Encyclopedia of Experiments: Immunology
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Placing two pairs of forceps on either side of the underlying mesentery between the blood vessels, gently rub the tips of the forceps together to create a hole in the membrane. Using angled forceps and a curved, taper point needle attached to a 5-centimeter suture, penetrate one side of the peritoneum through the hole in the membrane and up through the other side of the peritoneal lining, placing one suture at the top and another near the bottom of the incision to close the incision while keeping the loop of intestine externalized.
To increase the likelihood of success, do not tie off or tear any blood vessels while placing the abdominal structure, and limit any extraneous handling of the intestine.
Then, close the skin beneath the intestinal loop in the same manner, placing one suture in the middle of the incision between the previous sutures in the underlying peritoneum. Using an electrocautery, make a line of perforation along the anti-mesenteric border, and immediately apply a few drops of water to the surface of the intestine to prevent additional heat-induced tissue damage.
Blot with a Kimwipe to remove residual water. Use Vannas scissors to cut an approximately 1.5-centimeter horizontal slit at the distal edge of the cauterized tissue, along the length of the cauterized line toward the proximal end of the externalized tissue segment. When the mucosal surface is exposed, cover the abdomen with a moist Kimwipe to keep the tissue hydrated.
For spinning disk confocal microscopy imaging, transport the mouse to the microscope in a covered vessel. Launch the imaging software, and tilt the head of the microscope back, and add 150 microliters of 1 micromolar fleet AlexaFluor dye in Hank's buffered saline solution onto the glass coverslipped bottom. Position the mouse so that the opened mucosal surface directly contacts the coverslip.
Place the mouse and the dish onto the imaging stage in a pre-warmed incubator. Set the excitation intensity and exposure time for each laser to no more than 10 to 15 milliwatts, and 120 to 150 milliseconds respectively, and adjust the frame average to "2".
Turn on the electron-multiplying gain function to reduce the background noise, and select the 63X objective calibration to ensure a correct measurement of the pixel size. Using the 405-nanometer laser and the 20x air objective, manually visualize the nuclei to locate a field of villi that lack noticeable movement or drift, avoiding areas of artifactual movement due to respiration, peristalsis, or heartbeat.
Using the XY scan, record the x-y-coordinate of the field of interest, and switch to the glycerol immersion 63X objective. Acquire a live image on the 405-nanometer channel for up to 1 minute to confirm that the villi in the selected field are stable, while adjusting the focus to find the orthogonal plane just beneath the villous tip.
Then, acquire Z-stacks starting from just below the villous tip epithelium, about 15 to 20 micrometers, down to the villus, until it is difficult to resolve the nuclei using 1.5-micrometer steps. 3 to 5 minutes after beginning the acquisition, confirm the image stability, and the gamma-delta intraepithelial cell motility, and continue acquiring images for 30 to 60 minutes for each field of villi.
