Middle Cerebral Artery Occlusion Allowing Reperfusion via Common Carotid Artery Repair in Mice

This protocol and the data reported were conducted in accordance with the UK Animals (Scientific Procedures) Act, 1986 (Project license 60/4315) and following institutional ethical approval. All experiments are reported in accordance with the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guidelines¹⁶.

1. Preparation

1. Familiarize adult male C57BL/6 mice to post-operative care (e.g., environment, bedding, recovery foods) at least 48 h before the surgery.
   NOTE: Post-MCAO animals often have difficulty with eating and drinking following surgery.
   1. To prevent excessive weight loss following surgery, acclimatize the animals to any post-MCAO diet by, for example, placing rehydration gel, gel food, and soaked/wet normal diet pellets directly onto the cage floor.
   2. Change the cage bedding to post-operative bedding, such as white paper chip.
2. Sterilize all surgical tools before beginning the surgical set-up or procedures.
   1. Do this by autoclaving the tools (with a minimum of 121 °C, 15 psi, for 15 min) or through the use of ethylene oxide (following the correct manufacturer’s instructions, for 8–10 h).
   2. Disinfect all surfaces prior to setting up the procedure. Cover all surfaces with sterile surgical drapes or autoclaved foil for items that require handling during the surgery. Use aseptic technique for the duration of the procedure.
      NOTE: Autoclaved foil can be used to cover the instruments or equipment that need to be held during the surgery. Using sterile gloves and techniques, the foil can be applied and, then, the item can be placed into the sterile field. Following this, a sterile glove change will be required.

2. Middle Cerebral Artery Occlusion Surgery

1. Use adult male C57BL/6 mice weighing 24–31 g at the time of the surgery. Induce anesthesia using 5% isoflurane in 2 L/min of O₂, in a red plastic anesthesia chamber.
2. Following the induction, reduce the isoflurane to maintain an adequate anesthesia depth for the surgery (e.g., 1.5–2% in 70% N₂O₂/30% O₂), delivered by face mask combined with a scavenger system to lower the surgeon’s exposure to isoflurane.
   NOTE: Nitrous oxide (N₂O₂) can be used in combination with oxygen (O₂) to reduce the required amount of isoflurane for an adequate anesthesia depth.
3. Administer systemic analgesia pre-operatively [carprofen, 10 mg/kg subcutaneous (sc.)] and local anesthesia to the incision site (bupivacaine, 2 mg/kg sc., diluted 1:10 in 0.9% saline) peri-operatively.
4. Administer fluids by intraperitoneal (ip.) injections of 200 µL of sterile prewarmed 0.9% NaCl solution pre-operatively.
5. Shave the fur of the right temporal region and ventral neck region using small hair clippers to expose the skin. Disinfect the surgical skin area, using a 5% chlorhexidine solution for 3 min. Apply ocular lubricant onto both eyes, to prevent them from drying during the surgery.
   NOTE: If required for pulse oximetry readings to monitor blood oxygen saturation, heart rate, and breathing rate, use hair removal cream to clear the hind leg of hair. Applying the cream using clean cotton buds and, after the hair removal, wash the area with the dilute chlorhexidine solution. Perform this step in a pre-operative area to minimize the risk of loose fur contaminating the sterile surgical field. Researchers may prefer to use other surgical site preparations according to local practice.
6. Transfer the mouse to the surgical area and place it in the prone position on a homeothermic heat mat covered by a sterile drape. Maintain anesthesia via a nose cone. Insert a rectal probe to monitor the mouse’s body temperature and maintain it at 37.0 ± 0.6 °C.
   1. Prior to any surgical incisions, check the hind paw withdrawal reflex and blink reflex to confirm the anesthesia depth.
7. Attach a laser Doppler flowmetry (LDF) probe to record the blood flow to the MCA territory.
   1. Using a dissection stereoscope, make an incision of < 1 cm at the midpoint between the right eye and ear on the exposed temporal skin using a No. 15 scalpel. Bluntly dissect the underlying tissue covering the skull, gently scraping this away from the bone and drying the surface with sterile cotton swabs.
      NOTE: Care must be taken not to damage the temporalis muscle.
   2. Take hold of the 0.7 mm, flexible single fiber-optic probe attached to the LDF monitor, cut its end using a sharp scalpel to gain a flat edge, push this through the donut-shaped probe holder, and place a small amount of optical matching gel at the end of the fiber.
   3. Place a small amount of waterproof super-adhesive glue around the bottom edge of the probe holder; allow this to partially dry and become tacky.
   4. Dry the skull above the temporal bone and place a small amount of topical tissue adhesive in a circle to the bone, just enough for the probe holder.
   5. Place the probe holder, with the fiber optic probe already in place, on this area (6 mm lateral and 2 mm distal from the bregma).
   6. Allow time for the glue to dry; once dry and attached, begin recording the LDF data.
8. Gently turn the animal over to a supine position, taking care to support the laser Doppler probe and prevent its detachment. Gently tape the two forepaws down with microporous medical tape, sliding a pair of cotton buds (or something similar, such as closed forceps) under the neck to lift the area and create tension. Cover the mouse with a sterile drape to maintain aseptic coverage.
9. Begin the dissection and exposure of the CCA.
   1. Make a 1.5 cm midline incision on the exposed ventral neck using a No. 15 scalpel blade.
   2. Using gentle blunt dissection techniques, gently retract the salivary glands to the sides, exposing the trachea.
   3. Using blunt dissection, dissect the CCA free from the surrounding tissue and vagal nerve.
      NOTE: Avoid touching the vagal nerve directly, as any damage to the vagal nerve can impair mobility, feeding, and breathing.
10. Pass two small (2 cm) sections of a non-dissolvable 6-0 suture below the CCA, dorsal to the vessel and ventral to the vagal nerve. Draw one silk tie closer to the surgeon and tie it tightly around the CCA (proximal tie). Loosely tie the second silk tie (distal tie) toward the bifurcation of the ICA/ECA.
11. To begin to isolate a section of the CCA, apply a microvascular clip just above the distal tie but not obstructing the bifurcation. Using micro Vannas scissors, make a small hole into the CCA.
    NOTE: The ECA must remain patent at all times. The CCA incision must be no more than 40% of the width of the vessel and in an easily accessible ventral position; this will play an important role in the vessel repair stage post-MCAO.
12. Insert a 7-0 silicone-coated monofilament into the CCA, advancing toward the microvascular clip.
    NOTE: The filament size must be established based on the mouse’s weight prior to the surgery; refer to the manufacturer’s guidelines.
    1. Tighten the distal tie, enough to fasten the filament into place without damaging it. Remove the microvascular clip using clip holders.
       Note: No blood loss should be seen at this point. If there is a backflow of blood, the tie holding the filament is not tight enough.
13. Advance the filament into the ICA. 
    1. Ensure the filament remains within the ICA and does not pass into the pterygopalatine artery (PPA). Do this by slightly lifting and pulling the CCA, using one of the silk ties, to the outer side of the animal’s body, and steer the filament to bend past the internally facing opening of the PPA.
       Note: Once advanced to the MCA branch origin, a drop in the relative blood flow to the supplied territory will be visible on the LDF values; this confirms the filament placement.
14. Secure the filament in place with the distal silk tie, tying this tighter. Leave it in place for the duration of the occlusion time period.
    NOTE: Dependent on individual protocols, the animal can be moved to a recovery cage, following wound suturing, to recover, or remain under anesthesia for the duration of the occlusion period. In the latter case, ensure the wound is prevented from drying out by using sterile prewarmed 0.9% NaCl saline. 

3. Post-occlusion

1. At the end of the MCAO period, immediately retract the filament until the white filament head is clearly visible.
2. Then, Loosen the distal CCA tie just enough to remove the filament head most of the way out of the vessel.
   NOTE: The filament can be fully removed at this point if the surgeon is confident with speed to tie the distal tie to prevent blood loss.
3. Place one microvascular clip in a horizontal position toward the CCA bifurcation next to the distal tie. Loosen the distal tie and remove the filament fully. Add another microvascular clip below the proximal CCA tie toward the surgeon.
   NOTE: Ensure the vessel is far enough into the prongs of the clamp to prevent any slippage. The placement of the clips is essential to ensure clip removers easy access during later steps. The clips can be used to help lift the vessel to allow a better clearance and placement of the tissue patch, placing the clips horizontally across the surrounding muscles.
4. Carefully remove both silk ties using Dumont #5 forceps or micro Vannas scissors and dry the area using sterile cotton buds.
   NOTE: Care must be taken not to cut through/into the vessel.
5. In a sterile Petri dish, add fibrinogen and thrombin sealant solutions 1 and 2 (see the Table of Materials), ensuring the two substances stay separate, ready for mixing later.
   NOTE: Only very small volumes of the agents are required (< 0.25 mL each). Keeping the solutions separate prevents a premature reaction between the two constituents. Ensure the cap is replaced the same way as it was removed to prevent cross-contamination of the two syringes, which would cause the agent to react and set within the syringe. Prior to use, store the solutions at -20 °C. When required for the first surgery, thaw the sealant to room temperature. Do not refreeze the sealant; it must remain at room temperature and can be stored this way. The syringe can be used across multiple surgeries making it cost-effective; however, we recommend to not use the same vial for longer than 1 week to prevent contamination.
6. Use blunt dissection along the muscle with Dumont no 5 forceps and micro Vannas scissors to obtain a thin ventral slice of sternocleidomastoid muscle to use for the tissue pad, ensuring the slice is no more than 1 mm thick and runs along the top fibers of the muscle.
   NOTE: Do not cut through/across the muscle, as this will significantly impair its function. The tissue must be large enough to cover the CCA incision comfortably.
7. Using Dumont #5 forceps, take the tissue pad and mix the tissue evenly across the two fibrinogen and thrombin sealant solutions, forming a channel between the two reagents. Coagulation will occur quickly; as soon as coagulation begins, remove the tissue pad to the CCA incision. Place the tissue pad flat down with a medium firm pressure and open forceps.
   1. Swiftly remove the distal microvascular clip whilst still gently holding the tissue pad in place.
      Note: This allows some backflow of blood to further activate the fibrinogen and thrombin sealant reagents. Just enough pressure is required to hold the pad in place but not to fully block the vessel.
   2. Slowly alleviate the pressure from the tissue pad, allowing blood to flow under the incision area. Now, slowly and gently release the pressure from the proximal microvascular clip and fully remove.
   3. NOTE: In order to ensure the vessel becomes fully patent, the placement of the tissue pad and the removal of the microvascular clips must occur quickly to prevent the tissue pad from sealing to the inside of the CCA. However, if there is a small amount of blood leakage, re-place some light pressure onto the tissue pad to allow further time for clot formation and sealing to occur. If the blood leakage is substantial or the tissue pad does not appear to be sealing, maintain pressure to prevent blood loss and replace both CCA microvascular clips to isolate the incision and prevent further blood loss.
8. In the event the tissue pad does not seal to the vessel, a second attempt can be made, following steps 3.3–3.7.3.
9. Once the vessel is sealed, suture the wound using dissolvable 6-0 sutures. If LDF recording was continued throughout surgery, remove the LDF from the skull and suture wound using dissolvable 6-0 suture.

4. Post-operative Care

1. Place the animal into a prewarmed recovery cage (situated on a heated shelf/mat at 35 °C, or within a heated chamber).
   NOTE: Researchers may prefer to use other temperatures and durations according to local practice.
2. Provide all animals with 200 µL of prewarmed 0.9% NaCl saline sc. immediately post-operation, at 4 h post-operation, and 2x daily for 72 h.
   NOTE: The administration of prewarmed 0.9% NaCl is animal led. If more is required, more fluids can be administered to ensure a good recovery.
3. In the recovery cage, give the animals unrestricted access to soggy diet pellets, dry diet pellets, rehydration gel, and gel food, alongside ad libitum access to water.
4. Repeat the sc. carprofen injection at 24 h post-operation (see step 2.3).
   NOTE: All animals received the same dose of carprofen; any neuroprotective effect is likely to be negligible.
5. At regular intervals for 48 h, perform post-op mouse grimace scoring¹⁷ to assess pain levels to aid the decision to administer further analgesia.
6. Weigh the animals immediately before surgery and then daily following the procedure. Perform daily observations and complete welfare sheets to monitor their food and water intake and clinical signs.
7. Undertake functional observations at 24 h and 48 h post-operation. Assess the mice on a focal deficit scale. Evaluate their body symmetry, compulsory circling, gait, 45° grid climbing, circling behavior, front limb asymmetry, and whisker touch response^18,19,20.

5. Magnetic Resonance Imaging and Image Processing

1. Measure the lesion volume (LV) using structural magnetic resonance imaging (MRI).
   NOTE: Alternative methods, such as histological staining with triphenyltetrazolium chloride (TTC), have previously been used and correlate to structural MRI data. However, this method can only be used at the endpoint of a study and not longitudinally. Using longitudinal MRI scanning will reduce the number of animals required for a study.
   1. After 48 h, following the induction of the MCAO, anesthetize the mouse with isoflurane (5% isoflurane in 1 L/min of O₂ for induction, 1.5–2% isoflurane for maintenance).
   2. Transfer the mouse to the MRI cradle, place it on the respiration sensor for monitoring its breathing rate, and implant the rectal temperature probe for monitoring its temperature during the scanning. Place the 2-channel mouse-brain RF receive coil over the brain for a signal and place the cradle in a 9.4 T horizontal bore scanner.
      NOTE: Here, a volume coil with a 72 mm inner diameter was used for the RF transmission.
   3. Acquire T2-weighted scans using a fast spin-echo sequence. Set the repetition time (TR) to 3,000 ms and the echo time (TE) to 40 ms. Use 18 mm x 18 mm as the field of view (FOV), and obtain a 256 x 256 acquisition matrix with slices of 18 mm x 0.8 mm and three signal averages in approximately 10 min.
   4. Acquire diffusion tensor images (DTI) using a fast spin-echo sequence. Set the TR to 1,730 ms, the TE to 35 ms, the FOV to 20 mm x 20 mm, and obtain a 128 x 128 acquisition matrix with slices of 16 mm x 1 mm, two signal averages, 14 diffusion encoding directions, and a maximum b-value of 1,024 s/mm².
   5. Measure the LV on the T2-weighted images using an image display and measuring software package. Measure the lesioned area, collating the values together to calculate the total lesion volume while taking into account the MRI slice thickness (set during the MRI scan).
   6. Take into account any swelling and the percentage of area lesion volumes while measuring the full contralateral and ipsilateral hemispheres. Correct for any brain swelling due to edema using an indirect method to measure the lesion volume as described previously^21,22. Only include slices that contain cortex tissue and not frontal lobe or cerebellum tissue according to a standard mouse brain atlas to avoid overcorrection.
2. Measure the lesion diffusion parameters and obtain the core and penumbra regions of interest.
   1. Generate the apparent diffusion coefficient (ADC) and fractional anisotropy (FA) maps from the diffusion tensor images using appropriate MRI analysis software.
   2. Align the DTI images with the T2-weighted images using appropriate image analysis software capable of performing a linear registration of images. Following the registration, subtract the diffusion-weighted lesion masks (ischemic core) from the T2-weighted lesion masks (ischemic core and penumbra) to estimate the penumbra region. Apply the resultant masks (core and penumbra) to the ADC and FA maps to quantify the diffusion parameters within the core and penumbra.
   3. Translate the ipsilateral core and the penumbra masks about the brain midline in order to obtain contralateral ADC and FA values for comparison.