Erratum: A Mouse Model of Hemorrhagic Transformation Induced by Acute Hyperglycemia Combined with Transient Focal Ischemia

Summary

An erratum was issued for: A Mouse Model of Hemorrhagic Transformation Induced by Acute Hyperglycemia Combined with Transient Focal Ischemia. The Protocol section was updated.

Abstract

This corrects the article 10.3791/67371

Protocol

An erratum was issued for: A Mouse Model of Hemorrhagic Transformation Induced by Acute Hyperglycemia Combined with Transient Focal Ischemia. The Protocol section was updated from:

from:

The experimental protocol was approved by the Institutional Animal Care and Use Committee of Jianghan University (JHDXLL2024-080) and conducted in accordance with the Experimental Animal Ethical Guidelines issued by the Center for Disease Control of China. Adult male C57BL/6J mice weighing 21-26 g were used in this study. The details of the reagents and equipment used are listed in the Table of Materials.

1. Animal grouping and acute hyperglycemia inducing

1. House the mice in the laboratory animal center of Jianghan University in a 12-h light/dark cycle controlled environment (20-22 °C, 50%-55% humidity) with food and water ad libitum.
2. Randomly divide the mice into four groups: Sham + saline (n = 12), Sham + glucose (n = 12), MCAO + saline (n = 12), and MCAO + glucose (n = 20).

2. Preoperative preparation

NOTE: All experimental mice fasted for 12 h before surgery.

1. Administer systemic analgesia (meloxicam, 5 mg/kg, subcutaneous) and local anesthesia to the incision site (bupivacaine, 2 mg/kg, subcutaneous) perioperatively.
2. Sterilize all surgical instruments, cotton swabs, and surgical sutures by autoclaving at the conditions of 121 °C and 15 psi for 30 min.
3. Wipe the surgical platform and surrounding work area with 75% ethanol, and turn on the heating switch with the temperature set to 37 °C.
4. Anesthetize the mouse with a 3% isoflurane-20% oxygen gas mixture in the anesthesia induction chamber. Then, evaluate the toe pinch reflex to test the depth of anesthesia.
5. Move the mouse on the surgical platform in the prone position, and quickly insert the mouse's nose in the nose cone. Open the gas flow to the nose cone, closing off flow to the anesthesia induction chamber. Adjust the proportion of isoflurane to 2% and set the flow of the gas mixture to 0.4 mL/min for anesthesia maintenance.
6. Apply eye gel to the mouse's eyes to maintain moisture during surgery.
7. Shave the hair of the neck region by using the animal clipper. Clean and disinfect the skin by alternating applying with iodine and 70% ethanol using a disinfectant cotton stick.
8. Cover a sterile gauze on the neck and cut an opening to expose the operative region.

3. MCAO surgical procedure

NOTE: MCAO is performed using a modified thread-occlusion method, as previously described by Chiang et al.¹³.

1. Administer intraperitoneally 30% glucose (7.2 mL/kg) or normal saline to the mice 15 min before the surgical procedure.
   NOTE: Mice from Sham + glucose and MCAO + glucose groups received intraperitoneal injections of glucose. Mice from Sham + saline and MCAO + saline groups received intraperitoneal injections of normal saline.
2. Make a horizontal incision with an opening of 1.5 cm along the midline of the neck using a scalpel.
   NOTE: Ensure the incision is kept straight to achieve optimal visualization of the carotid artery.
3. Pull apart the subcutaneous tissue and superficial fascia by using surgical tweezers.
4. Beneath the superficial fascia, locate the submandibular gland and an inverted triangle formed by three muscles: the sternohyoid, positioned at the midline over the trachea; the posterior belly of the digastric muscle, recognizable by its shiny white tendinous part; and finally, the sternomastoid muscle.
5. Perform blunt dissection within the inverted triangle to identify and separate the left common carotid artery (CCA).
6. Dissect the vagus nerve adjacent to the CCA and mucosal tissue surrounding the blood vessels using ophthalmic forceps. Make a ringer on CCA using a 5.0 polyglycolic acid (PGA) absorbable suture, but keep the suture untensioned.
7. Separate upward along the CCA, a "Y" bifurcation can be observed to divide the CCA into the external carotid artery (ECA) and internal carotid artery (ICA).
8. Separate the bifurcation to fully expose the ECA and ICA. Loop and tightly tie a 5.0 PGA absorbable suture around the ECA distally from the bifurcation.
   NOTE: Ensure a sufficient distance between the permanent ligation site and vascular bifurcation to prevent the thread from dislodging during the rotation of the ECA.
9. Temporarily clamp both the CCA and ICA using two 8 mm x 2 mm light micro serrafine arterial artery clamps to block the blood flow from the CCA to the ICA.
10. Stretch the suture tie on the ECA distal and CCA to straighten the ECA segment.
11. Use microscissors to make a small incision between the two suture ties on the ECA.
12. Insert a silicon-coated monofilament suture (30 mm long, 3-4 mm coated silicon) into the ECA. Loop and slightly tie a second 5.0 PGA absorbable suture on the ECA near the bifurcation to prevent the monofilament suture from backing out.
13. Completely cut the ECA distal to the permanent ligation and remove the artery forceps clamp from the ICA.
14. Withdraw the suture to the bifurcation of the CCA, then carefully retract and rotate the ECA stump. Adjust the insertion direction of the suture and slowly insert it, approximately 9.0-10.0 mm from the CCA bifurcation into the ICA, to occlude the MCA.
15. Tighten the second PGA absorbable suture around the ECA and remove the artery clamp from the CCA.
16. Suture the muscle and the skin separately using 5.0 PGAabsorbable suture. Apply diclofenac sodium gel and mupirocin ointment to the wound.
17. Turn on the heating switch of the recovery cage with the temperature set to 37 °C. Place the mouse in the recovery cage during the post-occlusion period (60 min).

4. Monofilament removal and reperfusion

1. Re-anesthetize the mouse as previously described just before the occlusion period should end.
2. Clamp the CCA using a microclip artery clamp.
3. Partially retract the monofilament from the ICA until the silicon-coated tip becomes visible through the ICA.
4. Place another microclip artery clamp on the ICA above the silicon-coated tip.
5. Completely withdraw the monofilament and tightly ligate the ECA stump.
6. Remove the microclip artery clamp from ICA and CCA, respectively.
7. Suture the muscle and the skin layer by layer using 5.0 PGA absorbable suture.
8. Place the mouse in the recovery chamber and monitor all mice until they are fully awake. Administer meloxicam (5 mg/kg, subcutaneously) for pain relief every 12 h for up to 24 h.

5. Blood glucose measurement

NOTE: Blood glucose levels were measured at the following time points: (1) just before MCAO surgery (baseline), (2) immediately after the insertion of the monofilament (15 min after glucose injection), (3) immediately after the withdrawal of the monofilament immediately after the insertion of the monofilament (75 min after glucose injection).

1. Wipe the mouse' tail with an alcoholic cotton ball to make the tail vein fully hyperaemic.
2. Cut off the tail tip by 1-2 mm using surgical scissors.
3. Gently squeeze along the root of the tail to the tip of the tail, facilitating the blood to flow out of the incision.
4. Position the sample absorption tank of the test paper at the edge of the blood droplet.
   NOTE: The blood will be drawn into the test paper due to the siphoning effect.
5. Read the blood glucose meter reading and record the result.
   NOTE: Exclude the mice whose blood glucose levels did not rise above 10 mmol/L after 15 min of glucose injection.
6. Remove any surplus blood using a cotton ball and apply pressure to halt the bleeding.

6. Cerebral blood flow measurement

1. Prior to MCAO, shave the hair of the scalp region by using the animal clipper. Clean and disinfect the scalp by alternating applying with iodine and 70% ethanol using a disinfectant cotton stick.
2. Make a 1.0 cm long midline incision in the skin over the frontal region to expose the cranial fontanelle. Keep the skull moist with normal saline.
3. Prepare the Laser Doppler Flowmetry (LDF) instrument to measure relative changes in cerebral blood flow (CBF).
4. Hold the LDF probe tip perpendicular to the surface of the left parietal skull (1 mm posterior and 5 mm lateral to the bregma) until a stable FLUX is read and record this value.
   NOTE: The unit of measurement for FLUX is Perfusion Units (PU), with a measurement range of 0-1000 PU. The ideal baseline CBF on a Laser Doppler Flowmetry is >600 PU. A continuous period of 5 s with a fluctuation range of less than 10% is considered a valid FLUX reading.
5. Monitor the CBF prior to the procedure and immediately following MCA occlusion.
   NOTE: Mice that showed a blood flow reduction of less than 40% of the baseline value after MCAO were excluded from the study¹⁴.

7. 2,3,5-Triphenyltetrazolium Chloride (TTC) staining

1. Euthanize the mouse by intraperitoneal injection of an overdose of pentobarbital (300 mg/kg).
2. Once the mouse stops breathing, promptly immobilize it with a supine position.
3. Open the thorax using scissors, and cut out the diaphragm to expose the heart.
4. Insert a 23 G injection needle into the left ventricle of the mouse and incise the auricular appendix with scissors.
5. Perfuse 15 mL of normal saline until the fluid draining from the auricular appendix appears clear and transparent.
6. Decapitate the mouse and cut open the scalp to fully expose the skull using scissors. Insert the scissors tip 2 mm slightly in front of the coronal suture of the skull to pry it open.
7. Use tweezers to remove the mouse's brain tissue intact.
8. Place the brain tissue in a rodent brain matrice and cut it into 2 mm coronal slices.
9. Transfer the slices to a 24-well plate and incubate them with 2% TTC solution at 37 °C for 15 min.
10. Start the scanning software and set the parameter settings to a resolution of 1200 dpi and a JPG format for image analysis.
11. Clip the tissue sections out from the 24-well plate with curved forceps and arrange the sections on the glass scanning plate.
12. Scan the tissue sections and then export the images for measurement analysis.
13. Measure the infarct area by correcting for tissue swelling. Subtract the non-infarct area of the ipsilateral side from the area of the contralateral side using ImageJ software¹⁵. Assess the infarct size as a percentage of the contralateral hemisphere. For additional details on infarct area analysis, refer to Friedländer et al.¹⁶.

8. Gross observation

1. Repeat steps 7.1-7.8.
2. Clip the tissue sections out from the brain matrice with curved forceps and arrange the sections on the glass scanning plate.
3. Scanthe brain coronal sections as described in steps 7.10 and 7.12.

9. Hematoxylin and eosin (H&E) staining

1. Repeat steps 7.1-7.4. Slowly perfuse 15 mL of normal saline, followed by perfusion of approximately 15 mL of paraformaldehyde (PFA).
2. Repeat steps 7.6-7.7. Fix the brain tissue with 4% PFA for 24 h at room temperature.
3. Place the fixed brain tissue into the automated tissue dehydrator for dehydration, vitrification, and wax immersion.
4. Embed the tissue in paraffin wax and cut the brain tissue into 5 µm thick coronal slices by microtome.
5. Dewax the slice by immersing it in xylene three times, with each immersion lasting 8 min. Then, gradually immerse the slice in ethanol solutions of decreasing concentrations (100%, 95%, 90%, 80%, 70%), followed by distilled water, with each step lasting 5 min.
6. Stain the slice with hematoxylin solution for 3 min. Then, differentiate it by immersing it in 5% hydrochloric acid alcohol for 5 s.
7. Incubate the slice with eosin solution for 1 min, then dehydrate and hyalinize it with gradient ethanol solution (90%, 95%, 100%) and xylene.
8. Mount the section with neutral resins.
9. Capture the images of H&E stained brain slices under a bright field using fluorescence microscopy.

10. Determination of Evans Blue (EB) lekage

NOTE: For details on this procedure, please refer to Wang et al.¹⁷.

1. Inject 2% (w/v) Evans Blue (EB) solution (2 mL/kg) via the tail vein 23 h post-MCAO.
2. Repeat steps 7.1-7.7 one hour after EB injection, and capture a photo of the entire brain.
3. Place the brain in a rodent brain matrice and cut it coronally into 2 mm slices. Scanthe brain coronal sections as described in steps 7.10-7.12 to observe EB leakage.
4. Divide the slices into right and left portions, and put each one into a centrifuge tube.
5. Homogenize every slice of brain tissue in 50% trichloroacetic acid, with a ratio of 100 mg of brain tissue to 0.4 mL of trichloroacetic acid.
6. After being incubated at 4 °C overnight, centrifuge the homogenate for 30 min at 10,000 x g (at 4 °C).
7. Transfer the supernatant liquid into another centrifuge tube and dilute it four-fold with ethanol.
8. Measure the absorbance of the supernatant liquid at 620 nm using a spectrophotometer.
9. Convert absorbance values to the concentration of EB using a standard curve of EB in ethanol (e.g., at 31.25, 62.5, 125, and 250 ug/mL).
   NOTE: The result is presented as a microgram of EB per gram of brain tissue.

11. Determination of FITC-Dextran leakage

1. Inject FITC-Dextran (10 KDa, 6 mg /mL, diluted in 0.01M PBS; 4 mL/kg) into the tail vein 24 h post-MCAO, allowing it to circulate in the blood for 10 min.
2. Repeat steps 7.1-7.7 to perform perfusion and remove the intact brain tissue.
3. Fix the brain tissue in 4% PFA for 24 h at room temperature in darkness.
4. Transfer the fixed brain tissue into the sucrose solution with gradient concentrations of 10%, 20%, and 30% (diluted in 0.01M PBS) for dehydration.
5. Embed the brain tissue using optimal cutting temperature compound (OCT) and cut into 30 µm thick brain tissue sections. Transfer the sections onto a microscope slide using an inoculating loop and ensure they adhere to the surface of the slide.
6. Mount the brain tissue sections using a mounting medium containing 4',6-diamidino-2-phenylindole (DAPI).
7. Start the Laser confocal microscopy and its control software.
8. Fix the microscope slide on the object stage and locate the infarct area under 200x eyepiece.
9. Set the resolution to 1024 x 1024 and adjust the gain value and exposure time to obtain the clearest image possible.
10. Acquire the images of the FITC-Dextran stained region of cerebral infarction under an excitation wavelength of 488 nm.

to:

The experimental protocol was approved by the Institutional Animal Care and Use Committee of Jianghan University (JHDXLL2024-080) and conducted in accordance with the Experimental Animal Ethical Guidelines issued by the Center for Disease Control of China. Adult male C57BL/6J mice weighing 21–26 g were used in this study. The details of the reagents and equipment used are listed in the Table of Materials.

1. Animal grouping and acute hyperglycemia inducing

1. House the mice in the laboratory animal center of Jianghan University in a 12-h light/dark cycle controlled environment (20–22 °C, 50%–55% humidity) with food and water ad libitum.
2. Randomly divide the mice into four groups: Sham + saline (n = 12), Sham + glucose (n = 12), MCAO + saline (n = 12), and MCAO + glucose (n = 20).

2. Preoperative preparation

NOTE: All experimental mice fasted for 12 h before surgery.

1. Administer systemic analgesia (meloxicam, 5 mg/kg subcutaneous) and local anesthesia to the incision site (bupivacaine, 2 mg/kg subcutaneous) peri-operatively.
2. Sterilize all surgical instruments, cotton swabs, and surgical sutures by autoclaving at the conditions of 121 °C and 15 psi for 30 min.
3. Wipe the surgical platform and surrounding work area with 75% ethanol, and turn on the heating switch with the temperature set to 37 °C.
4. Anesthetize the mouse with a 3% isoflurane-20% oxygen gas mixture in the anesthesia induction chamber. Then, evaluate the toe pinch reflex to test the depth of anesthesia.
5. Move the mouse on the surgical platform in the prone position, and quickly insert the mouse’s nose in the nose cone. Open the gas flow to the nose cone, closing off flow to the anesthesia induction chamber. Adjust the proportion of isoflurane to 2% and set the flow of the gas mixture to 0.4 mL/min for anesthesia maintenance.
6. Apply eye gel to the mouse's eyes to maintain moisture during surgery.
7. Shave the hair of the neck region by using the animal clipper. Clean and disinfect the skin by alternating applying with iodine and 70% ethanol using a disinfectant cotton stick.
8. Cover a sterile gauze on the neck and cut an opening to expose the operative region.

3. Baseline cerebral blood flow measurement

1. Shave the hair of scalp region by using the animal clipper. Clean and disinfect the scalp by alternating applying with iodine and 70% ethanol using a disinfectant cotton stick.
2. Make a 1.0 cm long midline incision in the skin over the frontal region to expose the cranial fontanelle. Keep the skull moist with normal saline.
3. Prepare the Laser Doppler Flowmetry (LDF) instrument and hold the LDF probe tip perpendicular to surface of the left parietal skull (1 mm posterior and 5 mm lateral to the bregma); once the FLUX is stable and record the baseline value.
   NOTE: The unit of measurement for FLUX is Perfusion Units (PU), with a measurement range of 0-1000 PU. Ideal baseline CBF on a Laser Doppler Flowmetry are >600 PU. A continuous period of 5 seconds with a fluctuation range of less than 10% is considered as valid FLUX reading.
4. Suture the muscle and the skin separately using 5.0 PGA absorbable suture. Apply diclofenac sodium gel and mupirocin ointment to the wound.

4. MCAO surgical procedure

NOTE: MCAO is performed using a modified thread-occlusion method, as previously described by Chiang et al.¹³.

1. Gently turn the mouse over to a supine position and administer intraperitoneally 30% glucose (7.2 mL/kg) or normal saline to the mice 15 min before the surgical procedure.
   ​NOTE: Mice from Sham + glucose and MCAO + glucose groups received intraperitoneal injections of glucose. Mice from Sham + saline and MCAO + saline groups received intraperitoneal injections of normal saline.
2. Shave the hair of neck region by using the animal clipper. Clean and disinfect the skin by alternating applying with iodine and 70% ethanol using a disinfectant cotton stick. Cover a sterile gauze on the neck and cut an opening to expose the operative region.
   NOTE: Ensure the incision is kept straight to achieve optimal visualization of the carotid artery.
3. Make a 1.5 cm midline incision on the ventral aspect of the neck using a scalpel. Pull apart the subcutaneous tissue and superficial fascia by using surgical tweezers.
   NOTE: Make sure that the incision is kept straight to achieve optimal visualization of the carotid artery.
4. Beneath the superficial fascia, locate the submandibular gland and an inverted triangle formed by three muscles: the sternohyoid, positioned at the midline over the trachea; the posterior belly of the digastric muscle, recognizable by its shiny white tendinous part; and finally, the sternomastoid muscle.
5. Perform blunt dissection within the inverted triangle to identify and separate the left common carotid artery (CCA).
6. Dissect the vagus nerve adjacent to the CCA and mucosal tissue surrounding the blood vessels using ophthalmic forceps. Make a ringer on CCA using a 5.0 polyglycolic acid (PGA) absorbable suture, but keep the suture untensioned.
7. Separate upward along the CCA, a "Y" bifurcation can be observed to divide the CCA into the external carotid artery (ECA) and internal carotid artery (ICA).
8. Separate the bifurcation to fully expose the ECA and ICA. Loop and tightly tie a 5.0 PGA absorbable suture around the ECA distally from the bifurcation.
   NOTE: Ensure a sufficient distance between the permanent ligation site and vascular bifurcation to prevent the thread from dislodging during the rotation of the ECA.
9. Temporarily clamp both the CCA and ICA using two 8 mm x 2 mm light micro serrafine arterial artery clamps to block the blood flow from the CCA to the ICA.
10. Stretch the suture tie on the ECA distal and CCA to straighten the ECA segment.
11. Use microscissors to make a small incision between the two suture ties on the ECA.
12. Insert a silicon-coated monofilament suture (30 mm long, 3–4 mm coated silicon) into the ECA. Loop and slightly tie a second 5.0 PGA absorbable suture on the ECA near the bifurcation to prevent the monofilament suture from backing out.
13. Completely cut the ECA distal to the permanent ligation and remove the artery forceps clamp from the ICA.
14. Withdraw the suture to the bifurcation of the CCA, then carefully retract and rotate the ECA stump. Adjust the insertion direction of the suture and slowly insert it, approximately 9.0–10.0 mm from the CCA bifurcation into the ICA, to occlude the MCA.
15. Tighten the second PGA absorbable suture around the ECA and remove the artery clamp from the CCA.
16. Suture the muscle and the skin separately using 5.0 PGA absorbable suture. Apply diclofenac sodium gel and mupirocin ointment to the wound.
17. Gently turn the mouse over to the prone position and repeat step 3.4. to record the blood flow following MCA occlusion.
    NOTE: Mice exhibiting a blood flow reduction of less than 40% of the baseline value following MCAO were excluded from the study¹⁴.
18. Turn on the heating switch of recovery cage with the temperature set to 37°C. Place the mouse in the recovery cage during the post-occlusion period (60 min).

5. Monofilament removal and reperfusion

1. Re-anesthetize the mouse as previously described just before the occlusion period should end.
2. Clamp the CCA using a microclip artery clamp.
3. Partially retract the monofilament from the ICA until the silicon-coated tip becomes visible through the ICA.
4. Place another microclip artery clamp on the ICA above the silicon-coated tip.
5. Completely withdraw the monofilament and tightly ligate the ECA stump.
6. Remove the microclip artery clamp from ICA and CCA, respectively.
7. Suture the muscle and the skin layer by layer using 5.0 PGA absorbable suture.
8. Place the mouse in the recovery chamber and monitor all mice until they are fully awake. Administer meloxicam (5 mg/kg, subcutaneously) for pain relief every 12 h for up to 24 h.

6. Blood glucose measurement

NOTE: Blood glucose levels were measured at the following time points: (1) just before MCAO surgery (baseline), (2) immediately after the insertion of the monofilament (15 min after glucose injection), (3) immediately after the withdrawal of the monofilament immediately after the insertion of the monofilament (75 min after glucose injection).

1. Wipe the mouse’ tail with an alcoholic cotton ball to make the tail vein fully hyperaemic.
2. Cut off the tail tip by 1–2 mm using surgical scissors.
3. Gently squeeze along the root of the tail to the tip of the tail, facilitating the blood to flow out of the incision.
4. Position the sample absorption tank of the test paper at the edge of the blood droplet.
   NOTE: The blood will be drawn into the test paper due to the siphoning effect.
5. Read the blood glucose meter reading and record the result.
   NOTE: Exclude the mice whose blood glucose levels did not rise above 10 mmol/L after 15 min of glucose injection.
6. Remove any surplus blood using a cotton ball and apply pressure to halt the bleeding.

7. 2,3,5-Triphenyltetrazolium Chloride (TTC) staining

1. Euthanize the mouse by intraperitoneal injection of an overdose of pentobarbital (300 mg/kg).
2. Once the mouse stops breathing, promptly immobilize it with a supine position.
3. Open the thorax using scissors, and cut out the diaphragm to expose the heart.
4. Insert a 23 G injection needle into the left ventricle of the mouse and incise the auricular appendix with scissors.
5. Perfuse 15 mL of normal saline until the fluid draining from the auricular appendix appears clear and transparent.
6. Decapitate the mouse and cut open the scalp to fully expose the skull using scissors. Insert the scissors tip 2 mm slightly in front of the coronal suture of the skull to pry it open.
7. Use tweezers to remove the mouse's brain tissue intact.
8. Place the brain tissue in a rodent brain matrice and cut it into 2 mm coronal slices.
9. Transfer the slices to a 24-well plate and incubate them with 2% TTC solution at 37 °C for 15 min.
10. Start the scanning software and set the parameter settings to a resolution of 1200 dpi and a JPG format for image analysis.
11. Clip the tissue sections out from the 24-well plate with curved forceps and arrange the sections on the glass scanning plate.
12. Scan the tissue sections and then export the images for measurement analysis.
13. Measure the infarct area by correcting for tissue swelling. Subtract the non-infarct area of the ipsilateral side from the area of the contralateral side using ImageJ software¹⁵. Assess the infarct size as a percentage of the contralateral hemisphere. For additional details on infarct area analysis, refer to Friedländer et al.¹⁶.

8. Gross observation

1. Repeat steps 7.1–7.8.
2. Clip the tissue sections out from the brain matrice with curved forceps and arrange the sections on the glass scanning plate.
3. Scan the brain coronal sections as described in steps 7.10 and 7.12.

9. Hematoxylin and eosin (H&E) staining

1. Repeat steps 7.1–7.4. Slowly perfuse 15 mL of normal saline, followed by perfusion of approximately 15 mL of paraformaldehyde (PFA).
2. Repeat steps 7.6–7.7. Fix the brain tissue with 4% PFA for 24 h at room temperature.
3. Place the fixed brain tissue into the automated tissue dehydrator for dehydration, vitrification, and wax immersion.
4. Embed the tissue in paraffin wax and cut the brain tissue into 5 μm thick coronal slices by microtome.
5. Dewax the slice by immersing it in xylene three times, with each immersion lasting 8 min. Then, gradually immerse the slice in ethanol solutions of decreasing concentrations (100%, 95%, 90%, 80%, 70%), followed by distilled water, with each step lasting 5 min.
6. Stain the slice with hematoxylin solution for 3 min. Then, differentiate it by immersing it in 5% hydrochloric acid alcohol for 5 s.
7. Incubate the slice with eosin solution for 1 min, then dehydrate and hyalinize it with gradient ethanol solution (90%, 95%, 100%) and xylene.
8. Mount the section with neutral resins.
9. Capture the images of H&E stained brain slices under a bright field using fluorescence microscopy.

10. Determination of Evans Blue (EB) lekage

NOTE: For details on this procedure, please refer to Wang et al.¹⁷.

1. Inject 2% (w/v) Evans Blue (EB) solution (2 mL/kg) via the tail vein 23 h post-MCAO.
2. Repeat steps 7.1–7.7 one hour after EB injection, and capture a photo of the entire brain.
3. Place the brain in a rodent brain matrice and cut it coronally into 2 mm slices. Scan the brain coronal sections as described in steps 7.10–7.12 to observe EB leakage.
4. Divide the slices into right and left portions, and put each one into a centrifuge tube.
5. Homogenize every slice of brain tissue in 50% trichloroacetic acid, with a ratio of 100 mg of brain tissue to 0.4 mL of trichloroacetic acid.
6. After being incubated at 4 °C overnight, centrifuge the homogenate for 30 min at 10,000 x g (at 4 °C).
7. Transfer the supernatant liquid into another centrifuge tube and dilute it four-fold with ethanol.
8. Measure the absorbance of the supernatant liquid at 620 nm using a spectrophotometer.
9. Convert absorbance values to the concentration of EB using a standard curve of EB in ethanol (e.g., at 31.25, 62.5, 125, and 250 ug/mL).
   NOTE: The result is presented as a microgram of EB per gram of brain tissue.

11. Determination of FITC-Dextran leakage

1. Inject  FITC-Dextran  (10 KDa, 6 mg /mL, diluted in 0.01M PBS; 4 mL/kg) into the tail vein 24 h post-MCAO, allowing it to circulate in the blood for 10 min.
2. Repeat steps 7.1–7.7 to perform perfusion and remove the intact brain tissue.
3. Fix the brain tissue in 4% PFA for 24 h at room temperature in darkness.
4. Transfer the fixed brain tissue into the sucrose solution with gradient concentrations of 10%, 20%, and 30% (diluted in 0.01M PBS) for dehydration.
5. Embed the brain tissue using optimal cutting temperature compound (OCT) and cut into 30 μm thick brain tissue sections. Transfer the sections onto a microscope slide using an inoculating loop and ensure they adhere to the surface of the slide.
6. Mount the brain tissue sections using a mounting medium containing 4',6-diamidino-2-phenylindole (DAPI).
7. Start the Laser confocal microscopy and its control software.
8. Fix the microscope slide on the object stage and locate the infarct area under 200x eyepiece.
9. Set the resolution to 1024 x 1024 and adjust the gain value and exposure time to obtain the clearest image possible.
10. Acquire the images of the FITC-Dextran stained region of cerebral infarction under an excitation wavelength of 488 nm.