The authors thank the Vector Core at the University of North Carolina for providing the scAAV8-GFP vectors used in this study, the CGIBD Histology Core, and the laboratory of Dr. Brian C. Gilger for their assistance with the clinical assessment aspects of this study. This study was supported by the Pfizer-NC Biotech Distinguished Postdoctoral Fellowship and a Career Development Award from the American Society of Gene &#38; Cell Therapy and the Cystic Fibrosis Foundation.&#160;The content is solely the responsibility of the authors and does not necessarily represent the official views of the American Society of Gene &#38; Cell Therapy or the Cystic Fibrosis Foundation.

All animal procedures were performed in accordance with the regulations of the Institutional Animal Care and Use Committee at the University of North Carolina at Chapel Hill. The use of AAV vectors is a Biosafety Level 1 biohazard risk. Wear proper personal protective equipment, including a lab coat, gloves, and goggles when handling AAV. For the experiment described herein, a recombinant AAV vector packaged with the serotype 8 capsid and encoding a generic ubiquitous cytomegalovirus (CMV) promoter controlling the expression of green fluorescence protein (GFP) was utilized.
1. AAV vector handling and storage
<ol>
	<li>Store the virus in a -80 &#176;C freezer in 100 &#181;L aliquots in siliconized or low-retention microcentrifuge tubes.</li>
	<li>Thaw all vector stock solutions on ice before use. 
	&#8203;NOTE: Dyes such as sodium fluorescein solution (at a final concentration of 0.1-2%) are often mixed with the AAV vectors to visualize the injected solution. Additionally, visualization of injected solutions helps detect air bubbles and monitor AAV distribution and/or leakage after injection.</li>
</ol>
2. Subconjunctival (SCJ) injection
<ol>
	<li>Assemble the injection system.
	<ol>
		<li>To assemble the injection system, place a stereomicroscope and a syringe pump in a biosafety cabinet. 
		NOTE: An infusion pump is needed to perform injections with high precision. Herein, a Standard Infuse/Withdraw Programmable Syringe Pump is utilized (see the Table of Materials), which includes a tight grip and a secure syringe clamp for syringes ranging in volume from 0.5 &#181;L to 60 mL. This pump also offers enhanced flow performance with high accuracy and smooth flow rates from 1.28 pl/min to 88.28 ml/min.</li>
		<li>Cut the polyethylene tubing to a length of approximately 50 cm (see the Table of Materials).</li>
		<li>Insert the hub end of a 36 G needle into one of the ends of the tubing. 
		NOTE: Slide the needle hub end into the tube for ~3 mm to ensure that no leakage occurs. The 36 G needle is used for the subsequent SCJ injection. Needles ranging between 32 G and 36 G are the most commonly used sizes for SCJ injections. The use of a hemostat to assist this step is highly recommended to avoid the potential risk of sharps injury.</li>
		<li>Fill a disposable 3 mL syringe with sterile water; insert this disposable syringe into the side of the tube opposite the needle and flush the water throughout the tubing/needle. Repeat this step with 70% alcohol.</li>
		<li>Repeat step 2.1.4 three more times, alternating flushes with sterile water and 70% alcohol, to disinfect the tubing and ensure no leaks, clogs, or damage are observed throughout the tubing.</li>
		<li>Use the disposable 3 mL syringe to fill the tubing with sterile water and leave the tubing attached to the disposable syringe.</li>
		<li>Place a piece of parafilm on the bench surface and add a pool of sterile water to it (~1 mL). Submerge the portion of the tubing connected to the needle into the pool of sterile water. Pull the disposable syringe out from the tube opening at the opposite end to prevent any air from entering the tubing/needle system upon removal of the syringe. Leave the portion of the tubing connected to the needle submerged in the pool of water. 
		NOTE: Perform procedures 2.1.4 to 2.1.7 in a laminar hood.</li>
		<li>Fill a 10 &#181;L Hamilton syringe/needle with sterile water and avoid air in the syringe. Connect the Hamilton syringe/needle to the remaining open end of the tubing by submerging the tubing and needle tip of the Hamilton syringe into the pool of sterile water on the parafilm.</li>
		<li>Press the fast reverse button on the pump screen to move the pusher block to the approximate length of the syringe. Unscrew the bracket clamping knobs to loosen the retaining brackets on the pusher and the syringe holder blocks. Load the Hamilton syringe onto the syringe holder block and secure the syringe following the manufacturer&#39;s instructions. 
		NOTE: To secure the syringe, the syringe barrel clamp should be tight against the syringe barrel; however, do not overtighten, especially when using glass syringes.&#160;The syringe plunger should be secured by the pusher block retaining bracket.</li>
		<li>Adjust the parameters in the pump settings screen.
		<ol>
			<li>Press the Force button, and set the force level at 30%. Accept the changes to go back to the settings screen.</li>
			<li>Press the quick start button and select Method | Infuse/withdraw.</li>
			<li>For the Syringe, select Hamilton 1700, glass, 10 &#181;L. Select the infusion and withdraw rate and the injection volume. 
			NOTE: The Force level is set depending on the syringe type/material/capacity/manufacturers; see factory manufacturer instructions for suggested force for each syringe. The injection speed used in this experiment was 200 nL/s. SCJ injections are relatively safe, and there is less of a concern for the induction of elevated intraocular pressure (IOP) resulting from the injection. A slower injection speed is often desirable for certain applications to avoid reflux into the needle and maintain consistency in injections among animals.</li>
		</ol>
		</li>
		<li>Eject the water from the Hamilton syringe but leave the tubing and injection needle full of water. Slightly pull back on the Hamilton syringe by pressing the Reverse button to introduce a small air bubble in the tubing/needle. 
		NOTE: The air bubble will serve as a barrier between the water in the tube and the therapeutic drug (in this case, AAV), ensuring the accuracy of the administered dose.</li>
		<li>Withdraw the virus by placing the injection needle into an aliquot of the virus stock. Ensure that a visible air bubble remains between the virus and the water in the tubing. 
		NOTE: AAV vectors can bind to the plastic tubing and metal needle, leading to a loss of virus and/or inaccurate dosing regimens. Thus, to ensure rigor, reproducibility, and an accurate dose of AAV, precoating the surfaces that subsequently come into contact with the AAV is recommended. To coat the tubing/needle system with the virus, draw the viral vector solution into the tubing/needle and incubate it at room temperature for 10 min to allow for saturation of virus binding to the wall of the needle and/or tubing. Discard the virus.</li>
	</ol>
	</li>
	<li>Virus injection
	<ol>
		<li>Anesthetize the mouse with inhaled anesthesia (isoflurane) or intraperitoneal injection of ketamine/xylazine/acepromazine. Confirm the surgical plane of anesthesia by a lack of response to firm toe pinches. 
		NOTE: Use female and/or male C57BL/6J or BALB/c mice that are at least 6 weeks old. Ketamine/xylazine/acepromazine doses are as follows: ketamine at 70&#8201;mg/kg, xylazine at 7&#8201;mg/kg, and acepromazine at 1.5&#8201;mg/kg.</li>
		<li>Apply topical anesthesia to the eye that will receive the injection. 
		NOTE: Use 0.1% proparacaine hydrochloride and/or tetracaine hydrochloride ophthalmic solution (0.5%) for topical anesthesia.</li>
		<li>Apply topical ointment to the other eye that will not receive an injection to prevent dryness and injury.</li>
		<li>Position the mouse on the microscopic stage and expose the mouse eye under the stereomicroscope.</li>
		<li>Place two fingers on the eyelid and slightly pull it away from the mouse&#39;s eye to expose the conjunctiva, which is the inner membrane connecting the eyelid to the sclera.</li>
		<li>Grab the conjunctiva with forceps.</li>
		<li>Release the eyelid and hold the needle with the bevel facing upwards using the dominant hand.</li>
		<li>Insert the needle into the conjunctiva. Insert the needle until the bevel is completely covered by the conjunctival membrane. Lay the needle against the globe. 
		NOTE: As the conjunctiva is a clear membrane, the needle tip/bevel is easily visible.</li>
		<li>Start the injection by pressing the Start button using the footswitch. 
		NOTE: The movement of the air and the Hamilton plunger are synchronized; any delay indicates excess air in the injecting system or possibly a loose connection between the tubing, needle, and/or syringe components.</li>
		<li>After the injection is finished, hold the needle in place for 10 s before withdrawing the needle from the conjunctiva to decrease the chances of backflow. 
		NOTE: It is common for a bleb to appear at the site of the SCJ injection. Such blebs normally resolve completely within a few hours post injection.</li>
		<li>Put a drop of the topical lubricant gel on the mouse&#39;s eyes to prevent ocular dryness/injury, and then place the mouse on a heating pad to recover.</li>
		<li>Perform ocular examinations such as tear production, IOP, and slit-lamp examination in conjunction with corneal fluorescein staining to assess ocular abnormities following the injection. 
		NOTE: Tear production is measured by a Phenol Red Thread test, and a tonometer is often used to examine the IOP of the mouse eye. It is reported that some intraocular injections, such as intravitreal injections, might result in a significant increase in IOP; however, IOP changes after SCJ injection are not obvious13,22,23,24.</li>
	</ol>
	</li>
	<li>AAV biodistribution and transduction efficiency examination following subconjunctival injection
	<ol>
		<li>To investigate the viral genome biodistribution and/or transduction profile of AAV vectors delivered via SCJ, euthanize the mice by AVMA approved method. 
		NOTE: In this experiment, the mice were sacrificed 8 weeks post injection.</li>
		<li>For biodistribution and transgene expression in targeted ocular compartments, dissect the relevant tissue of interest such as the eyelids, cornea, conjunctiva, eye muscle, retina, and optic nerve. Flash-freeze all tissues and store at -80 &#176;C. To examine whole-body AAV biodistribution, collect organs such as the submandibular lymph nodes and liver, and flash-freeze and store them at -80 &#176;C.</li>
		<li>Using a DNA/RNA extraction kit, collect gDNA and RNA from the same sample to examine transgene expression and AAV biodistribution, respectively. If only vector biodistribution is desired, use a DNA extraction kit to extract gDNA.</li>
		<li>Perform standard qPCR and RT-qPCR to determine the AAV vector biodistribution and cDNA abundance using vector transgene-specific primers/probes13,25.</li>
		<li>For histology analysis, fix the eyes, embed them in paraffin, and section them at a thickness of 5 &#181;m. Perform standard immunofluorescence staining to reveal transgene expression26.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to disclose.

AAV-mediated gene therapy holds great potential for the treatment of ocular diseases. Current ocular gene therapy relies on two major local administration routes, intravitreal and subretinal injections. Unfortunately, both routes are invasive and can cause serious complications, including retinal detachment, cataract formation, and endophthalmitis. Thus, the investigation of relatively less invasive routes, such as SCJ injection, is of great interest.
Although this technique is relatively stra...

Ocular diseases encompass a broad range of both genetic and acquired disorders. In 2015, an estimated 36 million people were legally blind worldwide, and over 1 billion people suffer from at least some level of visual impairment, highlighting the need to scale up alleviation efforts at all levels1. The main methods for delivering ocular medications include both topical and local administration, such as eye drops or subconjunctival (SCJ), intracameral, intravitreal, and subretinal injections. Although noninvasive topical therapy is the most common delivery method for ophthalmic drugs and is extensively used for many anterior segment disorders, the presence of corneal anatomical barriers presents a challenge for the bioavailability, biodistribution, and efficacy of topically administrated substances, suggesting that it may not be the best candidate treatment route for many diseases of the inner eye. Local injection into the specific ocular compartment affected by the disease is likely to be a more effective and targeted drug delivery approach2. However, adverse effects resulting from repeated injections can complicate administration strategies. Ideally, a therapy should maintain long-term therapeutic efficacy following a single administration. Thus, gene therapy is a promising option for minimizing the number of required injections and providing sustained transgene expression for the treatment of ocular disease3,4.
Numerous viral and nonviral vectors are available for gene therapy; however, AAV vectors are of high interest due to their excellent safety profile. AAV is a small, single-stranded, non-enveloped DNA virus that was initially discovered as a contaminant of an adenovirus preparation in 1965 by Atchison et al.5,6 AAV was subsequently engineered as an efficient viral vector for gene delivery in the 1980s and has become the gene therapy vector of choice for many diseases, including ocular disorders, over the last few decades. The most notable of these is the first commercially available gene therapy drug, voretigene neparvovec, which was approved by the United States Food and Drug Administration to treat Leber&#39;s Congenital Amaurosis, a rare posterior eye disease. Although voretigene neparvovec has successfully overcome barriers to clinical development, challenges remain for the commercialization of additional ocular gene therapies. For example, voretigene neparvovec is administered to patients who retain viable retinal cells via subretinal injection. Thus, patients with more advanced forms of the disease who lack viable retinal cells are not eligible for treatment, as it would provide no clinical benefit. In addition, known complications associated with the subretinal injection procedure were observed, including eye inflammation, cataracts, retinal tearing, maculopathy, and pain7,8. Other concerns related to this procedure include the possibility of hemorrhage, retinal detachment, endophthalmitis, and revocation of the ocular immune privileged status through eye tissue destruction9,10,11,12. Thus, efforts to explore less invasive gene delivery routes such as SCJ injection have become increasingly important13,14,15,16,17.
The conjunctiva is a thin membrane containing 3-5 layers of cells and connecting the anterior eye to the interior eyelid. SCJ injections are used clinically for ophthalmic drug delivery to both the anterior and/or posterior segments of the eye for the treatment of ocular diseases such as age-related macular degeneration, glaucoma, retinitis, and posterior uveitis18,19. They are relatively simple to perform, employed routinely for ophthalmic drug delivery in an outpatient setting20, somewhat painless, do not compromise ocular immune privilege, and allow administered drugs to spread through a large periorbital region that encompasses the optic nerve. Hence, SCJ&#160;injections are an attractive route of administration for AAV gene therapy applications. Natural AAV serotypes administered via SCJ injection in mice have previously been characterized for safety, transduction efficiency, serum immunogenicity, biodistribution, and tissue specificity13,16,21. These data demonstrated that gene delivery to individual ocular tissues via SCJ administration is a formal possibility.
This paper describes a simple and adaptable protocol for SCJ injection to deliver AAV vectors in a mouse model. To ensure the reproducibility of this approach, an injection system consisting of a stereomicroscope, a programmable infusion/withdrawal syringe pump (which allows consistent and precise injection speed and pressure), and a gastight removable syringe coupled with microinjection needles is described. This system is adaptable for other intraocular administration routes such as intrastromal, intracameral, intravitreal, and subretinal injections in small animals. In addition, a fluorescein dye is often utilized to allow for visualization of the AAV injection site. The key practical steps in the administration route, setup for the injection platform, preparation of the injection, and tips from direct experience will be discussed in detail. Finally, common validation techniques for confirmation of AAV delivery to the desired tissues will be briefly discussed.

<table><tbody><tr><td>36 G NanoFil Needles</td><td>World Precision Instruments</td><td>NF36BV-2</td><td></td></tr><tr><td>AAV vector</td><td>&amp;nbsp; University of North Carolina at Chapel Hill</td><td>&amp;nbsp; /</td><td></td></tr><tr><td>Acepromazine</td><td>Henry Schein</td><td>NDC 11695-0079-8</td><td></td></tr><tr><td>anti-GFP antibody</td><td>AVES labs Inc.</td><td></td><td></td></tr><tr><td>Digital camera</td><td>Cannon</td><td>Cannon EOS T5i</td><td></td></tr><tr><td>DNA/RNA extraction kit</td><td>Qiagen</td><td>80204</td><td></td></tr><tr><td>&amp;nbsp;Forceps</td><td>Fine Science Tools</td><td>F6521</td><td></td></tr><tr><td>Hamilton syringe</td><td>Hamilton</td><td>7654-01</td><td></td></tr><tr><td>India ink</td><td>StatLab</td><td>NC9903975</td><td></td></tr><tr><td>Ketamine hydrochloride injection solution</td><td>Henry Schein</td><td>NDC 0409-2051-05</td><td></td></tr><tr><td>Moisture-resistant film</td><td>Parafilm</td><td>807-6</td><td></td></tr><tr><td>Polyethylene tubing</td><td>Becton Dickinson and Company</td><td>427401</td><td></td></tr><tr><td>Proparacaine 0.1%</td><td>Bausch Health US</td><td>NDC 24208-730-06</td><td></td></tr><tr><td>Rebound tonometer</td><td>Tonovet</td><td>/</td><td></td></tr><tr><td>Sodium fluorescein solution</td><td>Sigma-Aldich</td><td>46960</td><td></td></tr><tr><td>Standard Infuse/Withdraw Pump 11 Pico Plus Elite Programmable Syringe Pump</td><td>Harvard Bioscience</td><td>70-4504</td><td></td></tr><tr><td>Stereo microscopye</td><td>Leica</td><td>Mz6</td><td></td></tr><tr><td>Tetracaine Hydrochloride Ophthalmic Solution 0.5%</td><td>Bausch and Lomb</td><td>Rx only</td><td></td></tr><tr><td>Topical ointment</td><td>GenTeal</td><td>NDC 0078-0429-47</td><td></td></tr><tr><td>Xylazine</td><td>Akorn</td><td>NDC 59399-110-20</td><td></td></tr><tr><td>Zone-Quick Phenol Red Thread Box 100 Threads</td><td>ZONE-QUICK</td><td>PO6448</td><td></td></tr></tbody></table>

subconjunctival administration of adeno-associated virus vectors in small animal models

Ocular diseases include a wide range of inherited genetic and acquired disorders that are appealing targets for local drug delivery due to their relative ease of accessibility via multiple administration routes. Subconjunctival (SCJ) injections offer advantages over other intraocular administration routes as they are simple, safe, and usually performed in an outpatient setting. SCJ injections in small animals usually require the assistance of an operating microscope due to the size of the eye. Previous work has demonstrated that SCJ injection of specific adeno-associated virus (AAV) serotypes is a valid gene delivery strategy for targeted transduction of the ocular surface, eye muscle, cornea, and optic nerve, providing a potential approach for the treatment of many ocular diseases.
Herein, a detailed protocol is presented for SCJ injections in a mouse model using an injection system consisting of a programmable infusion/withdrawal syringe pump (which allows for consistent and precise injection speed and pressure) and a gastight removable syringe coupled with microinjection needles. The injection system is also adaptable for other intraocular administration routes such as intrastromal, intracameral, intravitreal, and subretinal injections in small animals. Although the delivery of adeno-associated viral vectors for ocular gene therapy studies is described, the protocol herein can also be adapted for a variety of ophthalmic solutions in small animal models. The key practical steps in the administration route, setup for the injection platform, preparation of the injection, and tips from direct experience will be discussed in detail. In addition, common validation techniques for AAV delivery confirmation to the desired tissues will also be briefly discussed.

Solution injected into the subconjunctival space presents as a bleb depending on the injection volume. 
In this experiment, 7 &#181;L of AAV (7 &#215; 109 viral genomes (vg)/eye) mixed with fluorescein at a final concentration of 0.1% was injected with a 36 G needle under a stereomicroscope, and the injection speed/pressure was held constant using a programmable syringe pump at 1 &#181;L/s. A bleb can appear upon injection (arrow). A microscopic view of AAV vector administration to the mu...

Watch this Scientific Journal Video about Subconjunctival Administration of Adeno-associated Virus Vectors in Small Animal Models at JoVE.com

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

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.

Ocular diseases include a wide range of inherited genetic and acquired disorders that are appealing targets for local drug delivery due to their relative ...

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Research

JoVE Journal

Medicine

2.8K Views.  University of North Carolina at Chapel Hill. 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. 

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

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 ...

Neuroscience

Using Adeno-associated Virus as a Tool to Study Retinal Barriers in Disease

M&#252;ller cells are the principal glial cells of the retina. Their end-feet form the limits of the retina at the outer and inner limiting membranes (ILM), and in conjunction with astrocytes, pericytes and endothelial cells they establish the blood-retinal barrier (BRB). BRB limits material transport between the bloodstream and the retina while the ILM acts as a basement membrane that defines histologically the border between the retina and the vitreous cavity. Labeling M&#252;ller cells is particularly relevant to study the physical state of the retinal barriers, as these cells are an integral part of the BRB and ILM. Both BRB and ILM are frequently altered in retinal disease and are responsible for disease symptoms.
There are several well-established methods to study the integrity of the BRB, such as the Evans blue assay or fluorescein angiography. However these methods do not provide information on the extent of BRB permeability to larger molecules, in nanometer range. Furthermore, they do not provide information on the state of other retinal barriers such as the ILM. To study BRB permeability alongside retinal ILM, we used an AAV based method that provides information on permeability of BRB to larger molecules while indicating the state of the ILM and extracellular matrix proteins in disease states. Two AAV variants are useful for such study: AAV5 and ShH10. AAV5&#160;has a natural tropism for photoreceptors but it cannot get across to the outer retina when administered into the vitreous when the ILM is intact (i.e., in wild-type retinas). ShH10 has a strong tropism towards glial cells and will selectively label M&#252;ller glia in both healthy and diseased retinas. ShH10 provides more efficient gene delivery in retinas where ILM is compromised. These viral tools coupled with immunohistochemistry and blood-DNA analysis shed light onto the state of retinal barriers in disease.

To investigate the blood-retinal barrier permeability and the inner limiting membrane integrity in animal models of retinal disease, we used several adeno-associated virus (AAV) variants as tools to label retinal neurons and glia. Virus mediated reporter gene expression is then used as an indicator of retinal barrier permeability.

M&#252;ller cells are the principal glial cells of the retina. Their end-feet form the limits of the retina at the outer and inner limiting membranes ...

Limbal Approach-Subretinal Injection of Viral Vectors for Gene Therapy in Mice Retinal Pigment Epithelium

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in retinopathies using non-viral and viral gene delivery to the retina and retinal pigment epithelium (RPE). Subretinal injection is the best approach to deliver viral vectors directly to RPE cells. Before the clinical trial of a gene therapy, it is inevitable to validate the efficacy of the therapy in animal models of various retinopathies. Thus, subretinal injection in mice becomes a fundamental technique for an ocular gene therapy. In this protocol, we provide the easy and replicable technique for subretinal injection of viral vectors to experimental mice. This technique is modified from the intravitreal injection, which is widely used technique in ophthalmology clinics. The representative results of RPE/choroid/scleral complex flat-mount will help to understand the efficacy of this technique and adjust the volume and titer of viral vectors for the extent of gene transduction.

Subretinal injection is a surgical technique for effective gene delivery to retinal pigment epithelium in the mouse eye. Here we describe an easy and replicable method for subretinal injection of viral vectors to retinal pigment epithelium in experimental mice.

The eye is a small and enclosed organ which makes it an ideal target for gene therapy. Recently various strategies have been applied to gene therapy in ...

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

Adeno-associated virus (AAV) vectors are a key reagent in the neurosciences for clustered regularly interspaced short palindromic repeats (CRISPR), optogenetics, cre-lox targeting, etc. The purpose of this manuscript is to aid the investigator attempting expansive central nervous system (CNS) gene transfer in the rat via tail vein injection of AAV. Wide-scale expression is relevant for conditions with widespread pathology, and a rat model is significant due to its greater size and physiologic similarities to humans compared to mice. In this example application, a wide-scale neuronal transduction is used to mimic a neurodegenerative disease that affects the entire spinal cord, amyotrophic lateral sclerosis (ALS). The efficient wide-scale CNS transduction can also be used to deliver therapeutic protein factors in pre-clinical studies. After a post-injection expression interval of several weeks, the effects of the transduction are evaluated. For a green fluorescent protein (GFP) control vector, the amount of GFP in the cerebellum is estimated quickly and reliably by a basic imaging program. For motor disease phenotypes that are induced by the ALS related protein transactive response DNA-binding protein of 43 kDa (TDP-43), the deficits are scored by escape reflex and rotarod. Beyond disease modeling and gene therapy, there are diverse potential applications for the wide-scale gene targeting described here. The expanded use of this method will aid in expediting hypothesis testing in the neurosciences and neurogenetics.

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.

Adeno-associated virus (AAV) vectors are a key reagent in the neurosciences for clustered regularly interspaced short palindromic repeats (CRISPR), ...

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

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, and excellent transduction efficacy in the CNS.&#160;Previous studies have demonstrated the therapeutic potency of AAV-delivered gene therapy in neurodegenerative disorders by IT administration. However, high rates of unpredictable failure due to the technical limitation of IT administration in small animals have been reported. Here, we established a scoring system to indicate the success extent of lumbar puncture in small animals by adding 1% lidocaine hydrochloride into the injection solution. We further show that the extent of transient weakness following injection can predict the transduction efficiency of AAV. Thus, this IT injection method can be used to optimize therapeutic trials in mouse models of CNS diseases that afflict wide regions of the CNS.

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.

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, ...

Purification of AAV Vectors by Single-Step Heparin Affinity Chromatography

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

Adeno-associated virus (AAV) has become an increasingly valuable vector for in vivo gene delivery and is currently undergoing human clinical trials. However, the commonly used methods to purify AAVs make use of cesium chloride or iodixanol density gradient ultracentrifugation. Despite their advantages, these methods are time-consuming, have limited scalability, and often result in vectors with low purity. To overcome these constraints, researchers are turning their attention to chromatography techniques. Here, we present an optimized heparin-based affinity chromatography protocol that serves as a universal capture step for the purification of AAVs.
This method relies on the intrinsic affinity of AAV serotype 2 (AAV2) for heparan sulfate proteoglycans. Specifically, the protocol entails the co-transfection of plasmids encoding the desired AAV capsid proteins with those of AAV2, yielding mosaic AAV vectors that combine the properties of both parental serotypes. Briefly, after the lysis of producer cells, a mixture containing AAV particles is directly purified following an optimized single-step heparin affinity chromatography protocol using a standard fast protein liquid chromatography (FPLC) system. Purified AAV particles are subsequently concentrated and subjected to comprehensive characterization in terms of purity and biological activity. This protocol offers a simplified and scalable approach that can be performed without the need for ultracentrifugation and gradients, yielding clean and high viral titers.

Author Spotlight: Improved Method for Production and Purification of Adeno-Associated Viral Vectors

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.

Adeno-associated virus (AAV) has become an increasingly valuable vector for in vivo gene delivery and is currently undergoing human clinical trials. ...

Bioengineering

Digital Droplet PCR Method for the Quantification of AAV Transduction Efficiency in Murine Retina

Many retinal cell biology laboratories now routinely use Adeno-associated viruses (AAVs) for gene editing and regulatory applications. The efficiency of AAV transduction is usually critical, which affects the overall experimental outcomes. One of the main determinants for transduction efficiency is the serotype or variant of the AAV vector. Currently, various artificial AAV serotypes and variants are available with different affinities to host cell surface receptors. For retinal gene therapy, this results in varying degrees of transduction efficiencies for different retinal cell types. In addition, the injection route and the quality of AAV production may also affect the retinal AAV transduction efficiencies. Therefore, it is essential to compare the efficiency of different variants, batches, and methodologies. The digital droplet PCR (dd-PCR) method quantifies the nucleic acids with high precision and allows performing absolute quantification of a given target without any standard or a reference. Using dd-PCR, it is also feasible to assess the transduction efficiencies of AAVs by absolute quantification of AAV genome copy numbers within an injected retina. Here, we provide a straightforward method to quantify the transduction rate of AAVs in retinal cells using dd-PCR. With minor modifications, this methodology can also be the basis for the copy number quantification of mitochondrial DNA as well as assessing the efficiency of base editing, critical for several retinal diseases and gene therapy applications.

This protocol presents how to quantify AAV transduction efficiency in mouse retina using digital droplet PCR (dd-PCR) together with small scale AAV production, intravitreal injection, retinal imaging, and retinal genomic DNA isolation.

Many retinal cell biology laboratories now routinely use Adeno-associated viruses (AAVs) for gene editing and regulatory applications. The efficiency of ...

Longitudinal In Vivo Imaging of Retinal Cells with Tailored Adeno-Associated Virus Serotypes via Confocal Scanning Laser Ophthalmoscopy

The dynamic nature of retinal cellular processes necessitates advancements in gene delivery and live monitoring techniques to enhance the understanding and treatment of ocular diseases. This study introduces an optimized adeno-associated virus (AAV) approach, utilizing specific serotypes and promoters to achieve optimal transfection efficiency in targeted retinal cells, including retinal ganglion cells (RGCs) and M&#252;ller glia. Leveraging the precision of confocal scanning laser ophthalmoscopy (CSLO), this work presents a non-invasive method for in vivo imaging that captures the longitudinal expression of AAV-mediated green fluorescent protein (GFP). This approach eliminates the need for terminal procedures, preserving the continuity of observation and the well-being of the subject. Furthermore, the GFP signal can be traced in AAV-infected RGCs along the visual pathway to the superior colliculus (SC) and lateral geniculate nucleus (LGN), enabling the potential for direct visual pathway mapping. These findings provide a detailed protocol and demonstrate the application of this powerful tool for real-time studies of retinal cell behavior, disease pathogenesis, and the efficacy of gene therapy interventions, offering valuable insights into the living retina and its connections.

This protocol describes the use of adeno-associated virus (AAV) vectors for cell-specific labeling and in vivo imaging using a confocal scanning laser ophthalmoscope (CSLO). This method enables the investigation of different retinal cell types and their contributions to retinal function and disease.

The dynamic nature of retinal cellular processes necessitates advancements in gene delivery and live monitoring techniques to enhance the understanding ...

AAV-Based Therapies via Intravenous Injection in Preclinical Mouse Models

Consistent Delivery of Adeno-Associated Virus via Lateral Tail-Vein Injection in Adult Mice

Many disorders affect multiple organs or involve different regions of the body, so it is critical to deliver therapeutics systemically to target the affected cells located in different sites. Intravenous injection is a widely used systemic delivery route in preclinical studies that assess treatments intended for body-wide administration. In adult mice, it involves the intravenous administration of the therapeutic agent into the mouse&#39;s lateral tail veins. When mastered, tail-vein injections are safe and fast, and only require simple and commonly available tools. However, tail-vein injections are technically challenging and require extensive training and continuous practice to ensure the accurate delivery of the intended dose.
Here we describe a detailed, optimized, lateral tail-vein injection protocol that we have developed based on our experience and on recommendations that had been previously reported by other groups. Other than the mouse restrainers and insulin syringes, this protocol requires only reagents and equipment that are readily available in most labs. We found that following this protocol results in consistently successful intravenous delivery of adeno-associated virus (AAV) into the tail veins of unsedated 7-9 week-old mice. Additionally, we describe the optimized protocols for the histological detection of fluorescent reporter proteins and vector genome per diploid genome (vg/dg) quantification used to assess AAV transduction and biodistribution. The goal of this protocol is to aid experimenters in easily performing tail-vein injections successfully and consistently, which can reduce the practice time needed to master the technique.

Author Spotlight: Developing Gene Therapies for Muscular Dystrophies and Neuromuscular Disorders

Here we detail an optimized protocol for mouse lateral tail-vein injection to systemically administer adeno-associated virus (AAV) in adult mice. Additionally, we describe protocols of commonly used assays to assess AAV transduction.

Many disorders affect multiple organs or involve different regions of the body, so it is critical to deliver therapeutics systemically to target the ...

Transconjunctival Approach for Injection into the Rat Optic Nerve

The rat serves as an important model for studying disorders of the optic nerve, including heritable, traumatic, neoplastic, and autoimmune conditions. Accessing the rat optic nerve for experimental manipulations and injections is challenging due to the small size of the orbit and the vascularity of the surrounding orbital tissues. Previous techniques have involved a cutaneous incision, which carries a higher risk of infection and requires wound closure. This study aims to describe a unique, less invasive, and potentially more efficient approach to accessing the rat optic nerve. A transconjunctival incision, along with a rat spinal cord hook, is used to isolate the optic nerve for experimental injections. In these experiments, an adenoviral vector is injected into the optic nerve with the goal of using this technique for direct gene therapy delivery in the future. This technique can also be applied by investigators requiring access to the rat optic nerve for other experimental studies, including drug delivery and optic nerve injury research.

This protocol describes a unique technique using a transconjunctival approach to access the rat optic nerve for injections.

The rat serves as an important model for studying disorders of the optic nerve, including heritable, traumatic, neoplastic, and autoimmune conditions. ...