The experimental protocol was approved by the Institutional Animal Care and Use Committee of Jianghan University and was conducted in accordance with Experimental Animals Ethical Guidelines issued by the Center for Disease Control of China. Adult male C57BL/6J mice, 10 weeks old, weighted 24-26 g, were used in this protocol. All mice were housed under a 12-h light/dark cycle controlled environment with food and water ad libitum.

1. Preoperative preparation

NOTE: The key instruments and equipment required for this protocol are shown in Figure 1.

1. Prior to the procedure, sterilize the surgical instruments by autoclaving.
2. Administer meloxicam subcutaneously at a dose of 5 mg/kg for analgesia 60 min before surgery.
3. Anesthetize the mouse with 5% isoflurane in an induction chamber.
4. Wipe the heating surgery board with 75% ethanol and activate the heating switch with the temperature set to 37 °C. Then, position the mouse in a left lateral position on the board, with the head positioned away from the surgeon and the tail facing towards the surgeon.
5. Attach a mask to the mouse face that provides a constant supply of 2% isoflurane and 0.3 L/min oxygen for maintenance anesthesia. Apply the eye ointment on the cornea to protect them from drying during the procedure.
6. Shave the fur from the left orbit to the left ear canal with a shaver. Use depilatory creams to remove the remaining fur for sterilization.
7. Prepare the surgical area by applying povidone-iodine disinfectant and 75% ethanol three times.

2. Distal MCAO model

1. Check if the mouse is deeply anesthetized by assessing the reflexes of the toe pinch.
2. Make a vertical incision between the left orbit and the left ear canal using a sterile scalpel blade.
3. Retract the skin's soft tissue with retractors and expose the temporal muscle.
4. Use straight microforceps to separate the apical and dorsal segments of the temporal muscle from the skull.
5. Identify the bifurcation of the MCA, which assumes a "Y" shape beneath the temporal bone with a bias towards the base of the skull (Figure 2A).
   NOTE: If the bifurcation of MCA is not visually discernible (due to an anatomically normal variation), ascertain the most rostral vessel.
6. Use an electric cranial drill to thin out the skull to make a bone window (4 mm in diameter) until the dura mater, with its translucent texture, becomes visible.
   NOTE: Intermittently add drops of normal saline to the skull to prevent overheating during this process.
7. Remove the dura mater above the artery with curved micro forceps to expose the MCA branch.
8. Set the electrocautery to 7 W. Coagulate the artery using the electric coagulation forceps at the sites of proximal and distal to the bifurcation of MCA (Figure 2A). When the bifurcation is not visible due to an anatomical variation, perform the coagulation on the accurately identified MCA branch (as mentioned above) at two separate locations approximately 1 mm apart¹³.
9. Monitor the blood flow of the left MCA cortical branch utilizing a laser speckle blood flow meter.
   NOTE: Mice exhibiting a blood flow reduction of less than 40% of the baseline value following electrocoagulation were excluded from the study.
10. Suture the muscle and the skin separately using 5-0 polyglycolic acid sutures. Apply diclofenac sodium gel and mupirocin ointment to the skin incision using a sterile cotton swab.
11. Place the mouse in a recovery chamber. Monitor all mice until they are fully awake. Provide meloxicam (5 mg/kg, SC) for analgesia every 24 h up to 72 h.

3. Sham Operation

1. Reproduce the same procedure as those mentioned above, excluding the process of arterial coagulation.

4. Behavioral tests

NOTE: Prior to the dMCAO, the mice underwent behavioral training twice daily for 3 days. On 3^rd day post-dMCAO, move the mice to the behavioral testing room for a 2 h environmental adaptation before testing.

1. Grip strength test¹⁴
   ​NOTE: The grip strength meter comprises a push-pull strain gauge and a metal horizontal bar.
   1. Grasp the base of the mouse tail. Gradually lower the mouse toward the metal grip bar until it grips the horizontal bar with both forepaws (Figure 2B).
   2. Pull the mouse backward at a constant speed of about 2 cm/s and keep its body horizontal.
   3. Record the peak force in grams (g) when the mouse releases its forepaws from the bar.
2. Pole test¹⁵
   1. Place the mouse on the top of a vertical wooden pole (50 cm high, 1 cm in diameter) with their head facing upwards, and allow them to climb down the pole (Figure 2B).
   2. Record the time taken by the mouse to turn around (T-turn) and the total time to climb down the pole (T-total) with a 60-s cut-off time.
3. Adhesive tape removal test¹⁶
   1. Hold the mouse and attach a patch of adhesive tape (2 × 2 mm) to the right forepaw of the mouse (Figure 2B).
   2. Put the mouse back into the rearing cage and allow it to move freely.
   3. Record the time taken by the mouse to remove adhesive with a limit of 60 s.
4. Cylinder test¹⁷
   1. Place the mouse in an open-top, clear glass cylinder (diameter: 14 cm; height: 20 cm) and record its spontaneous standing exploratory behavior for 3 min using a camera (Figure 2B).
      NOTE: During the experimental procedure, two mirrors were positioned alongside the cylinder to ensure optimal recording of forelimb movements.
   2. Wipe the cylinder with 75% ethanol to eliminate residual olfactory cues before testing the subsequent mouse.
   3. Reduce the playback speed of the video to 1/5 of the actual speed, and count the number of wall touches with the left (L) or right forepaw (R) or simultaneous use of both forepaws (B). Calculate the usage rate of the right forepaw as (L-R)/(L + R + B) × 100%.

5. Perfusion and sample preparation

1. Euthanize the mouse by peritoneal injection overdose of pentobarbital (300 mg/kg). Position the mouse supine and immobilize the limbs.
2. Make a midline incision on the skin, starting from the mid-abdominal cavity and extending towards the upper chest region.
3. Open the chest using surgical scissors and fold the xiphoid upwards using a hemostat. Carefully remove the lung and the diaphragm to expose the heart.
4. Insert the perfusion needle tip into the left ventricle and puncture the right atrium with scissors. Perfuse the heart with 20 mL of normal saline via the left ventricle.
5. Decapitate the mouse and cut open the scalp along the midline between the eyes. Cut the bone along both sides, starting from the skull base and opening up to the eye socket.
6. Lift the calvarium and gently detach the brain from the base of the skull by using a forceps.
   NOTE: Brains from half of the mice were subjected to 2,3,5-triphenyl tetrazolium chloride (TTC) staining, while brains from the other half underwent histological examination.
7. Soak the brains in 4% paraformaldehyde solution overnight for follow-up histological analysis.

6. Identification of infarction volume

1. Place the brain in a -20 °C freezer compartment of the refrigerator for 20 min.
2. Position the brain into the 1 mm brain matrix and slice the brain into coronary sections with a thickness of 2 mm by using microtome blades.
3. Transfer the brain sections to a 24-well culture plate and add 2% TTC to cover the brain tissue. Put the 24-well culture plate in a constant temperature oven for 30 min with the temperature set at 37 °C.
4. Carefully lift out the brain sections and fix them in 4% paraformaldehyde solution for 15 min.
5. Perform optical scanning of these brain sections with a resolution of 1200 x 1200 dpi by a scanister.
6. Measure the cerebral infarct volume by summing the infarct area of each section (area of the right hemisphere minus uninjured area of the left hemisphere) using Image J software (https://imagej.net/software/imagej/index)¹⁸. Calculate the infarct volume percentage as infarct volume/volume of the right hemisphere × 100%.

7. Histopathological and immunofluorescent staining analysis

1. Take the brain samples from the paraformaldehyde solution and flush them with running water for 1 h.
2. Transfer the brain samples into the automation-tissue-dehydrating machine. Launch the preset program for dehydration, vitrification, and wax immersion. Embed the dehydrated brain samples in paraffin and slice them into 5-µm thick sections by a microtome.
3. Hematoxylin and eosin (H&E) staining
   1. Dewax the sections with xylene 3 times for 8 min each.
   2. Rehydratethe sections by successively immersing in gradient ethanol (100%, 95%, 90%, 80%, 70%) and distilled water for 5 min each time.
   3. Stain the sections with the hematoxylin solution for 5 min.
   4. Rinse the sections with distilled water to wash the residue hematoxylin, then dip them in 5% hydrochloric acid alcohol for a 5-s differentiation. Wash the sections with distilled water for 2 min.
   5. Stain the sections with eosin solution for 30 s and rinse off excess staining fluid using distilled water.
   6. Immerse the sections in 90% ethanol, 95% ethanol, 100% ethanol, and xylene (2 times) successively for 5 min each time. Mount the sections with a neutral balsam mounting medium.
4. Immunofluorescence staining
   1. Dewax the sections with xylene 3 times for 8 min each time.
   2. Rehydrate the sections by successively immersing in gradient ethanol (100%, 95%, 90%, 80%, 70%) and distilled water for 5 min each time.
   3. Preheat the citrate antigen retrieval solution to boil in the microwave. Immerse the sections in the antigen retrieval solution. Reheat the antigen retrieval buffer with low power (20 W) for 20 min.
   4. Turn off the microwave and let the antigen retrieval buffer cool to room temperature (RT).
   5. Wash the sections thrice with 0.1 mM phosphate buffered saline (PBS) for 5 min each time.
   6. Incubate the sections with a blocking solution (5% bovine serum albumin) at RT for 1 h to block the non-specific binding of immunoglobulin.
   7. Incubate the sections with rabbit anti-NeuN antibody (1: 300), rabbit anti-Iba-1 antibody (1: 500) and mouse anti-GFAP antibody (1: 500) at 4˚C overnight.
   8. Wash the sections thrice with 0.1 mM PBS for 5 min each time.
   9. Incubate the sections with goat anti-rabbit Alexa 594-conjugated IgG (1:500) or goat anti-mouse Alexa 488-conjugated IgG (1:500) for 1 h at RT.
   10. Wash the sections with 0.1 mM PBS and mount them with an antifade mounting media containing 4',6-diamidino-2-phenylindole (DAPI).
   11. Take images of three microscopic fields (300 µm × 300 µm) within the ischemic penumbra using a fluorescent microscope at 594 nm and 488 nm, respectively.
   12. Using the ImageJ software (https://imagej.net/software/imagej/index), estimate the positively stained cell density by averaging counts in ischemic penumbra¹⁹.