Intracranial Stereotactic Injection of Adeno-Associated Viral Vectors Protocol

Experiment Summary

Adeno-associated viral (AAV) vectors provide a useful delivery vehicle for gene therapy. The broad array of AAV serotypes enable preferential transduction of different cell types and tissues. The applications of AAV in the central nervous system (CNS) include many aspects, such as research and treatment in neurological diseases and CNS injuries, as well as local transgenic studies in certain models of neurological diseases.

AAV-mediated transgenesis in the brain is usually not homogeneous due to differences in AAV serotypes, transcription factors and injection sites, and this heterogeneity is manifested in terms of time and space (specificity for different cell types). In addition, the technique of AAV virus preparation (affecting purity and titer) and the method of virus injection can greatly affect the efficiency of AAV-mediated transgenesis. Indeed, by injecting AAV directly into the brain or into a peripheral area of the brain (intramuscular or intravenous injection) different transduction patterns can be produced. It has been documented that AAV9 can cross the blood-brain barrier in mouse/cat and non-human primates. Moreover, AAV carriers can be retrogradely transported from muscle to spinal cord motor neurons. Experiments have shown that the retrograde efficiency of AAV1 is high. This paper focuses on the direct intracerebral injection of AAV by stereotaxic instrumentation.

Materials and Reagents

70% ethanol
30% hydrogen peroxide
UltraPure^TM distilled water
10% povidone-iodine topical anesthetic
Test article (AAV): Specific viral vectors expressing a gene of interest are available from Creative Biogene. ( You can choose the appropriate AAV vector according to your experimental needs, commonly used ones include HBV genome AAV particles, recombinase AAV particles, reporter AAV particles, etc.)
Diluent: injection buffer, such as lactated Ringer's, appropriate for CNS delivery
2-4% isoflurane/oxygen gas
Bupivacaine 0.5%
Lubricant eye ointment
Mice: Most of the mouse models used in our lab are on common backgrounds like FVB/NJ or C57Bl/6J and can be purchased.
Bucket of ice
10–100 µl pipette; pipette tips
10-µl Hamilton syringes; needles
Cotton-tipped applicators
Scalpel blades; scalpel handle
Sutures
2 Hemostats
Small Animal Trimmer
Forceps
Scissors
Stereotaxic injection rig
Injection pump
Heat lamp
0.5-ml insulin syringes to administer anesthetics/analgesics
Sterile surgical drape
Paper tissue
Gauze sponges
Dremel Bit (105 1/8-in shank)
Sterile gloves (at least 2 pair)

Procedure

Brain atlases of both adult male and female rats and mice are available in the literature and in books. The specific injection site can be determined based on the needs of the study.

A. Determine the amount of viral vector needed

Determine the amount of viral vector for use at the appropriate titer and volume needed for the experiment. Injection volumes are based on the volume of the target anatomical structure.
Thaw the appropriate volume of virus on ice.

A. Syringe preparation

Use the clean syringe to slowly draw up and expunge the entire volume of virus, minimizing bubble production.
Repeat this procedure 20 to 30 times with the same virus to adequately saturate the interior of the syringe, then discard the used virus.
Draw the virus into the saturated syringe.
Place the syringe into the injection pump.

B. Prepping mice for surgery

Place the mouse into the anesthesia induction chamber with 2% to 4% isoflurane. Adequate depth of anesthesia is achieved in 5 to 10 min.
Verify the absence of pedal reflexes indicative of surgical anesthesia while monitoring breathing.
Remove the fur between the mouse's ears using clippers or a hair removal solution.
Load mice into the ear bars being careful to keep the flat plane of the skull parallel to the base of the injection rig.
Place the front teeth in the tooth bar and then snugly fit the nose guard/nose cone to the mouse's nose to impede movement of the head.
Liberally apply lubricating ointment with a cotton-tipped applicator to the eyes to prevent drying of the corneas.
Inject an analgesic approved by the appropriate Institutional Animal Care and Use Committee.
Clean the shaved surface of the head by swabbing with 10% povidone-iodine followed by 70% ethanol. Repeat three time.
With a sterile scalpel blade, make a small incision rostral to caudal through the midline of the skin to expose the surface of the skull.
Swab the surface of the skull with a cotton-tipped applicator moistened with a small amount of 30% hydrogen peroxide to visualize the stereotaxic landmarks: bregma and lambda.

A. Target coordinates

Manipulate the arms of the stereotaxic rig to center the tip of the needle directly over bregma.
Move the needle to your target using x-axis (anterior/posterior) and y-axis (medial/lateral) measurements.
Use the autoclaved non-toxic marker to place a small dot where the needle will enter the skull.
Drill a small burr hole through the skull using the Dremel bit (105 1/8-in shank).
If measuring from the surface of the brain, lower the needle to the surface and make the appropriate calculations for depth (z-axis).
Slowly insert the needle tip to the desired target depth.
Begin the infusion.
Once the infusion is complete, keep the needle at the target location for an additional 5 min. This allows the virus to disperse away from the injection site. Slowly retract the needle out of the brain to prevent the virus from flooding back out the needle tract.
Repeat steps 21–27 for additional injections to the same mouse using appropriate coordinates.

Fig. 1 Loading a mouse into the stereotaxic rig.

B. Closing the incision

The incision can be closed by a variety of methods: sutures, wound clips, or VetBondR.
For suturing: hold the needle of the suture with a hemostat.
Using forceps, guide the needle through the skin on either side of the incision at the rostral end. Leave an inch of suture not pulled through the skin.
Loop the needle end of the suture around a second hemostat 2–3 times before clasping the tail end of the suture with the second hemostat and pulling the loops down the nose to form a knot.
Repeat 3 times alternating the direction of loops over the second hemostat.
Trim suture.
Repeat steps 33–36 until the entire incision is closed.
After closing the incision, add a topical analgesic such as bupivacaine. This will numb the area as well as dissuade cage mates from licking or gnawing at the sutures.
Place the mouse in a warm, dry environment free of loose bedding to emerge from anesthesia.
After the mouse is bright/alert/reactive, place them in their home cage.

* For research use only. Not intended for any clinical use.

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Intracranial Stereotactic Injection of Adeno-Associated Viral Vectors Protocol

Experiment Summary

Materials and Reagents

Procedure

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