help of sharp pointy scissors.
10. Clean the forceps and scissors with the cleaning solution.
11. Displace the bowels to find the kidneys beneath the intestines.
12. Separate the kidneys and place them in two cryogenic tubes for storage (if the kidneys are too big, cut them into pieces).
13. Clean the dissecting instruments with the cleaning solution.
14. Dissect the diaphragm and keep it in a cryopreservative tube.
15. Place the cryogenic tubes containing the dissected organs in a liquid nitrogen tank (Do not open the tank again and again rather put the tubes at one time to prevent nitrogen evaporation from the tank).
16. Wash the forceps and scissors with the solution containing bleach, water, and ethanol).
17. Identify the sex of the animal by observing the genital organs and counting the embryos or measuring the size of the testes.
18. Tag the leg of the animal. Using fine point forceps, perforate the skin and introduce the string.
19. Place the animal into an ethanol-filled jar.
20. Clean and wash the dissection board and the operative area.

Dissection of Mouse EDL and Soleus Muscles (Bröllochs, 2017)

1. Fix the leg of the animal with pins in a flexed position to the dissecting dish and cover the leg with Ringer solution.
2. Remove the skin of the leg and get rid of the excessive fascia surrounding the muscles.
3. With the help of the forceps, hold the Achilles tendon. Use the bodkin side of the scissors to rub the gap open between the Achilles tendon and the other tendons. Cut the Achilles tendon as close as possible to the foot when the gap is visible.
4. As the gap starts to open, support that gap to open up fully.
5. The Soleus muscle will be visible as the gap opens up to the knee. The Soleus muscle is dark pink, identify it by its color. Carefully cut the tendon near the knee.
6. Hold the Soleus muscle from its upper tendon gently with the forceps and pull it down slowly and carefully while (if needed) removing the surrounding fascia that is still holding it. Cut the lower tendon after freeing the whole muscle from the fascia.
7. Isolate the Soleus muscle from the leg and remove the leftover fascia. For more comfortable handling, one should move the muscle to a new dissecting dish, fix it with pins, infuse it with Ringer solution, and store it at 4 °C.
8. To dissect the EDL muscle, place and pin the leg into a stretched position first and remove the fascia.
9. Rub the lower leg’s tendons. The 4 EDL tendon ends are next to the V-looking Tibialis tendon. Carefully lift the tendons to observe the EDLs. The toes of the paw stretch while lifting. Make an incision at the ends of the tendon as close as possible to the foot.
10. A pocket will open up as the ends are cut. Remove the surrounding fascia. Isolate the EDL muscles from the pocket to visualize the upper tendon. Dissect the upper tendon then.
11. Isolate the EDL muscle from the leg and remove all the fascia and the fat.
12. Place both the dissected muscles in a clean dissecting dish. Lightly stretch the muscles and pin them. Cover the dissected muscles with a 3.7% PFA.
13. Store the dish at 4°C overnight after covering it with the parafilm.

Gross Postmortem Location and Examination of Heart, Lungs, Liver, Kidneys, and Spleen Method (Parkinson et al., 2011)

1. Examine the animal for skin and coat abnormalities, emaciation, or dehydration.
2. Keep a log of artificial manipulations, implants, or surgical scarring, if any.
3. With the help of the dissecting microscope examine all the external orifices (ears, eyes, nose, anus, genital openings, and oral cavity) for secretion or bleeding.
4. Place the euthanized animal in dorsal recumbency on a clean dissection board.
5. With the help of dissection scissors, make an incision through the skin from the length of the ventrum extending from the anus to the chin, reflecting the skin and incising the abdominal wall, opening the abdominal viscera, salivary and preputial/clitoral glands, and cervical and axillary lymph nodes. Examine the thoracic viscera by making 2 incisions laterally up on each side of the ribcage, then make a cut across, at the top of the sternum, to open a wide space enough to observe all the lobes of the lung thoroughly.
6. Observe the musculoskeletal structure.
7. Assess the condition of all the organs for abnormalities. Locate and identify the heart and the lungs in the thoracic cavity. Observe the liver, kidneys, and spleen in the abdominal cavity. Check the organs for any color changes, size differences, and absence or dislocation. Record the consistency of surfaces, any additional tissue (e.g., masses), fluid pockets, or the presence of fluid in the abdominal/thoracic cavities.
8. Examine the gastrointestinal tract for contents, lack of contents, thickened walls, masses, blood clots, and hemorrhage. Using a sharp blade, cut the kidneys (left-longitudinal section, right-cross section, on the midline, but off-center) to assess parenchyma for any abnormality. Check the presence of any large lymph nodes on the mesentery.
9. Observe the urogenital system and check for any blockage, fluid pockets, hemorrhage, or other abnormalities.  

Postmortem Collection of Heart, Liver, Kidneys, and Spleen for Histopathology

1. Collect a container filled with a suitable amount of 10% neutral buffered formalin (NBF), and make sure that the container is appropriately sized and labeled.
2. Adjust the amount of 10% NBF to obtain a 20:1 ratio of fixative to tissue.
3. Lay the animal’s carcass in dorsal recumbency on a clean dissection board and expose the tissues of interest.
4. Isolate the tissues from the animals’ carcass with the help of dissecting forceps and scissors.
5. Scrap the tissues to remove fat and unnecessary connective tissue carefully. The blood should be clean; use normal saline to rinse. Note: Do not use distilled or tap water to rinse the tissues.

6. Place the dissected tissues in the container containing 10% NBF.

Postmortem Collection and Perfusion of Lung Tissue

1. Fill a container with 10% NBF and adjust the volume of 10% NBF to obtain a 20:1 ratio of fixative to tissue.
2. Lay the animal to be dissected in dorsal recumbency on a clean dissection board.
3. Expose and display the trachea, heart, and lungs.
4. Remove the skin and muscle overlying the ventral thoracic and cervical regions using the dissecting forceps and scissors.
5. Trim the ribcage to expose the heart and lungs near the clavicle to open space, ample enough to observe all the lobes of the lung thoroughly.
6. Remove the neck muscles surrounding the sternum and ribs and ranging to the jaw, including those overlying the trachea.
7. Make two cuts by inserting the scissors under the anterior edge of the rib cage, one cut on either side, to remove the section of bone overlying the trachea.
8. With the forceps hold the trachea near the jaw and cut the trachea entirely by placing the scissors above the forceps
9. Pull out the trachea with the help of the forceps and cut the ventral tissue connections with scissors until all the thoracic tissues are removed from the body.
10. Place the lungs flat on the shelf.
11. Stitch the trachea with suturing material loosely.
12. Fill a syringe with a fixative and attach a needle, 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 is accurate).
13. Hold the trachea with the forceps and insert the needle into the trachea. Fill the lungs with fixative slowly. Keep filling until the lungs are fully inflated. Do not over-or underinflate. The amount of fixative needed depends on the age, strain, and health of the animal.
14. Fluid seeping and foaming from the tissues indicate over-inflation.
15. Flat lungs indicate under-inflation.
16. Withdraw the needle from the trachea.
17. Tighten the stitches on the trachea to prevent backflow of fixative out of the lungs.
18. Keep the inflated lungs into the fixative with a 20:1 fixative to tissue ratio.

Postmortem Collection of Brain

1. Bring an appropriately sized, labeled container(s) and fill in the required amount of 10% NBF and adjust the amount of 10% NBF to obtain a 20:1 ratio of fixative to tissue.
2. Place the animal carcass in ventral recumbency on a clean dissection board.
3. Remove the skin and the muscles surrounding the calvaria with the help of dissecting scissors.
4. Dislocate and remove the head from the animal’s body.
5. Insert the bottom blade in the foramen magnum (the opening where the skull opens into the spinal canal) with the help of small scissors, keep the scissor tips pointed upwards, and begin cutting through the midline of the calvaria.
6. Reflect both halves of the calvaria and expose the brain.
7. Place the exposed brain encase in the skull into a fixative.
8. Invert the skull gently so that the tissues fell from the skull because of the gravity.
9. Move the forceps under the brain starting at the olfactory lobes and along the outer edge of the brain, moving under the cerebrum and towards the cerebellum carefully. Gently pinch the connective tissue or nerves, with the help of the forceps, inhibiting the brain from falling from the skull.
10. Place and keep the brain into the fixative using an approximate 20:1 ratio of tissue to fixative.

Postmortem Collection of Respiratory Aspirate

1. Place the animal to be dissected in dorsal recumbency on a clean dissection board.
2. For bronchial aspirate collection in rats, get access to the respiratory tract through the trachea. For nasal aspirate in rats, reach the respiratory tract either through the trachea or the nasopharyngeal meatus. For bronchial or nasal aspirates in mice, approach the respiratory tract from the nasopharyngeal meatus.
3. Tracheal Access (recommended for rats):
   • Expose the subcutaneous tissues by moving the skin away from the cervical area.
   • Separate the salivary glands and cervical musculature to expose and visualize the trachea.
   • With the help of sterile dissecting instruments, cut the trachea to reach its lumen.
   • Note: Maintain asepsis throughout the collection.
4. Nasopharyngeal Meatus Access:
   • Cut the temporomandibular (jaw) joint and move the mandible away from the maxilla to expose the nasopharyngeal meatus.
   • Note: Maintain asepsis throughout the collection.
   • Draw approximately 1ml of sampling fluid into a sterile pipette.
   • Note: Sampling fluid may be normal saline, phosphate-buffered saline, or trypticase soy broth.
5. Bronchial Aspirate:
   • Inject the sampling fluid slowly into the bronchi and the lung by caudally inserting the pipette into the trachea. Pipette out the sampling fluid from the bronchi and the lungs and remove the pipette from the trachea.
6. Nasal Aspirate:
   • Cranially direct and insert the pipette into the nasopharyngeal meatus (mice) or tracheal lumen (rats), and pour the sampling fluid into the nasal cavity.
   • Access the nasal cavity by contacting 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.
   • Draw the sampling fluid from the nasal cavity into the pipette and remove the pipette from the meatus.
   • Transfer the sample to an appropriate media or container for testing.
   •  Maintain asepsis throughout the procedure.

Dissection of Mouse Parotid Glands

1. Euthanize the mice using the cervical dislocation method.
2. Cut and open the skin of the chest and head/neck region from the epigastrium toward the limbs.
3. Open the thorax by making a trapezoid cut.
4. Make a small incision in the apex of the left ventricle, insert the perfusion cannula into the incision and gently push into the aortic arch.
5. Perfuse [4% paraformaldehyde (PFA) in PBS with RNase-free water at pH 7.4] for 2–3 min.
6. Lift the parotid glands with a pair of forceps to locate the borders of the organ and cut out, including a part of the main excretory duct.
7. Separate the parotid glands from the surrounding tissues.