Intracranial Injection of Adeno-associated Viral Vectors

Viral vectors can be used to express fluorescent proteins in specific subsets of cells in tissue. Here a virus expressing engineered proteins is introduced into the primary visual cortex of the brain by first performing a small craniotomy over this area. Next, a glass micro pipette is lowered into the brain and a dental associated viral vector is injected into the brain and the surgical site is closed.

Once the animal has recovered, imaging can be performed to follow the fates of fluorescently labeled cells in vivo or in brain sections. The main advantages of this technique over existing methods such as fluorescent dye labeling and transgenic mouse lines, are that it can target individual cell types and its less expensive and time consuming than establishing a transgenic line. The use of adeno-associated viral vectors is approved for biosafety level one or BSL one.

Here, a lab coat and gloves will be worn in accordance with procedures for handling BSL one agents begin by preparing a biosafety cabinet Class two for Ali quoting virus. This will preserve the activity of the virus by avoiding repeated, freezing and thawing. Clear the biosafety cabinet of any unnecessary objects and sterilize the surface with 70%ethanol.

Place a beaker containing a 10%bleach solution in the biosafety cabinet for collecting a dental associated virus contaminated waste. Also placed sterile 0.5 milliliter tubes and a container of dry ice. Thaw the viral stock on ice outside of the biosafety cabinet.

Vortex the virus and open the tube inside the hood. Pipette the desired volume into a 0.5 milliliter tube. Close the tube and place it in the dry ice.

To flash, freeze the virus when all of the virus has been aliquot. Dispose of the empty tube and pipette tips in the 10%bleach waste container. Remove the waste container from the hood.

Add additional 10%bleach. Wait five to 10 minutes, then pour the bleach down the sink. Dispose of all plastic waste in a biohazard container.

According to institutional protocols, clean any equipment or surfaces that came in contact with the virus with 10%bleach. Remove and discard gloves. Store the aliquots in a minus 80 degree Celsius freezer.

Cover the surgical area with absorbent lab bench paper. The surgical tool should be sterile and surgery should be performed under aseptic conditions. Cover the area adjacent to the surgical area with absorbent lab bench paper.

This area will be dedicated to loading the micro pipettes with virus. Place an aliquot of the virus in a container of ice and allow it to thaw on ice as the surgery is being performed, set up a waste container of 10%bleach in the dedicated virus handling area for disposing any pipette tips or other items that come in contact with the virus. Pull a glass wire troll micro pipette to a tip diameter of approximately 20 microns.

Place a small drop of mineral oil on the blunt end of the micro pipette and insert the wire plunger provided with the micro pipettes. Secure the micro pipette in the clamp of the micro pump arm. Using surgical scissors.

Remove the hair from the top of the head of an anesthetized mouse. Then secure the mouse in a stereo attacks. Bathe the head in ethanol and Betadine to sterilize the area.

Place a drop of tobradex eye ointment on each eye to keep the eyes moist during surgery. Make an incision down the midline of the head and pull back the skin to expose the skull using fine tipped forceps. Carefully remove the facia from the skull.

Locate the area to be injected using stereotaxic coordinates and mark the skull with a surgical pen. Using a dental drill with a 1.4 millimeter burr, thin an area of the skull, approximately two millimeters in diameter until the skull cracks. Dividing the thinned area into several segments throughout this procedure.

Apply sterile saline to keep the skull moist. Perform a craniotomy by gently removing the thin skull segments using extra fine tip forceps. Place a Kim wipe over the lid of the tube of virus.

Then open the tube for a one microliter injection. Pipette 1.5 microliters onto a small piece of paraform. Next, place the tip of the micro pipette into the viral stock and manually withdraw the plunger.

If difficulty is experienced drawing viral stock into the micro pipette, the tip can be enlarged slightly by piercing a kim wipe with the micro pipette to break a small portion of the glass, lower the micro pump arm onto the plunger until a small amount of virus is dispelled from the tip of the micro pipette. Remove this drop with a cotton tip applicator and discard the applicator in a waste container. Apply a drop of mineral oil to the tip of the micro pipette to prevent clogging as the micro pipette is lowered into the brain, the using X and Y stereotaxic coordinates position the micro pipette over the area to be injected.

In this experiment, the stereotaxic coordinates used to locate the primary visual cortex are 2.7 millimeters posterior torema and 2.5 millimeters lateral to the midline. Very slowly lower the micro pipette at an approximate rate of one millimeter every minute. To the desired depth, enter the desired injection parameters into the CYS micro four micro pump controller box.

For this experiment, one microliter over 10 minutes will be used as a rate of injection. Initiate the injection. When the injection is finished.

Leave the pipette to rest for one to two minutes to prevent influx of virus during removal. After this period, very slowly, remove the micro pipette from the brain, suture the scalp, and seal it with tissue glue. Allow the animal to recover under a heat lamp until it is ambulating.

Then return it to its cage. Imaging experiments can be performed days to weeks after viral injection. Following the surgery and injection, rinse the micro pipette with 10%bleach and discard it in a sharps container.

Add additional 10%bleach to the waste container. Wait five to 10 minutes, then pour the bleach down the sink. Dispose of waste in a biohazard container according to institutional guidelines.

Discard the lab bench paper into a biohazard bin. Wipe down all of the surfaces and instruments that may have come in contact with the virus. With 10%bleach, unused virus may be frozen.

Again, keeping in mind that repeated freeze haw cycles cause degradation of the virus. This image shows transduced neurons after injection of doublet, stranded, adeno-associated virus serotype.One. The cell body and the proximal and distal dendrites are clearly visible in fixed sections.

Here, labeling typical of an intracranial viral injection in primary visual cortex is shown. Note the extent of viral spread as well as labeled neurons, glia, and processes. Injection of viral vector into the hippocampus also results in clearly visible labeled cells as shown in this image, Once mastered, this technique can be done in one hour.

While attempting this procedure, it's important to remember to use great care when lowering and raising the microFIT in the brain. Don't forget that working with viral vectors can be extremely hazardous. Always follow the proper procedures for handling biohazardous agents set by the Centers for Disease Control.

Please consult your institution safety office for approval and guidance before performing experiments using virus.