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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>&#160; University of North Carolina at Chapel Hill</td>
			<td>&#160; /</td>
			<td></td>
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		<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>&#160;Forceps</td>
			<td>Fine Science Tools</td>
			<td>F6521</td>
			<td></td>
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			<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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2.7K 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

High-Efficiency Transduction of Liver Cancer Cells by Recombinant Adeno-Associated Virus Serotype 3 Vectors

Recombinant vectors based on a non-pathogenic human parvovirus, the adeno-associated virus 2 (AAV2) have been developed, and are currently in use in a number of gene therapy clinical trials. More recently, a number of additional AAV serotypes have also been isolated, which have been shown to exhibit selective tissue-tropism in various small and large animal models1. Of the 10 most commonly used AAV serotypes, AAV3 is by far the least efficient in transducing cells and tissues in vitro as well as in vivo.
However, in our recently published studies, we have documented that AAV3 vectors transduce human liver cancer - hepatoblastoma (HB) and hepatocellular carcinoma (HCC) - cell lines extremely efficiently because AAV3 utilizes human hepatocyte growth factor receptor as a cellular co-receptor for binding and entry in these cells2,3.
In this article, we describe the steps required to achieve high-efficiency transduction of human liver cancer cells by recombinant AAV3 vectors carrying a reporter gene. The use of recombinant AAV3 vectors carrying a therapeutic gene may eventually lead to the potential gene therapy of liver cancers in humans.

In this article, we describe the identification of the adeno-associated virus serotype 3 (AAV3) as the most efficient vector for targeting human liver cancer cells.

Recombinant vectors based on a non-pathogenic human parvovirus, the adeno-associated virus 2 (AAV2) have been developed, and are currently in use in a ...

Protocol for Long Duration Whole Body Hyperthermia in Mice

Hyperthermia is a general term used to define the increase in core body temperature above normal. It is often used to describe the increased core body temperature that is observed during fever. The use of hyperthermia as an adjuvant has emerged as a promising procedure for tumor regression in the field of cancer biology. For this purpose, the most important requirement is to have reliable and uniform heating protocols. We have developed a protocol for hyperthermia (whole body) in mice. In this protocol, animals are exposed to cycles of hyperthermia for 90 min followed by a rest period of 15 min. During this period mice have easy access to food and water. High body temperature spikes in the mice during first few hyperthermia exposure cycles are prevented by immobilizing the animal. Additionally, normal saline is administered in first few cycles to minimize the effects of dehydration. This protocol can simulate fever like conditions in mice up to 12-24 hr. We have used 8-12 weeks old BALB/Cj female mice to demonstrate the protocol.

This paper describes a protocol for whole body hyperthermia in mice that can stimulate fever like conditions up to 12-24 hr.

Hyperthermia is a general term used to define the increase in core body temperature above normal. It is often used to describe the increased core body ...

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 ...

High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal systems. HFUS has the ability to detect structures as small as 30 &#181;m, a property that has been utilized for visualizing superficial lymph nodes in rodents in brightness (B)-mode. Combining power Doppler with B-mode imaging allows for measuring circulatory blood flow within lymph nodes and other organs. While HFUS has been utilized for lymph node imaging in a number of mouse&#160; model systems, a detailed protocol describing HFUS imaging and characterization of the cervical lymph nodes in mice has not been reported. Here, we show that HFUS can be adapted to detect and characterize cervical lymph nodes in mice. Combined B-mode and power Doppler imaging can be used to detect increases in blood flow in immunologically-enlarged cervical nodes. We also describe the use of B-mode imaging to conduct fine needle biopsies of cervical lymph nodes to retrieve lymph tissue for histological&#160; analysis. Finally, software-aided steps are described to calculate changes in lymph node volume and to visualize changes in lymph node morphology following image reconstruction. The ability to visually monitor changes in cervical lymph node biology over time provides a simple and powerful technique for the non-invasive monitoring of cervical lymph node alterations in preclinical mouse models of oral cavity disease.

This protocol describes the application of high-frequency ultrasound (HFUS) for imaging mouse cervical lymph nodes. This technique optimizes visualization and quantification of cervical lymph node morphology, volume and blood flow. Image-guided biopsy of cervical lymph nodes and processing of lymph tissue for histological evaluation is also demonstrated.

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal ...

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

Adeno-Associated Virus-Mediated Delivery of CRISPR for Cardiac Gene Editing in Mice

The clustered, regularly interspaced, short, palindromic repeat (CRISPR) system has greatly facilitated genome engineering in both cultured cells and living organisms from a wide variety of species. The CRISPR technology has also been explored as novel therapeutics for a number of human diseases. Proof-of-concept data are highly encouraging as exemplified by recent studies that demonstrate the feasibility and efficacy of gene editing-based therapeutic approach for Duchenne muscular dystrophy (DMD) using a murine model. In particular, intravenous and intraperitoneal injection of the recombinant adeno-associated virus (rAAV) serotype rh.74 (rAAVrh.74) has enabled efficient cardiac delivery of the Staphylococcus aureus CRISPR-associated protein 9 (SaCas9) and two guide RNAs (gRNA) to delete a genomic region with a mutant codon in exon 23 of mouse Dmd gene. This same approach can also be used to knock out the gene-of-interest and study their cardiac function in postnatal mice when the gRNA is designed to target the coding region of the gene. In this protocol, we show in detail how to engineer rAAVrh.74-CRISPR vector and how to achieve highly efficient cardiac delivery in neonatal mice.

Here we provide a detailed protocol to carry out in vivo cardiac gene editing in mice using recombinant Adeno-Associated Virus(rAAV)-mediated delivery of CRISPR. This protocol offers a promising therapeutic strategy to treat dystrophic cardiomyopathy in Duchenne muscular dystrophy and can be used to generate cardiac-specific knockout in postnatal mice.

The clustered, regularly interspaced, short, palindromic repeat (CRISPR) system has greatly facilitated genome engineering in both cultured cells and ...

Continuous Blood Sampling in Small Animal Positron Emission Tomography/Computed Tomography Enables the Measurement of the Arterial Input Function

For quantitative analysis and bio-kinetic modeling of positron emission tomography/computed tomography (PET/CT) data, the determination of the temporal blood time-activity concentration also known as arterial input function (AIF) is a key point, especially for the characterization of animal disease models and the introduction of newly developed radiotracers. The knowledge of radiotracer availability in the blood helps to interpret PET/CT-derived data of tissue activity. For this purpose, online blood sampling during the PET/CT imaging is advisable to measure the AIF. In contrast to manual blood sampling and image-derived approaches, continuous online blood sampling has several advantages. Besides the minimized blood loss, there is an improved resolution and a superior accuracy for the blood activity measurement. However, the major drawback of online blood sampling is the costly and time-consuming preparation to catheterize the femoral vessels of the animal. Here, we describe an easy and complete workflow for catheterization and continuous blood sampling during small animal PET/CT imaging and compared it to manual blood sampling and an image-derived approach. Using this highly-standardized workflow, the determination of the fluorodeoxyglucose ([18F]FDG) AIF is demonstrated. Further, this procedure can be applied to any radiotracer in combination with different animal models to create fundamental knowledge of tracer kinetic and model characteristics. This allows a more precise evaluation of the behavior of pharmaceuticals, both for diagnostic and therapeutic approaches in the preclinical research of oncological, neurodegenerative and myocardial diseases.

Here a protocol for continuous blood sampling during PET/CT imaging of rats to measure the arterial input function (AIF) is described. The catheterization, the calibration and setup of the system and the data analysis of the blood radioactivity are demonstrated. The generated data provide input parameters for subsequent bio-kinetic modeling.

For quantitative analysis and bio-kinetic modeling of positron emission tomography/computed tomography (PET/CT) data, the determination of the temporal ...

Self-Administration of Drugs in Mouse Models of Feeding and Obesity

Preclinical studies in mice often rely on invasive protocols, such as injections or oral gavage, to deliver drugs. These stressful routes of administration have significant effects on important metabolic parameters including food intake and body weight. Although an attractive option to circumvent this is to compound the drug in rodent food or dissolve it in water, these approaches also have limitations as they are affected by drug stability at room temperature for extended periods of time, the drug's solubility in water, and that the dosing is highly dependent on timing of food or water intake. The constant availability of the drug also limits translational relevance on how drugs are administered to patients. To overcome these limitations, drugs can be mixed with highly palatable food, such as peanut butter, allowing mice to self-administer compounds. Mice reliably and reproducibly consume the drug/peanut butter pellet in a short time frame. This approach facilitates a delivery approach with minimal stress compared with an injection or gavage. This protocol demonstrates the approach of drug preparation, animal acclimatization to placebo delivery, and drug delivery. The implications of this approach are discussed in studies related to timing of drug administration and the circadian rhythm.

The overall goal of this procedure is to describe a method for self-administration of drugs that can be used in mouse models of feeding and obesity.

Preclinical studies in mice often rely on invasive protocols, such as injections or oral gavage, to deliver drugs. These stressful routes of ...

AAV Isolation by Double Iodixanol Density Gradient Centrifugation

Suspension Culture Production and Purification of Adeno-Associated Virus by Iodixanol Density Gradient Centrifugation for In Vivo Applications

This protocol describes recombinant adeno-associated virus (rAAV) production and purification by iodixanol density gradient centrifugation, a serotype-agnostic method of purifying AAV first described in 1999. rAAV vectors are widely used in gene therapy applications to deliver transgenes to various human cell types. In this work, the recombinant virus is produced by transfection of Expi293 cells in suspension culture with plasmids encoding the transgene, vector capsid, and adenoviral helper genes. Iodixanol density gradient centrifugation purifies full AAV particles based on particle density. Additionally, three steps are included in this now-ubiquitous methodology in order to increase total virus yield, decrease the risk of precipitation due to contaminating proteins, and further concentrate the final virus product, respectively: precipitation of viral particles from cell media using a solution of polyethylene glycol (PEG) and sodium chloride, the introduction of a second round of iodixanol density gradient centrifugation, and buffer exchange via a centrifugal filter. Using this method, it is possible to consistently achieve titers in the range of 1012 viral particles/mL of exceptional purity for in vivo use.

Author Spotlight: Efficient Adeno-Associated Virus Isolation for Pre-Clinical Applications

Adeno-associated virus is produced in suspension cell culture and purified by double iodixanol density gradient centrifugation. Steps are included to increase total virus yield, decrease the risk of virus precipitation, and further concentrate the final virus product. Expected final titers reach 1012 viral particles/mL and are suitable for pre-clinical in vivo use.