Isolation of Lymphocytes from a Mouse Peyer's Patch

Begin by placing a mouse in the supine position, and sterilizing the abdomen with 70% ethanol. Make a midline abdominal incision through the skin and peritoneum from the pubis to the rib cage to open the peritoneal cavity and locate the cecum and the ileocecal junction.

Make as distal as possible cut, at the junction to separate the small intestine from the cecum, and cut the mesentery with scissors to carefully remove the entire small intestine up to the pyloric sphincter.

Snip the junction between the pylorus and the duodenum to completely detach the small intestine from the abdominal cavity, and place the isolated small intestine into one well of a six-well plate containing cold RPMI medium, supplemented with 10% Fetal Bovine Serum or FBS on ice. Then, gently agitate the tissues manually until all of the segments are submerged in the cold medium.

When all of the intestines have been harvested, use forceps to grasp the mesenteric fat of one edge of the intestine and place the sample — mesenteric side down — on a paper towel. Moisten the entire intestinal segment with RPMI plus 10% FBS to avoid tissue dehydration and stickiness, and locate the Peyer's patches, which appear as white multilobulated aggregates with a cauliflower-like shape on the anti-mesenteric side of the intestinal wall.

To harvest the Peyer's patches, use curved surgical scissors with the curve facing up to gently excise each patch from its distal and proximal borders, taking care to exclude the surrounding intestinal tissue, and place the Peyer's patches into individual wells of a 12-well plate containing ice-cold RPMI plus 10% FBS, on ice, as they are collected.

When all of the patches have been acquired, use scissors to cut the tip of a 1-milliliter micropipette tip so the diameter is large enough to aspirate individual Peyer's patches, and transfer the Peyer's patches into 150-milliliter conical tube per mouse, containing 25-milliliters of 37 degrees Celsius RPMI plus 10% FBS.

Then, place the tubes in an orbital shaker at 37 degrees Celsius with continuous agitation at 125 to 150 RPM for 10 minutes. At the end of the agitation, transfer the Peyer's patches onto 140-micrometer cell strainer per mouse, and use the rubber end of 110-milliliter syringe plunger per animal to gently disrupt the Peyer's patches through the mesh into individual 50-milliliter conical tubes.

Rinse the strainers with 15 to 20 milliliters of cold RPMI plus 10% FBS, and collect the cells by centrifugation. After carefully discarding the supernatant, re-resuspend the cells at an approximately 1 x 10⁷ cells per milliliter of RPMI plus 10% FBS concentration for counting. Then, transfer 2 to 2.5 x 10⁶ cells per 200 micro-liters of medium, to each well of a 96-well round bottom plate, and pellet the cells by centrifugation.