Mouse Stereotaxic Surgeries for Intracranial Viral Injection

Prepare a clean empty mouse cage on a heating pad and a clean mouse cage with gel food for post-op care

Set up sterile working area including stereotaxic frame

Weigh mouse

Anesthetize mouse in induction chamber (recommended: 2.5% isoflurane, 200ml/min flow rate)

Hair over surgery area can be quickly clipped before transferring the mouse onto the stereotaxic frame

Once the mouse is deeply anesthetized (~0h 5m 0s), stop anesthesia and move the mouse to the stereotaxic frame over the heating pad with the temperature probe and secure the mouse mouth on the nose cone.

Restart anesthesia (directed towards the nose cone)

The heating pad settings should be adjusted so that the temperature probe placed under the mouse should read a body temperature between 33°C - 37°C

Apply ophthalmic ointment over eyes

Inject appropriate volume (based on mouse weight and desired dosage) of analgesic; an appropriate amount of saline can also be injected to prevent dehydration during the procedure

Carefully insert and secure the ear-bars. The position of the mouse head will be verified and adjusted once the skull is exposed, but it is recommended to make sure that the head is not visibly tilted

Clean the area of the incision with the povidone-iodine swab followed by the ethanol swab

Repeat Step 11, 3 times

It is preferred to apply line-block anesthetic (0.15% bupivacaine) under the skull skin before starting the procedure rather than applying EMLA cream on the sutured skin at the end of the surgery

With the fine scissor, expose the skull by making an anterior-posterior incision

Visually identify bregma and lambda

Insert a glass pipette (a small volume of non-toxic food dye can be used to help marking the relevant spots) on the stereotaxic arm holder and lower it onto the skull

Mark bregma by gently touching the intersection of the coronal/sagittal sutures with the pipette tip, and zero the coordinates on the reader

Move to lambda (intersection of lambdoid and sagittal sutures) and measure its position relative to bregma

Minimize the deviation of dorso/ventral (D/V) and medio/lateral (M/L) distance between lambda and bregma by adjusting the position of the head

Re-zero the coordinates at bregma and repeat bregma/lambda measurements until satisfactory

Once the head is in the correct position, it is possible to identify the desired injection spot

It is recommended to use the measured anterior/posterior (A/P) distance between bregma and lambda to calculate an adjustment factor for the final coordinates: the measured B-L distance will be divided by the reference distance of 4.21. For an adult mouse, the obtained value (“adjustment ratio”) should be close to 1, and in this case no coordinates adjustment is required (but still optional). For smaller mice, the reference coordinates should be multiplied by the calculated adjustment ratio to obtain the final coordinates for the specific mouse

Move the pipette to the spot indicated by the adjusted A/P and M/L coordinates and mark it

Whether performing uni-lateral or bi-lateral injections, it is recommended to mark the spots on both sides of the skull, and to measure their relative dorso-ventral position. Their relative D/V deviation should be minimized by adjusting the position of the head

Once the desired spot has been marked, the marker pipette can be removed, and a hole is drilled in the skull at the indicated position

Blood and debris are cleaned with sterile saline and sterile cotton swabs

Insert micropipette with volumetric references in the holder and connect it to a syringe to apply positive/negative pressure

Draw up desired volume of viral solution in the syringe by applying negative pressure

Lower pipette loaded with the viral solution into the hole until the tip touches the dura. Zero the dorso-ventral coordinate

Gradually lower the pipette tip into the brain until the desired dorso-ventral coordinate is reached

Slowly inject the desired volume of viral solution (recommended: ~150nl/min) by gently and gradually applying positive pressure

Release pressure and leave the pipette in position for ~5-10 min so that the viral solution can spread and be absorbed by the tissue

Slowly retract viral injection pipette and discard it in an appropriate waste collection bin

Suture Skin

Optional: Repeat saline injection to prevent dehydration

Remove animal from stereotaxic frame and place it in the clean, empty cage on heating pad until deambulatory (~0h 10m 0s - 0h 15m 0s

Once awake and deambulatory, mouse can be moved to the clean cage with gel food, also on heating pad

24h 0m 0s after surgery, a second dose of Metacam is administered and antibiotic ointment is applied on the sutured skin

The health status of the mouse is monitored over the following days. If needed, additional doses of Metacam or saline can be administered

Mouse is normally kept in a cage on heating pad for at least 4 days and is then moved to standard housing

Mice are sacrificed for experiments at least 10 days after surgeries