Intracranial injection of adeno-associated viral vectors | protocol (2023)

1. Virus Handling and Storage

1. Appropriate protection and handling techniques should be selected based on the biosecurity level of the virus to be used. These practices can be found in Biosafety in Microbiological and Biomedical Laboratories 5^IsEdition, available on the CDC website
   (http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm). The use of AAV vectors is approved for Biosafety Level 1 (BSL-1). For the experiment shown here, a lab coat and gloves are worn when handling the virus according to BSL-1 agent handling procedures.
2. To preserve viral activity, it is best to break it up into small aliquots to avoid repeated freezing and thawing.
3. Prepare a class II biosafety cabinet (BSC) to aliquot the virus by cleaning the BSC of unnecessary objects and sterilizing the surface with 70% EtOH. Place a beaker with a 10% bleach solution in the hood to collect AAV-contaminated residue. Sterile 0.5 mL tubes and a container of dry ice are also placed in the BSC.
4. Thaw virus stocks on ice outside of the BSC.
5. Swirl the virus and open the tube inside the BSC.
6. Pipet the desired aliquot volume (e.g. 5 L) into a 0.5 mL tube, cap the tube and place on dry ice to freeze the virus.
7. When all virus has been aliquoted, discard the virus tube and pipette tips in the 10% bleach waste container.
8. Remove the waste bin from the hood and add an additional 10% bleach, then pour the bleach into the sink. Dispose of plastic waste in a biohazard container according to the instructions of your institution's biosafety officer.
9. Clean any equipment or surfaces that have been in contact with the virus with 10% bleach. Discard the gloves.
10. Store aliquots in a -80 °C freezer.

2. Surgery

1. Cover the surgical area with absorbent laboratory paper. Surgical instruments must be sterilized and the surgery must be performed under aseptic conditions according to your facility's biosafety and animal use policies.
2. Select an area adjacent to the surgical area dedicated to loading the micropipettes with virus and cover with absorbent laboratory paper. Place an aliquot of the virus in a container of ice in this area and allow the virus to thaw on ice while the surgery is performed.
3. Place a 10% bleach waste container in the dedicated virus handling area to discard pipette tips, etc. that come in contact with the virus.
4. Draw a Wiretrol glass micropipette to a tip diameter of approximately 20 µm. Place a small drop of mineral oil on the blunt end of the micropipette and insert the wire plunger provided with the micropipettes.
5. Secure the micropipette in the micropump arm clamp.
6. Inject a mouse subcutaneously with buprenorphine at a dose of 0.5 mg/kg. Anesthetize the mouse with avertin by intraperitoneal injection at a dose of 200 mg/kg and trim the hair on the top of the head with surgical scissors, taking care to leave large enough margins to prevent hair clipping get into the incision.
7. Attach an electric blanket to the base of a stereotax to maintain a body temperature of 37 °C during surgery and secure the mouse in the stereotax.
8. Bathe the head with three alternating scrubs of ethanol and betadine to sterilize the area.
9. Place a drop of Tobradex eye ointment in each eye to keep eyes moist during surgery.
10. Make a midline incision on the head and peel back the skin to expose the skull.
11. Carefully remove the fascia from the skull using fine forceps.
12. Locate the area to be injected using stereotactic coordinates and mark the skull with a surgical pen.
13. Using a dental bur with a 1.4 mm bur, thin an area of ​​the skull approximately 2 mm in diameter until the skull fractures and divide the thinned area into multiple segments.
14. Keep the skull wet with sterile saline during this procedure.
15. Perform a craniotomy by gently removing the thinned skull segments with extra-fine-tipped forceps.

3. Injection preparation

1. With a KimWipe placed over the cap, open the virus tube and pipette 1.5 µl (for a 1 µl injection) of virus onto a small piece of parafilm.
2. Place the micropipette tip into the virus stock and manually pull back the plunger. If it is difficult to draw viral material into the micropipette, the tip can be easily enlarged by piercing a KimWipe with the micropipette to break a small portion of the glass.
3. Lower the micropump arm onto the plunger until a small amount of virus detaches from the micropipette tip. Remove this small drop with a cotton swab and dispose of the applicator in a waste container.
4. Apply a drop of mineral oil to the tip of the micropipette to prevent clogging when advancing the micropipette into the brain.

4. Virus injection

1. Using the stereotactic X and Y coordinates, position the micropipette over the area to be injected. In this experiment, the stereotactic coordinates used to locate the primary visual cortex are 2.7 mm posterior to the bregma and 2.5 mm lateral to the midline. Very slowly (about 1 mm/1 minute) lower the micropipette to the appropriate Z position.
2. Enter the desired injection parameters into the SYS Micro4 micropump control panel and start the injection. For this experiment, 1 microliter over 10 minutes is used as the injection rate.
3. After the injection is complete, leave the pipette for a minute or two to prevent the virus from escaping during the extraction. After this time, very slowly withdraw the micropipette from the brain (same speed as above).
4. Sew the scalp and seal with tissue glue. Inject the animal subcutaneously with buprenorphine at a dose of 0.1 mg/kg every 8-12 hours for the next 72 hours or as long as the animal shows signs of pain. Allow the animal to recover under a heat lamp until it is migratory and ready to return to its cage. Imaging experiments can be started after the desired incubation time (days or weeks after virus injection).

5. Cleaning

1. Rinse the micropipette with 10% bleach and discard in a puncture-resistant container.
2. Dispose of the waste container in the same manner as in Section 1.
3. Dispose of lab bench paper in the biowaste bin and clean all surfaces and instruments that may have been exposed to the virus with 10% bleach.
4. Unused virus can be refrozen, bearing in mind that repeated freeze/thaw cycles will cause virus degradation.

6. Representative Results

Illustration 1.Transduced neuron following injection of double-stranded adeno-associated virus serotype 1 (dsAAV S1) carrying green fluorescent protein (GFP) under the control of the CMV promoter. The cell body as well as the proximal and distal dendrites are clearly visible in still sections obtained with an epifluorescence microscope. Scale bar = 100 µm.

Figure 2.Typical labeling of an intracranial virus injection into the primary visual cortex with dsAAV S1, showing extent of virus spread and labeled neurons, glia, and processes. Scale bar = 250 µm.

Figure 3.Labeling of cells in the hippocampus with dsAAV S1. Scale bar = 250 µm. These numbers are adjusted from Loweryet al.2009¹

Virus-mediated gene delivery has great potential for studying neurological processes and treating brain diseases.^1,2,3. The great versatility of this technique can also be exploited to both fluorescently label cells for imagingin-vitrojlive⁴. Here we demonstrate a detailed procedure for the transduction of neurons and glial cells in the mouse visual cortex with a double-stranded adeno-association virus expressing enhanced green fluorescent protein.

Although this technique is relatively simple, there are a number of technical details that need to be considered. An important factor is the injection-induced tissue damage; Therefore, it is important to be very careful during surgery and when inserting/removing the micropipette. A failed surgical procedure could result in destruction of the structures to be imaged. Also, after loading the virus into the micropipette, care must be taken to eliminate air bubbles in the pipette tip to ensure the correct volume is injected. Dropping the micropipette slightly past the Z target coordinate and then retracting it to the correct position can prevent virus from leaking out of the injection site. When selecting an incubation period, care should be taken to allow sufficient time for expression of the viral gene. This varies depending on the virus used. The injection can produce an impaired immune response. In our experience, this swelling is generally confined to the needle path and can be avoided by imaging away from the injection site (approximately 50 μm or more). Finally, if brain sections are to be mounted on slides, the use of an anti-fading mounting medium to preserve fluorescence is suggested.

Intracranial injection of viral vectors has several technical advantages over other labeling techniques. By using stereotactic coordinates and modulating the injected volume, the fluorescent label can be precisely localized to an area of ​​interest. The amount of transduction can be altered by adjusting the volume injected, virus titer, or survival time, allowing for visualization of cell groups or single cells. In addition, the use of different viruses (e.g., lentivirus, herpesvirus, adenovirus) can also modulate the time and amount of fluorescent protein expressed, as well as target cell types. Overall, this technique can provide a very versatile labeling of different brain elements for imaging.