Postoperative Ileus Murine Model

Animal care and experimental procedures were conducted in accordance with the Guiding Principles in the Care and Use of Animals (China) and were approved by the Ethics Review Committee of Beijing Friendship Hospital (NO. 20-2056). C57BL/6 mice (8-12 weeks old) were used for the study.

1. Preparation for surgery

1. Fast all mice for 12 h prior to the planned modeling to prevent aspiration death caused by reflux of stomach contents after anesthesia (Figure 1A).
2. Prepare and sterilize surgical instruments.
3. Prepare the anesthetic tribromoethanol and the heating pad. Monitor the heating pad to prevent overheating and keep it at a consistent temperature (37.5 °C).

2. Anesthesia

1. Anesthetize each mouse with tribromoethanol.
   1. Dilute tribromoethanol to prepare a 20 mg/mL solution in normal saline. Administer 0.2 mL of the tribromoethanol working solution per 10 g of body weight via intraperitoneal injection.
   2. Ensure mice are completely anesthetized within 5 min and remain anesthetized for 20 min.
2. Assess the depth of anesthesia by observing the inability of the mouse to remain upright and check for muscle relaxation. Observe the loss of voluntary movement, blink reflex, and response to reflex stimulation (toe or tail pinch with firm pressure).
3. Assess the respiratory rate and pattern of the mouse by monitoring the movement of the chest wall and abdomen. Ensure the breathing rate is ~ 55-65 breaths/min under optimal anesthesia.
4. Place the anesthetized mouse on a board and secure it with tape.
5. After the mice were completely anesthetized, use an electric hair shaver to remove hair from the abdomen. After shaving, wipe off all loose hair on the abdomen using a saline-moistened cotton ball.
   NOTE: Ensure proper ventilation and use of a fume hood or biosafety cabinet when handling tribromoethanol to minimize exposure risks. During the procedure, wear appropriate personal protective equipment, including gloves and lab coats, to prevent accidental contact with chemicals and biological materials.

3. Surgery

1. Fully extend the limbs to expose the abdomen. Ensure that the mouse's head is positioned to maintain a clear airway (Figure 1B).
2. Disinfect the skin of the surgical area twice using a cotton ball soaked with 75% alcohol. After disinfection, use a dry, sterile medical gauze to remove excess alcohol from the abdomen.
3. Make an incision through the skin and lift the rectus abdominis muscle in the middle of the abdomen using tweezers.
   1. Make a small incision along the median line of the rectus abdominis muscle, taking care to avoid injury to various organs in the abdomen.
   2. Ensure the approximate range of the incision is as follows. Ensure that the upper margin of the incision is 6-8 mm from the xiphoid process of the sternum, the lower margin is 6-8 mm from the external genitalia, and the length of the incision is ~1 cm.
4. Place a piece of gauze pre-moistened with normal saline on both sides of the abdominal incision. Use hemostatic forceps to fix the gauze on the upper and lower edges of the incision, exposing the incision (Figure 1C).
   NOTE: In the sham group, the incision was covered with wet gauze for 5 min without any surgical operation.
5. Properly fix the gauze, then use two cotton swabs pre-moistened with saline to gently press against both sides of the abdominal wall adjacent to the incision. Squeeze out a small amount of the intestinal tube through the incision and expose it by placing it on the gauze (Figure 1D).
6. Completely moisten two cotton swabs with normal saline. Gently grasp the intestinal tissue with the moistened cotton swabs and carefully remove the small intestine.
   1. Locate the caecum and then take out the intestine with the cotton swabs until 2 cm before the stomach to avoid touching the pancreas. Extend the intestine from the proximal end of the pancreato-duodenal ligament to the distal end of the ileocecal region.
7. Apply consistent pressure along the entire small intestine from the proximal to the distal ends. Ensure uniform force application for 5 min until small bleeding spots emerge on the intestinal surface (Figure 1E).
8. After 5 min, use pre-moistened cotton swabs to carefully place all the small intestines back into the abdominal cavity, following the normal physiological anatomical position of the small intestine.
9. Gently massage the abdomen across the gauze and abdominal wall for 3-5 s to ensure that the bowel is restored to its natural anatomical position and to prevent artificial mechanical intestinal obstruction or mesenteric torsion after surgery.
10. Inject 100 µL of saline into the abdominal cavity to replace lost fluids during the operation and lubricate the abdominal tissue.
11. Remove the gauze and close the abdomen using 6-0 surgical sutures. Begin by closing the muscle layer with continuous sutures, taking care to avoid damage to the abdominal organs. Lift the rectus abdominis muscle during suturing (Figure 1F).
12. After completely closing the rectus abdominis muscle, suture it completely with simple intermittent sutures using a 6-0 surgical suture. Maintain a stitch length of ~0.5 mm and a needle spacing of ~2 mm.
13. After closing the abdominal incision, gently wipe the area near the incision with dry sterile medical gauze to keep it dry and clean from blood, tissue fluid, or normal saline.
14. Apply a small amount of medical incision adhesive to and around the incision to avoid splitting after surgery.
15. After the medical incision adhesive has completely dried, transfer the mice to a cage equipped with dry, cleaning bedding, and place the cage on a heated blanket maintained at a constant temperature of 37.5 °C until the mice fully recover. Do not leave the animals unattended until they have regained sufficient consciousness to maintain sternal recumbency.
16. Transfer the awake mice to the animal feeding barrier environment and allow them to drink freely until the detection time.

4. Gastrointestinal transit assay

1. Administrate 200 µL of FITC-dextran solution (5 mg/mL in PBS) to the fasted mice via gavage 22.5 h after surgery (Figure 1A).
2. Euthanize the mice using a method approved by the Institutional Animal Care and Use Committee (IACUC), such as CO₂ inhalation followed by cervical dislocation.
   1. To isolate the digestive tract, make a midline incision from the diaphragm to the pelvis, carefully removing the surrounding connective tissues and organs.
   2. Separate the digestive tract from the surrounding tissues and organs, ensuring that the entire tract from the stomach to the anus is included.
3. Divide the digestive tract into 15 equal segments (from stomach to colon), numbered sequentially from 1 to 15 based on physiological position (sequence: St -S1:S10-Ce-L1:L3) (Figure 2A).
4. Cut each segment of intestinal tissue with its content into pieces measuring 1-4 mm and place them into separate 1.5 mL centrifuge tubes containing 1 mL of DPBS.
5. Vortex the centrifuge tubes for 10 s to homogenize the samples.
6. Centrifuge the tubes at 500 × g for 1 min. Collect the supernatant for FITC fluorescence quantification using a multimode microplate reader.
7. Describe the gastrointestinal transit by calculating the geometric center (GC) of the FITC-dextran and calculate using the following formula: (Σ[% FITC per segment x segment number])/100.

5. Paraffin embedding and hematoxylin and eosin (HE) staining

1. Euthanize the mice by CO₂ inhalation followed by cervical dislocation after 24 h. Perform paraffin embedding and gastrointestinal assays on the same animal.
2. Open the abdominal cavity and collect small intestinal tissue for further analysis.
3. Fix the entire bowel with intestinal content in Carnoy's fixative (60% methanol + 10% acetic acid + 30% chloroform) at 4 °C for 2 h.
4. Deacidify the tissue by rinsing twice with methanol for 30 min each time.
5. Replace methanol with ethanol twice for 30 min each time.
6. Deacidify the tissue by rinsing twice with methanol for 30 min each time. To achieve tissue transparency with xylene, immerse the tissue in xylene and incubate for 1 h.
7. Embed the tissue in paraffin wax using a paraffin embedding machine. Cut the embedded tissue into 4 µm thick slices using a microtome.
   NOTE: Divide the small intestine into different waxes for improved performance. Properly label the segments to identify their location.
8. Perform HE staining using the HE staining kit. Seal the slides and examine them under a microscope for analysis.

6. Immune cell isolation and flow cytometry

1. Prepare the following solutions.
   1. Predigestion solution: Prepare Hank's balanced salt solution without Ca²⁺ and Mg²⁺ with 1 mM dithiothreitol (DTT) and 10 mM ethylenediaminetetraacetic acid (EDTA).
   2. Digestion solution: Prepare a solution containing 200 µg/mL DNAase I, 500 µg/mL collagenase IV, 4% FBS, and 100 µM HEPES buffer in RPMI 1640.
   3. Magnetic-activated cell sorting works (MACS) buffer: Prepare a buffer containing 5 g/L BSA and 2 mM EDTA in PBS without Ca²⁺ or Mg²⁺.
2. Remove fecal contents by washing the intestines with normal saline. Remove Peyer's patches and connective tissue of the small intestine. Cut the small intestines open longitudinally and then cut into 1 cm segments in ice-cold PBS.
3. Wash the intestine segments with predigestion solution at 37 °C for 20 min on an orbital shaker. Collect the predigestion solution and filter it through 70 µm cell strainers to obtain intraepithelial lymphocytes (IELs).
4. Digest the segments in the digestion solution for 30 min and then filter the suspension through 70 µm cell strainers to obtain lamina propria lymphocytes (LPLs).
5. Centrifuge the LPL cells at 500 × g for 10 min, and discard the supernatant.
6. Wash and suspend the cells with DPBS (sodium azide, Tris, and protein-free) at a concentration of 1 x 10⁶ cells/100 µL.
7. Incubate the cells with a fluorescent viability dye (1:500) for 15 min at room temperature (RT) in the dark.
8. Suspend the cells with MACS buffer and centrifuge them at 500 × g for 5 min.
9. Count the stained cells using a hemocytometer or automated cell counter. Resuspend the cells at a concentration of 1 x 10⁶ cells/100 µL of MACS buffer. Perform antibody staining (1:200-1:400) by adding the appropriate volume of antibody to the cell suspension and incubating at 4 °C for 15 min.
10. Wash the cells as described in step 6.8.
11. Detect the samples using flow cytometry and analyze data with the associated flow cytometry software^7,8.