Name: Intracranial Injection of Adeno-associated Viral Vectors (Video) | JoVE Journal
Uploaded: 2010-11-17T18:00:00.000+00:00
Duration: 8 min 47 s
Description: Learn intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex. This method enables targeted cell type imaging in vivo with cost-effective advantages over transgenic approaches.

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5 minutes

This work was made possible by grants from the NIH (EY012977), a Career Award in the Biomedical Sciences from the Burroughs Wellcome Fund, the Whitehall Foundation, and the Sloan Foundation (A.K.M.).

1. Virus Handling and Storage
<ol>
<li>Proper protection and handling techniques should be chosen based on the biosafety level of the virus to be used. These practices can be found in Biosafety in Microbiological and Biomedical Laboratories 5th edition, available on the CDC website (<a href="http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm">http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm</a>). The use of AAV vectors is approved for Biosafety Level 1 (BSL-1). For the experiment shown here, a lab coat and gloves will be worn when handling the virus, in accordance with procedures for handling BSL-1 agents.</li>
<li>To preserve the activity of the virus it is best to divide it into small aliquots to avoid repeated freezing and thawing.</li>
<li>Prepare a Biosafety Cabinet (BSC) Class II for aliquoting virus by clearing the BSC of any unnecessary objects and sterilizing the surface with 70% EtOH. Place a beaker containing 10% bleach solution in the hood for collecting AAV-contaminated waste. Also placed in the BSC are sterile 0.5mL tubes and a container of dry ice.</li>
<li>Thaw viral stock on ice outside the BSC.</li>
<li>Vortex virus and open the tube inside the BSC.</li>
<li>Pipette the desired aliquot volume (e.g. 5 μL) into a 0.5mL tube, close the tube and place it in the dry ice to flash-freeze the virus.</li>
<li>When all virus has been aliquoted, dispose of the virus tube and pipette tips in the waste container containing 10% bleach.</li>
<li>Remove the waste container from the hood and add additional 10% bleach, then pour the bleach down the sink. Dispose of plastic waste in a biohazard container as instructed by your institute's biosafety officer.</li>
<li>Clean any equipment or surfaces that came in contact with the virus with 10% bleach. Discard gloves.</li>
<li>Store aliquots in -80°C freezer.</li>
</ol>
2. Surgery
<ol>
<li>Cover the surgical area with absorbent lab bench paper. Surgical tools should be sterilized and surgery done under aseptic conditions in accordance with your institution's biosafety and animal use guidelines.</li>
<li>Choose an area adjacent to the surgical area that will be dedicated to loading the micropipettes with virus, and cover it with absorbent lab bench paper. Place an aliquot of virus in a container of ice in this area, and allow the virus to thaw on ice as the surgery is being performed.</li>
<li>Setup a waste container of 10% bleach in the dedicated virus handling area for disposing of pipette tips, etc. that come in contact with the virus.</li>
<li>Pull a glass Wiretrol micropipette to a tip diameter of approximately 20 microns. Place a small drop of mineral oil on the blunt end of the micropipette and insert the wire plunger provided with the micropipettes.</li>
<li>Secure the micropipette in the clamp of the micropump arm.</li>
<li>Inject a mouse subcutaneously with buprenorphine at a dose of 0.5 mg/kg. Anesthetize the mouse with Avertin via intraperitoneal injection with a dose of 200 mg/kg and remove the hair from the top of the head with surgical scissors, making sure to leave margins large enough to prevent hair from entering the incision.</li>
<li>Attach a heating blanket to the base of a stereotax to maintain a body temperature of 37°C throughout the surgery and secure the mouse in the stereotax.</li>
<li>Bathe the head with three alternating scrubs of ethanol and betadine to sterilize the area.</li>
<li>Place a drop of Tobradex eye ointment on each eye to keep the eyes moist during surgery.</li>
<li>Make an incision down the midline of the head and pull the skin back to expose the skull.</li>
<li>Carefully remove the fascia from the skull using fine-tipped forceps.</li>
<li>Locate the area to be injected using stereotaxic coordinates and mark the skull with a surgical pen.</li>
<li>Using a dental drill with a 1.4mm burr, thin an area of the skull approximately 2mm in diameter until the skull cracks, dividing the thinned area into several segments.</li>
<li>Throughout this procedure, keep the skull moist with applications of sterile saline.</li>
<li>Perform a craniotomy by gently removing the thinned skull segments using extra-fine tipped forceps.</li>
</ol>
3. Injection Preparation
<ol>
<li>With a KimWipe placed over the lid, open the tube of virus and pipette 1.5uL (for a 1uL injection) of virus onto a small piece of parafilm.</li>
<li>Place the tip of the micropipette into the viral stock and manually withdraw the plunger. If difficulty is experienced drawing viral stock into the micropipette, the tip can be enlarged slightly be piercing a KimWipe with the micropipette to break a small portion of the glass.</li>
<li>Lower micropump arm onto plunger until a small amount of virus is dispelled from the tip of the micropipette. Remove this small drop with a cotton tip applicator and discard applicator in waste container.</li>
<li>Apply a drop of mineral oil to the tip of the micropipette to prevent clogging as the micropipette is lowered into the brain.</li>
</ol>
4. Virus Injection
<ol>
<li>Using X and Y stereotaxic coordinates, position the micropipette over the area to be injected. In this experiment, the stereotaxic coordinates used to locate primary visual cortex are 2.7mm posterior to bregma and 2.5mm lateral to the midline. Very slowly lower the micropipette (at an approximate rate of 1mm/1minute) to the proper Z position.</li>
<li>Enter the desired injection parameters into the SYS Micro4 micropump controller box and initiate the injection. For this experiment, 1 microliter over 10 minutes will be used as a rate of injection.</li>
<li>When the injection is finished, leave the pipette to rest for one-two minutes to prevent efflux of virus during removal. After this period, very slowly remove the micropipette from the brain (same rate as above).</li>
<li>Suture the scalp and seal it with tissue glue. Inject the animal subcutaneously with buprenorphine at a dose of 0.1 mg/kg every 8-12 hours over the next 72 hours, or as long as the animal is exhibiting signs of pain. Allow the animal to recover under a heat lamp until it is ambulating and ready to be returned to its cage. Imaging experiments can be initiated after the desired incubation time (days to weeks after viral injection).</li>
</ol>
5. Cleanup
<ol>
<li>Rinse the micropipette with 10% bleach and discard it in a sharps container.</li>
<li>Dispose of waste container in the same manner as Section 1.</li>
<li>Discard lab bench paper into biohazard bin and wipe down all surfaces and instruments that may have come in contact with the virus with 10% bleach.</li>
<li>Unused virus may be frozen again, keeping in mind that repeated freeze/thaw cycles cause degradation of the virus.</li>
</ol>

6. Representative Results
<img src="/files/ftp_upload/2140/2140fig1.jpg" alt="Neuron fluorescence microscopy; green dye highlights dendritic structures; cell imaging analysis." data-altupdatedat="1785211204000" data-alt="Figure 1"> 
Figure 1. Transduced neuron after injection of double-stranded adeno-associated virus serotype 1 (dsAAV S1) carrying green fluorescent protein (GFP) under control of the CMV promotor. The cell body as well as proximal and distal dendrites are clearly visible in fixed sections imaged using an epifluorescent microscope. Scale bar= 100 μm.
<img src="/files/ftp_upload/2140/2140fig2.jpg" alt="Fluorescence microscopy image of neural tissue highlighting neuron activity through GFP labeling." data-altupdatedat="1785211204000" data-alt="Figure 2"> 
Figure 2. Labeling typical of an intracranial virus injection in primary visual cortex using dsAAV S1 showing the extent of viral spread as well as labeled neurons, glia and processes. Scale bar= 250 μm.
<img src="/files/ftp_upload/2140/2140fig3.jpg" alt="Fluorescence microscopy image of neuron cells; visualizing cellular structures for neurological research." data-altupdatedat="1785211204000" data-alt="Figure 3"> 
Figure 3. Labeling of cells in hippocampus using dsAAV S1. Scale bar= 250 μm. These figures are adapted from Lowery et al. 20091

<ol><li>Kaplitt, M. G., Leone, P., Samulski, R. J., Xiao, X., Pfaff, D. W., O'Malley, K. L., During, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-term+gene+expression+and+phenotypic+correction+using+adeno-associated+virus+vectors+in+the+mammalian+brain%5BTitle%5D)+AND+%22Nat+Genet%22%5BJournal%5D)">Long-term gene expression and phenotypic correction using adeno-associated virus vectors in the mammalian brain.</a> Nat Genet. 8, 148-154 (1994).</li>
<li>Harding, T. C., Dickinson, P. J., Roberts, B. N., Yendluri, S., Gonzalez-Edick, M., Lecouteur, R. A., Jooss, K. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Enhanced+gene+transfer+efficiency+in+the+murine+striatum+and+an+orthotopic+glioblastoma+tumor+model%2C+using+AAV-7-+and+AAV-8-pseudotyped+vectors%5BTitle%5D)+AND+%22Hum+Gene+Ther%22%5BJournal%5D)">Enhanced gene transfer efficiency in the murine striatum and an orthotopic glioblastoma tumor model, using AAV-7- and AAV-8-pseudotyped vectors.</a> Hum Gene Ther. 17, 807-820 (2006).</li>
<li>Lo, W. D., Qu, G., Sferra, T. J., Clark, R., Chen, R., Johnson, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Adeno-associated+virus-mediated+gene+transfer+to+the+brain%3A+duration+and+modulation+of+expression%5BTitle%5D)+AND+%22Hum+Gene+Ther%22%5BJournal%5D)">Adeno-associated virus-mediated gene transfer to the brain: duration and modulation of expression.</a> Hum Gene Ther. 10, 201-213 (1999).</li>
<li>Lowery, R. L., Zhang, Y., Kelly, E. A., Lamantia, C. E., Harvey, B. K., Majewska, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Rapid+long-term+labeling+of+cells+in+the+developing+and+adult+rodent+visual+cortex+using+double-stranded+adeno-associated+viral+vectors%5BTitle%5D)+AND+%22Dev+Neurobiol%22%5BJournal%5D)">Rapid long-term labeling of cells in the developing and adult rodent visual cortex using double-stranded adeno-associated viral vectors.</a> Dev Neurobiol. 69, 674-688 (2009).</li>
</ol>

The animal and experimental protocols were approved by the University of Rochester University Committee on Animal Resources (UCAR) in accordance with the PHS Policy on Humane Care and Use of Laboratory Animals.

Virally-mediated gene delivery holds great potential for the study of neurological processes and treatment of brain disorders1,2,3. The great versatility of this technique can also be exploited to fluorescently label cells for imaging both in vitro and in vivo4. Here we demonstrate a detailed procedure for the transduction of neurons and glia in mouse visual cortex using a double-stranded adeno-association virus expressing enhanced green fluorescent protein.
<p class="jove_con...

<table><tbody><tr><td>St&amp;#8211;lting Mouse and Neonatal Rat Adaptor</td><td>Stoelting Co.</td><td>51625</td><td>Regular stereotax for securing animals for surgery may be substituted</td></tr><tr><td>Extra Fine Bonn Scissors, 8.5cm, straight tip, cutting edge 13mm</td><td>Fine Science Tools</td><td>14084-08</td><td></td></tr><tr><td>Eye Dressing Forceps, 10cm, tip width 0.5mm, curved</td><td>Fine Science Tools</td><td>11152-10</td><td></td></tr><tr><td>Dumont #5/45 Forceps- Dumoxel Standard Tip, 11cm, angled</td><td>Fine Science Tools</td><td>11251-35</td><td>Extra-fine tipped forceps for performing craniotomy</td></tr><tr><td>Standard Pattern Forceps, straight, 2.5mmx1.35mmtip, 12cm</td><td>Fine Science Tools</td><td>11000-12</td><td></td></tr><tr><td>Microtorque Control Box and Tech2000 handpiece</td><td>Ram Products, Inc.</td><td>TECH2000ON/OFF</td><td>Dental drill</td></tr><tr><td>Micro Drill Stainless Steel Burrs 1.4mm tip diameter</td><td>Fine Science Tools</td><td>19008-14</td><td></td></tr><tr><td>Wiretrol micropipettes, to deliver 1-5 Ul</td><td>VWR international</td><td>5-000-1001 or 53480-287</td><td></td></tr><tr><td>Mineral oil</td><td>VWR international</td><td>29447-338</td><td></td></tr><tr><td>Manual Micromanipulator and Tilting Base (right-handed)</td><td>World Precision Instruments, Inc.</td><td>M3301-M3-R</td><td>Used for determining stereotaxic co-ordinates</td></tr><tr><td>UltraMicroPump (UMP3) (one) with SYS-Micro4 Controller</td><td>World Precision Instruments, Inc.</td><td>UMP3-1</td><td></td></tr><tr><td>Sutures</td><td>VWR international</td><td>95056-952</td><td></td></tr><tr><td>P-97 Flaming/Brown Micropipette Puller</td><td>Sutter Instrument Co.</td><td>P-97</td><td></td></tr><tr><td>Tobradex</td><td></td><td></td><td>Available from your institution&amp;rsquo;s veterinary services</td></tr></tbody></table>

intracranial injection of adeno-associated viral vectors

Intracranial Injection of Adeno-associated Viral Vectors

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific subsets of cells in different brain regions both in vivo and in brain sections. Unlike the injection of fluorescent dyes, viral labeling offers targeting of individual cell types and is less expensive and time consuming than establishing transgenic mouse lines. In this technique, an adeno-associated viral (AAV) vector is injected intracranially using stereotaxic coordinates, a micropipette and an automated pump for precise delivery of AAV to the desired area with minimal damage to the surrounding tissue. Injection parameters can be tailored to individual experiments by adjusting the animal age at injection, injection location, volume of injection, rate of injection, AAV serotype and the promoter driving gene expression. Depending on the conditions chosen, virally-induced transgene expression can allow visualization of groups of cells, individual cells or fine cellular processes, down to the level of dendritic spines. The experiment shown here depicts an injection of double-stranded AAV expressing green fluorescent protein for the labeling of neurons and glia in the mouse primary visual cortex.

Here we present the intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex.

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific ...

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.

intracranial-injection-of-adeno-associated-viral-vectors

Research

JoVE Journal

Neuroscience

Source: Pereira, M. et. al., In Vivo Direct Reprogramming of Resident Glial Cells into Interneurons by Intracerebral Injection of Viral Vectors. J. Vis. Exp. (2019)This video demonstrates stereotaxic injection of an adeno-associated virus into the cerebral cortex of a transgenic mouse to deliver Cre-dependent neuronal reprogramming and fluorescent reporter genes into glial cells.

Stereotaxic Injection of Viral Vectors for Glial Cell Reprogramming in a Mouse Model

Source: Li, D., et al. Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice. J. Vis. Exp. (2019)This video demonstrates the procedure for the intrathecal injection of adeno-associated viruses (AAV) into the cerebrospinal fluid (CSF) of an awake mouse, covering syringe preparation, spinal alignment, and precise needle insertion. This method validates targeted gene delivery, supporting its potential for CNS therapies.

Intrathecal Delivery of Viruses in a Mouse Model for Treating Central Nervous System Diseases

Source: Grames, M. S., et al. Methods and Tips for Intravenous Administration of Adeno-associated Virus to Rats and Evaluation of Central Nervous System Transduction. J. Vis. Exp. (2017)This video demonstrates the tail vein injection of recombinant adeno-associated virus (AAV) in rats. After injection into the lateral tail vein, the virus enters the bloodstream, circulates through the heart, and reaches the brain, where it enters neurons and facilitates target protein production for...

 Intravenous Administration of Recombinant Adeno-Associated Virus Vectors in a Rat Model 

Here we demonstrate a technique for widespread neuronal transduction by intraventricular injection of adeno-associated virus into the neonatal mouse brain.  This method provides a rapid and easy way to attain lifelong expression of virally-delivered transgenes.

Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction

The brain is a unique site with qualities that are not well represented by in vitro or ectopic analyses. Orthotopic mouse models with reproducible location and growth characteristics can be reliably created with intracranial injections using a stereotaxic fixation instrument and a low pressure syringe pump.

Creating Anatomically Accurate and Reproducible Intracranial Xenografts of Human Brain Tumors

Methods for a wide-scale central nervous system gene delivery in the rat are covered. In this example, the purpose is to mimic a disease that affects the entire spinal cord. The widespread transduction can be used to deliver a therapeutic protein to the CNS from a one-time, peripheral administration.

Methods and Tips for Intravenous Administration of Adeno-associated Virus to Rats and Evaluation of Central Nervous System Transduction

Here we present a direct intrathecal injection technique using 1% lidocaine hydrochloride in a viral solution to ensure efficient adeno-associated virus delivery to small animals and establish a scoring system to predict transduction efficiency in the central nervous system according to the degree of transient weakness induced by lidocaine.

Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice

In this manuscript, subconjunctival injection is demonstrated as a valid vector delivery method for ocular tissues in mice using an injection system consisting of an infusion/withdrawal syringe pump and a gastight removable syringe coupled with microinjection needles. This injection system is also adaptable for other intraocular administration routes.

Subconjunctival Administration of Adeno-associated Virus Vectors in Small Animal Models

A surgical procedure is described to perform injections into the lumbar cistern of the juvenile rat. This approach has been used for the intrathecal delivery of gene therapy vectors, but it is anticipated that this approach can be used for a variety of therapeutics, including cells and drugs.

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

This manuscript describes a detailed protocol for the generation and purification of adeno-associated viral vectors using an optimized heparin-based affinity chromatography method. It presents a simple, scalable, and cost-effective approach, eliminating the need for ultracentrifugation. The resulting vectors exhibit high purity and biological activity, proving their value in preclinical studies.

Isolation of Adeno-Associated Viral Vectors Through a Single-Step and Semi-Automated Heparin Affinity Chromatography Protocol

A new technique for widespread delivery of adeno-associated virus that uses subarachnoid virus infusion is described. This method not only ensures widespread transduction of mouse neocortical neurons in superficial layers but also results in selective expression of the transgene in layer five pyramidal neurons, even when using a non-selective promoter.

Widespread Transduction of Mouse Neocortical Neurons by Subarachnoid Injection of AAV2

Learn intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex. This method enables targeted cell type imaging in vivo with cost-effective advantages over transgenic approaches.

Intracranial Injection of Adeno-associated Viral Vectors

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