The overall goal of this procedure is to study focal cerebral ischemia in animal models. This is accomplished by first making a small incision on the skin between the left eye and the base of the ear. Next, a craniotomy is performed at the junction of the zygomatic arch and the squamal bone to expose the middle cerebral artery, the middle cerebral artery is then ligated using a small vessel cauterizer.
Finally, the skin is sutured to close up the wound. Ultimately, results can be obtained that show an infarct located on the left hemisphere of the mouse brain. The brain can be stained with TTC or scanned using MRI for the visualization of a stroke.
The stroke brain can also be used for determination of protein or mRNA expression. Hi, I'm Garza Chola. I'm a graduate student in Gale Johnson's lab at University of Rochester.
This method can help answer key questions in the stroke field, such as illustrating the mechanisms which lead to cell that in ischemic brain. To prepare for surgery, sterilize all the surgical instruments, gauze sponges, and cognitive applicators by autoclave. Keep surgical tools in 70%ethanol during the surgery and air dry on sterile gauze right before use.
Spray the work area with 70%ethanol two hours before the surgery. Inject the mouse with 0.05 milligrams per kilogram of buprenorphine. The drug will be administered again immediately after surgery, and then every three to five hours during the first 24 hour postoperative period.
Use an anesthetic vaporizer to an anesthetize the mouse with a 3%ISO fluorine, 20%oxygen gas mixture. Adjust the oxygen amount with a flow meter. Test the level of anesthesia by toe or tail pinching.
Maintain the level of anesthesia with 2%ISO fluorine. Apply artificial tears to the mouse's eyes and use caution to avoid damaging the eye. During the surgical procedure.
Place the mouse on its right side in a lateral position using the animal clippers. Shave the area on the left side between the left eye and the base of the left ear. Cleanse the area by alternating between a Betadine solution and 70%eth ethanol using cotton tip applicators and lightly rubbing the area.
Repeat is necessary. Place the mouse onto a heating pad connected to a rectal probe To maintain body temperature at 37 degrees Celsius, insert the rectal probe by using mineral oil. Position the mouse on its right side under the microscope and secure with tape.
Cut a window in a gauze sponge and cover the surgical area with fine straight scissors. Make a vertical incision between the left eye and the base of the left ear. Use curved hemostats to keep the surgical area open if excessive drying of the tissue occurs during the surgery.
Use cotton tip applicators to apply sterile PBS. Use spring scissors to make a horizontal incision on the temporal muscle and slowly pull with forceps to slightly separate the temporal muscle from the skull. Hold the jawbone with curved forceps and use an 18 gauge needle to make a small subtemporal craniotomy at the junction of zygomatic arch and squam sole bone.
With bone run jurors remove small pieces of the skull to expose the MCA. The zygomatic arch and orbital contents should not be damaged during this process. Using a small vessel cauterizer ligate the distal portion of the MCA, place the temporal muscle back into its original position, and use five zero nylon sutures.
To close the incision site, discontinue anesthesia and remove the rectal probe. Inject the mouse with a second dose of buprenorphine subcutaneously. Return the mouse to the cage, which was kept at 37 degrees Celsius.
With a heating panel. Closely monitor the mouse for the next 24 hours for any discomfort, including decreases in appetite or water consumption, hunched posture, increased respiration or pilo erected hair. Provide the mouse with recovery gel immediately after the surgery.
Wait two to three hours after surgery. Then e inject the mouse with buprenorphine subcutaneously every three to five hours up to 24 hours. 24 hours after surgery, deeply anesthetize the mouse and perfuse the mouse with 4%TTC.
Remove the brain and place it into a 4%paraform aldehyde solution overnight. The next day, position the brain into the sectioning block. Use razor blades to slice the brain into one millimeter thick slices.
Place the slices next to a millimeter scale ruler. Using a digital camera connected to a dissecting microscope. Photograph the slices in the image J software.
Click on the file menu and open the desired photo. Click on the straight line tab in the program. Draw a straight line between the two margins on the ruler.
Click on analyze menu and select set scale. Set the known distance as one and unit of length as millimeter. Click on freehand circle tab.
Draw circle to outline the contralateral hemisphere. Click on analyze menu and select measure. The calculation window will pop up showing the area of the circle drawn.
Draw another circle around the ipsilateral hemisphere, excluding the white stroke area. Measure the area as shown previously. The new value will be added to the calculation window.
The difference between the first and second values represents the area of the infarct. Calculate the stroke area in each slice by repeating the above steps for slice thicknesses of one millimeter. Calculate the stroke volume by summing the stroke area calculated in each slice.
This figure shows the TTC stain brain 24 hours post permanent MCA ligation in a wild type C 57 BL six mouse. The stroke site is white in appearance and can be seen from different magnification and angles. Shown here are one millimeter thick slices of the TTC stained stroke brain from anterior to posterior infarct.
Volume was calculated 24 hours post-surgery. The volume of the infarct is approximately 23 cubic millimeters Once mustard. This technique can be done in 30 minutes if it's performed properly.
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