Intravital Visualization of Gamma Delta Intraepithelial Lymphocytes

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Mouse Preparation

NOTE: The following procedure, including animal preparation and surgery, will take 30–40 min. Prior to the surgery, turn on the microscope and warm up the enclosed incubator on the microscope to 37 °C.

1. Perform experiments on 8–10 weeks old TcrdEGFP mice on a C57BL/6 background, which were obtained from Bernard Malissen (INSERM, Paris, France). According to the IACUC-approved procedure, anesthetize the mouse by i.p. injection of ketamine (120 mg/kg) and xylazine (10 mg/kg) based on the weight of the animal. Evaluate the level of anesthesia by respiration rate (55–65 breaths per minute), toe pinch, and palpebral reflex.
2. Once surgical plane anesthesia is reached, secure the hind legs of the mouse using tape to a heating pad to maintain body temperature, which can be monitored by a rectal thermometer. If there are signs that the anesthesia is wearing off (e.g., increased respiration rate, blinking and/or eye muscle twitching response to palpebral reflex), re-administer 50 μL of ketamine/xylazine at a time until the mouse is fully sedated.
3. Prepare the nuclear dye by diluting 50 µL of Hoechst 33342 (10 mg/mL) with 315 µL of 0.9% saline; the final concentration is 37.5 mg/kg based on the weight of the mouse (200 µL approximate volume). Once the mouse is anesthetized, slowly inject the Hoechst using a tuberculin syringe through the retro-orbital venous sinus.
   NOTE: Wait 1–2 min before proceeding to the next step to allow the mouse to acclimate to the change in circulating volume following retro-orbital injection.
4. Place a small amount of lubricant over the eyes to prevent them from drying out.

2. Mouse Surgery: Laparotomy to Expose Intestinal Mucosa

1. Make a 2 cm vertical incision through the skin and peritoneum along the midline of the lower abdomen to externalize the region of the intestine to be imaged.
2. Using angled forceps, carefully pull the cecum out of the peritoneal cavity and identify the area of the small intestine to be imaged. Be careful not to tear mesenteric blood vessels during this process. To minimize potential damage or bleeding, avoid grabbing or pinching the intestine with forceps.
3. Locate a suitable region of the small intestine (2–3 cm) for imaging that contains minimal fecal contents. Carefully return the cecum and remaining small intestine into the peritoneal cavity, while leaving the segment of interest externalized.
   NOTE: Avoid puncturing the cecum or disrupting the mesenteric vasculature during this step.
4. Place two pairs of forceps on either side of the underlying mesentery between blood vessels, and gently rub the tips of the forceps together to create a hole in the membrane (Figure 1). This will allow the needle to pass through the mesentery when closing the peritoneal cavity beneath the intestine.
5. Close the incision while keeping the loop of the intestine externalized. Using angled forceps and a curved, taper point needle attached to a 5 cm suture, penetrate one side of the peritoneum, go through the hole made in the previous step, and up through the other side of the peritoneal lining. Place one suture at the top, and another near the bottom of the incision.         
   NOTE: Avoid damaging the vasculature during this step.
6. Repeat this process to close the skin beneath the intestinal loop, by placing one suture in the middle of the incision, between the previous sutures in the underlying peritoneum.
   NOTE: It is important to only externalize the area that is to be imaged; this will reduce extraneous motion during imaging. Be sure to avoid tying off the mesenteric arteries.
7. Use an electrocautery to make a line of perforation along the anti-mesenteric border. Immediately after cauterization, 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.
8. Cut a small horizontal slit at the distal edge of the cauterized tissue using Vannas scissors and proceed to cut along the length of the cauterized line toward the proximal end of the externalized tissue segment (approximately 1.5 cm in length) to expose the mucosal surface.    
   NOTE: If necessary, use forceps to gently remove any excess fecal content remaining on the exposed mucosal surface.
9. Cover the abdomen of the mouse with a moist Kimwipe to prevent the tissue from becoming dehydrated. Transport the mouse to the microscope in a covered vessel.

3. Image Acquisition by Spinning Disk Confocal Microscopy

1. Pipette 150 µL of 1 µM AlexaFluor 633 in HBSS onto the glass coverslipped bottom of a 35 mm dish. Position the mouse so that the opened mucosal surface directly contacts the coverslip.
2. Tilt the head of the microscope back and place the mouse on the coverslipped dish on the imaging stage in a pre-warmed incubator. Alternatively, cover the mouse with a heating blanket to maintain body temperature.
3. Launch imaging software. The excitation intensity and exposure time for each laser should not exceed 10–15 mW or 120–150 ms, respectively, to avoid photobleaching and to minimize the interval between time points. Adjust the frame average to "2" and turn on the EM gain function to reduce background noise. Select 63x objective calibration to ensure the correct measurement of pixel size.
   NOTE: The settings detailed above are meant to provide an initial reference, but will vary depending on individual microscope configurations.
4. Using the 405 nm laser and the 20x air objective, visualize the nuclei to locate a field of villi that lack noticeable movement or drift by eye. Avoid areas of artifactual movement due to respiration, peristalsis, or heartbeat.
5. Using the XY scan, record the XY coordinate of the field of interest and switch to the glycerol immersion 63X objective.
6. Confirm that the villi in the selected field are stable by acquiring a live image on the 405 nm channel for up to 1 min.
7. While acquiring a live image on the 405 nm channel, adjust the focus to find the orthogonal plane just beneath the villous tip. Acquire Z-stacks starting from just below the villous tip epithelium down the villus until it is difficult to resolve the nuclei, about 15–20 µm, using a 1.5 µm step.        
   NOTE: When imaging with three channels (405, 488, 640 nm) using the exposure times described above, it is possible to acquire all three channels sequentially at approximately 20 s intervals.
8. Open the software in analysis mode, 3–5 min after beginning the acquisition, to confirm the image stability and γδ IEL motility. Continue to acquire images for 30–60 min for each field of villi. Record 2–3 fields for each mouse. While imaging, monitor the mouse every 5 min.