A Modified Transcranial Middle Cerebral Artery Occlusion Model to Study Stroke Outcomes in Aged Mice

Summary

This protocol demonstrates a unique mouse stroke model with a medium-sized infarct and an excellent survival rate. This model allows preclinical stroke researchers to extend the ischemia duration, use aged mice, and assess long-term functional outcomes.

Abstract

In experimental stroke research, middle cerebral artery occlusion (MCAO) with an intraluminal filament is widely used to model ischemic stroke in mice. The filament MCAO model typically exhibits a massive cerebral infarction in C57Bl/6 mice that sometimes includes brain tissue in the territory supplied by the posterior cerebral artery, which is largely due to a high incidence of posterior communicating artery atresia. This phenomenon is considered a major contributor to the high mortality rate observed in C57Bl/6 mice during long-term stroke recovery after filament MCAO. Thus, many chronic stroke studies exploit distal MCAO models. However, these models usually produce infarction only in the cortex area, and consequently, the assessment of post-stroke neurologic deficits could be a challenge. This study has established a modified transcranial MCAO model in which the MCA at the trunk is partially occluded either permanently or transiently via a small cranial window. Since the occlusion location is relatively proximal to the origin of the MCA, this model generates brain damage in both the cortex and striatum. Extensive characterization of this model has demonstrated an excellent long-term survival rate, even in aged mice, as well as readily detectable neurologic deficits. Therefore, the MCAO mouse model described here represents a valuable tool for experimental stroke research.

Introduction

Nearly 800,000 people suffer a stroke in the US every year, and most of these strokes are ischemic in nature¹. Timely restoration of the cerebral blood flow with tissue plasminogen activator (tPA) and/or thrombectomy is currently the most effective treatment for stroke patients; however, the full recovery of neurologic functions in the long term is rare^2,3. Thus, searching for novel stroke therapy that targets functional improvement is an intense area of research that requires clinically relevant animal models of stroke.

The most common ischemic stroke model in rodents uses intraluminal middle cerebral artery occlusion (MCAO) to induce stroke. In this model, initially developed by Zea Longa in 1989, a nylon filament is introduced into the internal carotid artery (ICA) to block the blood flow to the middle cerebral artery (MCA)⁴. However, this model has limitations. First, when the filament is inserted into the ICA, the blood flow to the posterior cerebral artery (PCA) could be partially blocked as well, especially in mice. Critically, the posterior communicating artery (PcomA), a small artery that connects anterior and posterior cerebral circulation, is frequently underdeveloped in some mouse strains, such as C57Bl/6, the strain predominantly used in experimental stroke research. This patency of the PcomA is believed to contribute to the variability in lesion size in mice after stroke⁵. Indeed, when blood flow to the PCA drops precipitously during MCAO, and the PcomA is unable to provide sufficient collateral blood flow, the stroke infarct can expand into the territory of the PCA. Moreover, in this model, a long duration of ischemia leads to a higher chance of mortality in mice. Consequently, a short MCAO duration of 30-60 min is typically used in mice. However, most stroke patients experience a few hours of ischemia before reperfusion treatment. Thus, a mouse stroke model with an extended duration of ischemia is of high clinical relevance.

The overall goal of this procedure is to model ischemic stroke in mice that have a medium-sized infarct and an excellent survival rate. This transcranial MCAO model addresses critical attributes of clinical stroke, as prolonged ischemia can be performed, and aged mice tolerate this model well, allowing for the long-term assessment of functional recovery.

Protocol

All procedures described in this work are conducted in accordance with the NIH guidelines for the care and use of animals in research, and the protocol was approved by the Duke Institute Animal Care and Use Committee (IACUC). Young (8-10 weeks old) and aged (22 months old) male C57Bl/6 mice were used for the present study. An overview of this protocol is illustrated in Figure 1.

1. Surgical preparation

1. Examine the mouse for gross abnormalities and behavioral deficits.
   NOTE: Before surgery, it's important for surgeons to wear appropriate PPE (protective personal equipment), including surgical mask, cap, gloves, and gown.
2. Weigh the mouse; program the ventilator (see Table of Materials) based on the body weight.
3. Place the mouse in a 4 in x 4 in x 7 in anesthesia induction box. Turn on the oxygen flow meter (see Table of Materials), set at 30, and the nitrous oxide flow meter, set at 70. Turn on the vaporizer with 5% isoflurane.
4. Insert the guide wire into the 20 G intravenous (IV) catheter.
5. Take the mouse out of the induction box when its respiratory rate is reduced to 30-40 breaths per minute.
6. Lay the mouse on the surgical bench in a supine position. Pull the mouse's tongue out and hold it with the fingers of the left hand. Insert a laryngoscope (see Table of Materials) into the animal's mouth to visualize the vocal cord.
7. Stabilize the mouse chin on the laryngoscope using the right middle finger. Free the left hand to hold the 20 G IV catheter.
8. Insert the guide wire slightly into the vocal cord, and then slowly push the 20 G IV catheter into the trachea until the wing part of the catheter becomes even with the nose tip.
   NOTE: If the mouse is moving, do not insert the wire. This may cause trauma to the trachea and bleeding.
9. Turn on the ventilator (see Table of Materials), and connect it with the 20 G IV catheter intubated in the mouse. Reduce the isoflurane to 1.5%, and ensure that both lungs are mechanically ventilated.
   ​NOTE: Do not forget to reduce the isoflurane concentration. Otherwise, the mouse will receive an overdose of anesthesia.
10. Apply eye ointment onto both eyes and inject 5 mg/kg carprofen subcutaneously.
11. Keep the mouse in a lateral position with the right temporal area facing up. Maintain the rectal temperature at 37 °C using a heating pad (35 °C) and a heat lamp controlled by a temperature regulator (see Table of Materials).
12. Shave the surface area between the right eye and ear, and disinfect the surgical area at least three times with iodine and alcohol swabs.

2. MCAO surgery

1. Open the sterile instrument package for MCAO surgery. Wear sterile gloves and make a 1 cm skin incision between the right eye and right ear using surgical scissors.
   NOTE: Monitor skin color, body temperature, and response to toe pinch every 15 min.
2. Dissect the underlying fascia with forceps to expose the temporal and masseter muscles.
   NOTE: Be careful not to damage the parotid gland.
3. Use forceps to touch the lower part of the temporal muscle and detect the location of the zygomatic arch. Carefully pull aside the branches of the facial nerve.
4. Use the tip of a high-temperature cautery loop (see Table of Materials) to cut a 5 mm transverse incision on the temporal muscle.
5. Use two forceps to dissect the underlying zygomatic arch and expose the joint between the maxilla and zygomatic bones.
6. Use scissors to cut a 3 mm portion of the zygomatic arch and remove it. Separate the masseter muscle from the skull base.
   NOTE: Be careful not to fracture the retro-orbital sinus and superficial temporal vein.
7. Apply four small retractors positioned in different directions to expose the cranial skull base, with the trigeminal nerve branches pulled laterally by one retractor.
   NOTE: A sulcus on the external surface of the cranial base marks the location of the lateral fissure between the frontal and temporal lobes. The MCA lies here (Figure 2A), and its trunk and branches are visible through the thin, transparent skull (Figure 2B). The relationship of this artery to other major cerebral arteries is displayed in Figure 2A.
8. Apply a drop of 0.9% normal saline on the skull above the MCA trunk and proximal to the branch of the rhinal cortex. Use an electrical grinder to thin the skull until a small fracture is visible.
   NOTE: Do not push the grinder against the skull, as it may penetrate the skull and injure the underlying artery.
9. Use the tip of the forceps to lift the thinned skull and remove it. For sham mice, stop here, and do not ligate the artery.
   NOTE: A small rectangular window across the MCA trunk is formed.
10. Place a single-strand loop of black braided silk on top of the MCA (Figure 2C). Insert an 8-0 microsurgical needle to lift the MCA trunk, and tie the suture (see Table of Materials) under the needle, leaving both ends of the needle on the top of the silk thread loop knot (Figure 2D).
11. For transient MCAO, tighten the silk thread knot slightly under the needle to block arterial blood flow (Figure 2E), representing MCAO onset.
12. Use the forceps to hold the suture, and slowly remove the needle at the end of the ischemia (e.g., 60 min or longer).
    NOTE: When the needle is removed, the silk thread knot is slipped off the MCA, and the brain is reperfused (Figure 2F).
13. For permanent MCAO, firmly tighten the silk thread loop around the artery, and remove the needle. Cut and remove the excess suture materials.
14. Apply a drop of 0.25% bupivacaine to the skin incision, and suture the muscle and skin separately using 6-0 nylon sutures intermittently (see Table of Materials). Apply antibiotic ointment to the surface of the skin incision.
    NOTE: The skin incision can also be closed with sterile staples or glue.

3. Post-surgical care

1. Turn the isoflurane off to awaken the mouse. Disconnect the ventilator when spontaneous respiration is restored.
2. Transfer the mouse to a recovery chamber (see Table of Materials) with a controlled temperature.
3. Extubate the mouse when its righting reflex is restored, or it begins to move.
4. Closely monitor the mouse in a temperature- and humidity-controlled chamber. Return the mouse to the home cage after it gains full consciousness (recovery period ~2 h). Administer 5 mg/kg Carprofen subcutaneously daily for 3 days.

4. Laser speckle contrast imaging (LSCI)

1. Six and 24 h post-MCAO, mount the anesthetized mouse on the stereotaxic frame. Shave the top of the head, and clean it with three alternating swabs of iodine and alcohol.
   NOTE: Anesthetization was performed as mentioned in step 1.3. LSCI is also performed before MCAO.
2. Make a 3 cm midline skin incision and dissect the skin from the skull. Apply four small needle retractors to expose the skull top.
3. Move the laser speckle camera (see Table of Materials) above the head, and adjust the focus of the camera. Image the cerebral blood flow.

5. 2,3,5-triphenyltetrazolium chloride (TTC) staining

1. Deeply anesthetize the mouse with 5% isoflurane at the end of the experiment, typically on day 1, 3, or 28 after stroke. Pinch the tail to ensure there is no pain response.
2. Decapitate the mouse using a surgical scissor and harvest the brain. Incubate the brain in ice-cold saline for 20 min.
3. Put the brain in a brain slicer matrix on ice and drop cold saline on the brain. Cut the brain into 1 mm slices using thin razor blades.
4. Immerse the brain slices at the same orientation into a dish of 2% TTC solution (see Table of Materials). Keep the dish in the dark at room temperature for 15 min.
   NOTE: Normal brain tissue becomes red and ischemic tissue remains white.
5. Transfer the brain slices to 10% formalin for 24 h of fixation. Image the brain slices and measure the infarct area.

Results

With a direct view under a surgical microscope, it can be visually confirmed that MCA blood flow is blocked during ischemia. Our previous study showed a >80% blood flow reduction in the ischemic area using a laser Doppler monitor⁶. In order to determine post-MCAO blood flow changes, LSCI can be used to further confirm the ischemic insult and reperfusion (Figure 1). Indeed, in Figure 3A, it is observed that the blood supply was reduced ...

Discussion

The first transcranial MCA occlusion model was established in rats in 1981^11,12, and replaced by the no-craniectomy MCAO model in 1989⁴. The initial transcranial MCA occlusion had a wide surgical field, such that the entire zygomatic arch was removed and the muscles pulled laterally. Local tissues were swollen after surgery, causing stress and decreased food intake for the animals. In our modified transcranial MCAO model, the incision is l...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.25% bupivacaine	Hospira	NDC 0409-1159-18
0.9% sodium chloride	ICU Medical	NDC 0990-7983-03
2,3,5-Triphenyltetrazolium Chloride (TTC)	Sigma or any available vendor
20 G IV catheter	BD	381534	20 GA 1.6 IN
30 G needle	BD	305106
4-0 silk suture	Look	SP116	Black braided silk
8-0 suture with needle	Ethilon	2822G
Alcohol swabs	BD	326895
Anesthesia induction box	Any suitable vendor		Pexiglass make
Electrical grinder	JSDA	JD 700
High temperature cautery loop tip	Bovie	AA03
Isoflurane	Covetrus	NDC 11695-6777-2
Laser doppler perfusion monitor	Moor Instruments	moorVMS-LDF1
Lubricant eye ointment	Bausch + Lomb	339081
Mouse rectal probe	Physitemp	RET-3
Nitrous Oxide	Airgas	UN1070
Otoscope	Welchallyn	728	2.5 mm Speculum Otoscope served as a laryngoscope to visualize vocal cords in mice
Oxygen	Airgas	UN1072
Povidone-iodine	CVS	955338
Recovery box	Brinsea	TLC eco
Rimadyl (carprofen)	Zoetis	6100701	Injectable 50 mg/mL
Rodent ventilator	Kent Scientific	Rodent Jr.
Temperature controller	Physitemp	TCAT-2DF
Triple antibioric & pain relief	CVS	NDC 59770-823-56
Vaporizer	RWD	R583S