This study was supported by the SANPORC and ALSA Foundation grant (Martin Marsala); the National Sustainability Programme, project number LO1609 (Czech Ministry of Education, Youth and Sports); and RVO: 67985904 (Stefan Juhas and Jana Juhasova).

These studies were carried out under a protocol approved by the Institutional Animal Care and Use Committee of the University of California, San Diego and were in compliance with the Association for Assessment of Laboratory Animal Care guidelines for animal use. All studies were performed in such a manner as to minimize group size and animal suffering.
1. General Animal and Surgical Preparation
<ol>
	<li>Before starting the surgical procedure, thaw the virus (AAV9-UBI-GFP; 5 &#181;L aliquots)8. Prepare a 5% dextran (10,000 MW) solution by mixing dextran powder in distilled water. Mix the virus solution with 5% dextran solution 1:1 to a final dextran concentration of 2.5%.
	<ol>
		<li>Store the virus solution on ice (4 &#176;C).</li>
	</ol>
	</li>
	<li>Use adult C57BL/6J mice (male and female, 20-30 g). Anesthetize the mice using 5% isoflurane (in O2, 1 L/min) and maintain them at 2-3% inhaled isoflurane (in O2, 1 L/min) by nose cone during surgery, depending on the breathing rate and paw pinch response.</li>
	<li>Shave the back of the animals with shaving clippers and clean the skin with 2% chlorhexidine.</li>
	<li>In chronic recovery studies follow a strict sterile technique.</li>
	<li>If lumbar subpial injections are to be performed, cut the skin overlaying the Th8-L1 vertebrae with a scalpel and detach the paravertebral muscle from Th10-12 spinal vertebrae using scissors.
	<ol>
		<li>Mount the animal into a standard stereotaxic frame using mouse spinal clamps.</li>
		<li>Shave the both sides of the lamina of the Th10-12 vertebrae using a dental drill (drill bit: 0.9 mm, speed: 20,000 rpm) until cracks appear.</li>
		<li>Remove cracked bone fragments with forceps and expose the dorsal surface of the lumbar spinal cord.</li>
		<li>Cut open the dura about 1 cm using a 30 G stainless steel needle and forceps.</li>
	</ol>
	</li>
	<li>If cervical subpial injections are to be performed, incise the dorsal neck skin 1.5-2 cm using scissors and expose the C1-C2 segments.
	<ol>
		<li>Remove the atlanto-occipital membrane of the cisterna magna using a 23G stainless steel needle and forceps.</li>
		<li>Clean the incision site of any tissue and bone debris using cotton swabs.</li>
		<li>Cut open the dura about 2-3 mm using a 30 G stainless steel needle and forceps.</li>
	</ol>
	</li>
</ol>
2. Opening the Pial Membrane and Inserting the Subpial Needle for AAV9 Delivery
<ol>
	<li>Mount the 34 G pia-penetrating needle into the Z-arm of an XYZ manipulator using a glass capillary holder (Figure 1A and B). 
	NOTE: To manufacture the pia-penetrating needle, the original beveled tip of the 34G needle is sharpened using a glass capillary beveller with diamond abrasive plate - coarse (5.0 &#181;m to 50 &#181;m tip sizes) and grinding angle of 15 - 20&#176;. The tip of the needle (1 mm length, measured from the tip) is then gently bent to about 90&#176; (Figure 1B, left insert).</li>
	<li>Using a surgical dissecting scope, set to 8-10 X magnification, penetrate the pia with the pia-penetrating needle by about 1 mm (Figure 1C) using the X-arm.
	<ol>
		<li>Keep the angle of the penetrating needle to the tissue surface at 5-10&#176;.</li>
	</ol>
	</li>
	<li>After the pia opening, remove the pia-penetrating needle horizontally from the subpial space (Figure 1 D) using the X-arm. 
	NOTE: Remember the penetrated site by a landmark, such as a blood vessel.</li>
	<li>Load a blunt 36G injection needle with AAV9-UBI-GFP virus using a 50-&#181;L microsyringe connected to the injection needle with PE-10 or PE-20 tubing.</li>
	<li>Mount the needle into the Z-arm of a second XYZ manipulator (Figure 1A and B) using a glass capillary holder (Figure 1B, right insert). 
	NOTE: To manufacture the subpial AAV9 injection needle, the blunt tip of a 36 G needle is polished using a glass capillary beveller with diamond abrasive plate - coarse (5.0 &#181;m to 50 &#181;m tip sizes) to remove the sharp edges. The tip of the needle (2-3 mm length, measured from the tip) is then gently bent to about 90&#176;. The pia-penetrating and subpial injection needles are inserted into 1-2 cm-long 20G slave stainless steel tubing (10 mm from the end of the needle) and glued with epoxy. The use of 20 G (0.91 mm-diameter) tubing is required for a secure attachment to the glass capillary holder.</li>
	<li>By manipulating the X, Y, and Z arms of the second manipulator, position the tip of the AAV9 injection needle into the pia-penetrated site and then advance it about 2-3 mm into the subpial space through the previous pial membrane opening using the X-arm ( Figure 1E and F). 
	NOTE: 1) The AAV9-UBI-GFP is prepared according to previously reported protocols9,10, and the final titers are adjusted to 1.2 x 1013 genome copies per mL (gc/mL). 2) There is no need to mark the site of the pial opening because it is readily identifiable (Figure 1D).</li>
	<li>Inject the AAV9-UBI-GFP (1.5, 3, or 5 &#181;L) into the subpial space using a 50 &#181;L microsyringe (see Table 1 for experimental groups).</li>
	<li>Remove the injection needle from the subpial space after the AAV9-UBI-GFP injection is complete.</li>
	<li>Close the muscle and skin using 4.0 monofilament suture and surgical clips. 
	NOTE: There is no need to seal the open vertebra.</li>
	<li>Allow the animals to recover on a heating pad.</li>
	<li>For pain control inject Buprenorphine 0.05 mg/kg/sc &#160;every 12 h for 2-3 days post-surgery.</li>
</ol>
3. Perfusion-fixation, Tissue Cryoprotection, and Immunofluorescence Staining
<ol>
	<li>At a predetermined time point after the subpial AAV9 injections, deeply anesthetize the mice with euthanasia solution (see the Table of Materials, 0.3 mL) and transcardially perfuse them with 20 mL of heparinized saline followed by 20 mL of 4% paraformaldehyde in PBS.</li>
	<li>Dissect the spinal cords and brains using a bone rongeur and post-fix them in 4% formaldehyde in PBS overnight at 4 &#176;C.</li>
	<li>Cryoprotect the spinal cords and brains with 30% sucrose in PBS for a minimum of 5-7 days.</li>
	<li>Cut coronal, transverse, or longitudinal/horizontal frozen sections (30 &#181;m-thick) on a cryostat and store them in PBS at 4 &#176;C.</li>
</ol>
4. Immunofluorescence Staining of Spinal Cord and Brain Sections (See the Table of Materials)
<ol>
	<li>Incubate free-floating sections in primary antibodies overnight.</li>
	<li>After incubation with primary antibodies, wash the sections three times in PBS and incubate with fluorescence-conjugated donkey anti-rabbit, donkey anti-chicken, and donkey anti-goat secondary antibodies.</li>
	<li>Mount the sections on microscopy slides, dry them at room temperature, and cover them with an anti-fade medium.</li>
	<li>Capture images using epifluorescence fluorescence microscope (objectives: 10X, NA-0.3; 20X, NA-0.8; and 63X, NA-1.4).</li>
</ol>

<ol>
	<li>Foust, K. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intravascular+AAV9+preferentially+targets+neonatal+neurons+and+adult+astrocytes.">Intravascular AAV9 preferentially targets neonatal neurons and adult astrocytes.</a> Nat Biotechnol. 27 (1), 59-65 (2009).</li><li>Duque, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intravenous+administration+of+self-complementary+AAV9+enables+transgene+delivery+to+adult+motor+neurons.">Intravenous administration of self-complementary AAV9 enables transgene delivery to adult motor neurons.</a> Mol Ther. 17 (7), 1187-1196 (2009).</li><li>Gray, S. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preclinical+differences+of+intravascular+AAV9+delivery+to+neurons+and+glia:+a+comparative+study+of+adult+mice+and+nonhuman+primates.">Preclinical differences of intravascular AAV9 delivery to neurons and glia: a comparative study of adult mice and nonhuman primates.</a> Mol Ther. 19 (6), 1058-1069 (2011).</li><li>Meyer, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improving+single+injection+CSF+delivery+of+AAV9-mediated+gene+therapy+for+SMA:+a+dose-response+study+in+mice+and+nonhuman+primates.">Improving single injection CSF delivery of AAV9-mediated gene therapy for SMA: a dose-response study in mice and nonhuman primates.</a> Mol Ther. 23 (3), 477-487 (2015).</li><li>Foust, K. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Therapeutic+AAV9-mediated+suppression+of+mutant+SOD1+slows+disease+progression+and+extends+survival+in+models+of+inherited+ALS.">Therapeutic AAV9-mediated suppression of mutant SOD1 slows disease progression and extends survival in models of inherited ALS.</a> Mol Ther. 21 (12), 2148-2159 (2013).</li><li>Passini, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translational+fidelity+of+intrathecal+delivery+of+self-complementary+AAV9-survival+motor+neuron+1+for+spinal+muscular+atrophy.">Translational fidelity of intrathecal delivery of self-complementary AAV9-survival motor neuron 1 for spinal muscular atrophy.</a> Hum Gene Ther. 25 (7), 619-630 (2014).</li><li>Bell, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+neuron+transduction+after+intracisternal+delivery+of+AAV9+in+a+cynomolgus+macaque.">Motor neuron transduction after intracisternal delivery of AAV9 in a cynomolgus macaque.</a> Hum Gene Ther Methods. 26 (2), 43-44 (2015).</li><li>Miyanohara, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Potent+spinal+parenchymal+AAV9-mediated+gene+delivery+by+subpial+injection+in+adult+rats+and+pigs.">Potent spinal parenchymal AAV9-mediated gene delivery by subpial injection in adult rats and pigs.</a> Mol Ther Methods Clin Dev. 3, 16046 (2016).</li><li>Xu, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+gene+knockdown+in+rat+dorsal+root+ganglia+mediated+by+self-complementary+adeno-associated+virus+serotype+5+following+intrathecal+delivery.">In vivo gene knockdown in rat dorsal root ganglia mediated by self-complementary adeno-associated virus serotype 5 following intrathecal delivery.</a> PLoS One. 7 (3), 32581 (2012).</li><li>Xiao, X., Li, J., Samulski, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production+of+high-titer+recombinant+adeno-associated+virus+vectors+in+the+absence+of+helper+adenovirus.">Production of high-titer recombinant adeno-associated virus vectors in the absence of helper adenovirus.</a> J Virol. 72 (3), 2224-2232 (1998).</li></ol>

Martin Marsala is a co-founder of Neurgain Technologies, Inc. (San Diego, USA).

The current study describes a technique of subpial vector (AAV9) delivery in adult mice. As demonstrated in the accompanying video, this approach and technique can effectively be used, provided that the required instruments and pia-penetrating needle and subpial injection needle are properly manufactured, according to the established and tested specifications.
Critical Technical Variables in Performing a Consistent and Safe Subpial Injection in Mice: 
As demonstrated, a s...

The use of AAV vectors to treat a variety of spinal cord and CNS neurodegenerative disorders is becoming a well-accepted platform to effectively upregulate or silence the expression of gene(s) of interest. One of the key limitations to the more effective utilization of this technology to treat CNS/spinal cord disorders is the limited ability to deliver AAV vector(s) to the deep brain or spinal cord parenchyma in adult mammals.
It was demonstrated, for example, that the systemic delivery of AAV9 in adult rodents, cats, or non-human primates is only moderately effective at inducing transgene expression in neurons in the brain and spinal cord1,2,3. The more effective intrathecal delivery of AAV9 vectors has also been shown to lead to only limited transgene expression in anatomically-defined pools of neurons. More specifically, it has been demonstrated that cisternal or lumbo-sacral intrathecal AAV9 delivery in non-human primates, pigs, or rodents leads to a high level of transgene expression in spinal &#945;-motoneurons and segmental dorsal root ganglion neurons. However, minimal or no expression in spinal interneurons or ascending or descending axons in the white matter is seen4,5,6,7. Collectively, these data show that a highly effective biological-anatomical barrier exists, which prevents the diffusion of intrathecally delivered AAV into deeper spinal parenchyma.
In a previous study using adult rats and pigs, a novel subpial vector delivery technique was developed8. Using this approach, highly potent and multi-segmental transgene expression was demonstrated after a single-bolus subpial AAV9 delivery. Intense GFP expression was consistently seen in neurons, glial cells, and descending/ascending axons through the injected spinal segments. This study demonstrated for the first time that the pia mater represents the primary barrier limiting effective AAV9 diffusion into the spinal parenchyma from the intrathecal space. While this previously developed technique and subpial injection device is relatively easy to use in large rodents (like rats) or adult pigs, the system is not suitable for use in small animals, such as adult mice. Because of the high number of available transgenic mouse models of a variety of neurodegenerative disorders, there is a clear need for the development of an effective spinal-parenchymal vector delivery technique in mice. The availability of such a technique would permit the study of the effect of specific gene silencing (e.g., using shRNA) or upregulation using cell-non-specific (e.g., cytomegalovirus-CMV or Ubiquitin) or cell-specific (e.g., synapsin or glial fibrillary acidic protein (GFAP)) promoters during early postnatal development or under diseased conditions.
Accordingly, in the present study, we have developed and validated a miniature subpial vector delivery system that can effectively be used in adult mice. Similarly, as in previous rat and pig studies, this work demonstrates potent transgene expression throughout the spinal parenchyma after a single-bolus subpial AAV9 delivery in mice. The simplicity of this approach, the very good tolerability of injected mice to subpial AAV9 delivery, and the high potency of transgene expression in the spinal parenchyma suggest that this technique can effectively be implemented in any laboratory setting and used in experiments targeting spinal gene expression.

<table><tbody><tr><td>C57BL/6J Mice</td><td>Jackson Labs</td><td>664</td><td></td></tr><tr><td>Lab Standard Stereotaxic for Mice</td><td>Harvard Apparatus</td><td>72-9568</td><td></td></tr><tr><td>Mouse Spinal Adaptor</td><td>Harvard Apparatus</td><td>72-4811</td><td></td></tr><tr><td>XYZ Manipulator</td><td>Stoelting</td><td>51604</td><td></td></tr><tr><td>Manual Infusion Pump</td><td>Stoelting</td><td>51218</td><td></td></tr><tr><td>34G Beveled Nanofill Needle</td><td>World Precision Instruments</td><td>NF34BV-2</td><td></td></tr><tr><td>36G Blunt Nanofill needle</td><td>World Precision Instruments</td><td>NF-36BL-2</td><td></td></tr><tr><td>Fluriso, Isoflurane</td><td>MWI Veterinary Supply</td><td>502017</td><td></td></tr><tr><td>Chlorhexidine Solution</td><td>MWI Veterinary Supply</td><td>501027</td><td></td></tr><tr><td>20G Stainless Steel Needle</td><td>Becton-Dickinson</td><td>305175</td><td></td></tr><tr><td>23G Stainless Steel Needle</td><td>Becton-Dickinson</td><td>305145</td><td></td></tr><tr><td>30G Stainless Steel Needle</td><td>Becton-Dickinson</td><td>305128</td><td></td></tr><tr><td>Cotton Tipped Applicator</td><td>MWI Veterinary Supply</td><td>27426</td><td></td></tr><tr><td>Glass Capillary Beveller&amp;nbsp;</td><td>Narishige International</td><td>SM-25B</td><td></td></tr><tr><td>Slide Microscope Superfrost</td><td>Leica Microsystems</td><td>M80</td><td></td></tr><tr><td>50 &amp;mu;L Microsyringe&amp;nbsp;</td><td>Hamilton</td><td>81242</td><td></td></tr><tr><td>BD Intramedic PE-20 Tubing</td><td>Becton, Dickinson</td><td>427406</td><td></td></tr><tr><td>BD Intramedic PE-10 Tubing</td><td>Becton, Dickinson</td><td>427401</td><td></td></tr><tr><td>4-0 monofilament suture</td><td>VetOne</td><td>V1D397</td><td></td></tr><tr><td>Glass Capillary Beveller&amp;nbsp;</td><td>Narishige</td><td>Pipet Micro Grinder EG-40&amp;nbsp;</td><td></td></tr><tr><td>5 min Epoxy (Epoxy Clear)</td><td>Devcon</td><td>14310</td><td></td></tr><tr><td>Euthanasia Solution</td><td>MWI Veterinary Supply</td><td>11168</td><td></td></tr><tr><td>Heparin Inj 1,000 U/mL</td><td>MWI Veterinary Supply</td><td>54254</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>158127</td><td></td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S0389</td><td></td></tr><tr><td>Anti NeuN Antibody</td><td>EMD-Millipore</td><td>ABN78</td><td>Primary Rabbit Polyclonal Antibody, 1:1000</td></tr><tr><td>Anti-Choline Acetyltransferase (CHAT) Antibody</td><td>EMD-Millipore</td><td>AB144P</td><td>Primary Goat Polyclonal Antibody, 1:100</td></tr><tr><td>Anti GFP Antibody</td><td>Aves Labs</td><td>GFP-1020</td><td>Primary Chicken Polyclonal Antibody, 1:1000</td></tr><tr><td>Donkey anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 594</td><td>ThermoFisher Scientific</td><td>A21207</td><td>Secondary Antibody, 1:1000</td></tr><tr><td>&amp;nbsp;Donkey anti-Rabbit IgG Secondary Antibody, Alexa Fluor 680</td><td>ThermoFisher Scientific</td><td>A10043</td><td>Secondary Antibody, 1:1000</td></tr><tr><td>Donkey anti-Chicken IgY Secondary Antibody, Alexa Fluor 488</td><td>Jackson Immunoresearch Labs</td><td>703-545-155</td><td>Secondary Antibody, 1:1000</td></tr><tr><td>Donkey Anti-Goat IgG H&amp;amp;L (Alexa Fluor 647</td><td>Abcam</td><td>ab150131</td><td>Secondary Antibody, 1:1000</td></tr><tr><td>Slide Microscope Superfrost</td><td>Fisher Scientific</td><td>12-550-143</td><td></td></tr><tr><td>ProLong Gold Antifade Mountant</td><td>Fisher Scientific</td><td>P36930</td><td></td></tr><tr><td>Epifluorescence Microscope</td><td>Zeiss</td><td>Zeiss AxioImager M2</td><td></td></tr><tr><td>Fluorescence Confocal Microscope</td><td>Olympus</td><td>Olympus FV1000</td><td></td></tr><tr><td>Dextran</td><td>Polysciences, Inc</td><td>19411</td><td></td></tr><tr><td>AAV9-UBC-GFP</td><td>UCSD Viral Vector Core Laboratory</td><td></td><td></td></tr></tbody></table>

subpial adeno-associated virus 9 (aav9) vector delivery in adult mice

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. Using subpially-placed polyethylene catheters (PE-10 or PE-5) for AAV9 delivery, potent transgene expression through the spinal parenchyma (white and gray matter) in subpially-injected spinal segments has been demonstrated. Because of the wide range of transgenic mouse models of neurodegenerative diseases, there is a strong desire for the development of a potent central nervous system (CNS)-targeted vector delivery technique in adult mice. Accordingly, the present study describes the development of a spinal subpial vector delivery device and technique to permit safe and effective spinal AAV9 delivery in adult C57BL/6J mice. In spinally immobilized and anesthetized mice, the pia mater (cervical 1 and lumbar 1-2 spinal segmental level) was incised with a sharp 34 G needle using an XYZ manipulator. A second XYZ manipulator was then used to advance a blunt 36G needle into the lumbar and/or cervical subpial space. The AAV9 vector (3-5 &#181;L; 1.2 x 1013 genome copies (gc)) encoding green fluorescent protein (GFP) was then injected subpially. After injections, neurological function (motor and sensory) was assessed periodically, and animals were perfusion-fixed 14 days after AAV9 delivery with 4% paraformaldehyde. Analysis of horizontal or transverse spinal cord sections showed transgene expression throughout the entire spinal cord, in both gray and white matter. In addition, intense retrogradely-mediated GFP expression was seen in the descending motor axons and neurons in the motor cortex, nucleus ruber, and formatio reticularis. No neurological dysfunction was noted in any animals. These data show that the subpial vector delivery technique can successfully be used in adult mice, without causing procedure-related spinal cord injury, and is associated with highly potent transgene expression throughout the spinal neuraxis.

Potent Transgene Expression in Subpially AAV9-injected Segments: 
The analysis of transgene (GFP) expression in spinal cord sections at 14 days after AAV9 delivery showed AAV9-dose dependent GFP expression throughout the spinal parenchyma. First, two bilateral 3 &#181;L injections of AAV9-UBI-GFP injected into the upper lumbar subpial space were associated with the near-complete infection of the white and gray matter in the whole lumbar spinal cord, extending to the ...

Watch this Scientific Journal Video about Subpial Adeno-associated Virus 9 AAV9 Vector Delivery in Adult Mice at JoVE.com

Subpial Adeno-associated Virus 9 AAV9 Vector Delivery in Adult Mice

The goal of the present study was to develop and validate the potency and safety of spinal adeno-associated virus 9 (AAV9)-mediated gene delivery by using a novel subpial gene delivery technique in adult mice.

Subpial Adeno-associated Virus 9 (AAV9) Vector Delivery in Adult Mice

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. ...

subpial-adeno-associated-virus-9-aav9-vector-delivery-in-adult-mice

Research

JoVE Journal

Neuroscience

17.4K Views.  University of California, San Diego.  Het doel van de huidige studie was het ontwikkelen en valideren van de kracht en de veiligheid van de lever-adeno-geassocieerde virus 9 (AAV9) -gemedieerde genafgifte door gebruik te maken van een nieuwe subpiale genafgifte techniek bij volwassen muizen. 

Video: Subpial Adeno-associated Virus 9 AAV9 Vector Delivery in Adult Mice

Preparation of rAAV9 to Overexpress or Knockdown Genes in Mouse Hearts

Controlling the expression or activity of specific genes through the myocardial delivery of genetic materials in murine models permits the investigation of gene functions. Their therapeutic potential in the heart can also be determined. There are limited approaches for in vivo molecular intervention in the mouse heart. Recombinant adeno-associated virus (rAAV)-based genome engineering has been utilized as an essential tool for in vivo cardiac gene manipulation. The specific advantages of this technology include high efficiency, high specificity, low genomic integration rate, minimal immunogenicity, and minimal pathogenicity. Here, a detailed procedure to construct, package, and purify the rAAV9 vectors is described. Subcutaneous injection of rAAV9 into neonatal pups results in robust expression or efficient knockdown of the gene(s) of interest in the mouse heart, but not in the liver and other tissues. Using the cardiac-specific TnnT2 promoter, high expression of GFP gene in the heart was obtained. Additionally, target mRNA was inhibited in the heart when a rAAV9-U6-shRNA was utilized. Working knowledge of rAAV9 technology may be useful for cardiovascular investigations.

In this manuscript, a method to prepare recombinant adeno-associated virus 9 (rAAV9) vectors to manipulate gene expression in the mouse heart is described.

Controlling the expression or activity of specific genes through the myocardial delivery of genetic materials in murine models permits the investigation ...

Genetics

Systemic Delivery of MicroRNA Using Recombinant Adeno-associated Virus Serotype 9 to Treat Neuromuscular Diseases in Rodents

RNA interference via the endogenous miRNA pathway regulates gene expression by controlling protein synthesis through post-transcriptional gene silencing. In recent years, miRNA-mediated gene regulation has shown potential for treatment of neurological disorders caused by a toxic gain of function mechanism. However, efficient delivery to target tissues has limited its application. Here we used a transgenic mouse model for spinal and bulbar muscular atrophy (SBMA), a neuromuscular disease caused by polyglutamine expansion in the androgen receptor (AR), to test gene silencing by a newly identified AR-targeting miRNA, miR-298. We overexpressed miR-298 using a recombinant adeno-associated virus (rAAV) serotype 9 vector to facilitate transduction of non-dividing cells. A single tail-vein injection in SBMA mice induced sustained and widespread overexpression of miR-298 in skeletal muscle and motor neurons and resulted in amelioration of the neuromuscular phenotype in the mice.

Here we describe the delivery of microRNA using a recombinant adeno-associated virus serotype 9 in a mouse model of a neuromuscular disease. A single peripheral administration in mice resulted in sustained miRNA overexpression in muscle and motor neurons, providing an opportunity to study miRNA function and therapeutic potential in vivo.

RNA interference via the endogenous miRNA pathway regulates gene expression by controlling protein synthesis through post-transcriptional gene silencing. ...

Effective Systemic Delivery in Mice Using Retrobulbar Injection Technique

Systemic Treatment for Postnatal, Juvenile, and Runted Adult Mice by Retrobulbar Sinus Injection

While tail vein injections are frequently used as a systemic route of delivery in adult mice, retrobulbar injections are an alternative method for systemic delivery with fewer limitations. First, tail vein injections (TVIs) are limited to adult mice where the size of the tail vein is suitable for access. Being restricted to treating adult mice can be problematic when dealing with mouse models that do not survive to adulthood. Second, TVIs are not feasible for mouse models with growth retardation phenotypes where the mice never achieve the size of adult wildtype mice. Therefore, retrobulbar injections can be successfully used to treat both young and small adult mice. Lastly, retrobulbar injections are performed under anesthesia, which is less stressful on the mice than TVIs that are commonly performed without anesthesia. This article presents a protocol and detailed instructions for retrobulbar injections that can be used for systemic delivery to small and young mice.

This article provides a protocol and an accompanying video for the retrobulbar sinus injection of up to a total volume of 150 &#181;L for postnatal, juvenile, and runted adult mice. This procedure is particularly well suited for the injection of small mice (15 g) when tail vein injection is not feasible.

While tail vein injections are frequently used as a systemic route of delivery in adult mice, retrobulbar injections are an alternative method for ...

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, ...

Production, Purification, and Quality Control for Adeno-associated Virus-based Vectors

Gene delivery tools based on adeno-associated viruses (AAVs) are a popular choice for the delivery of transgenes to the central nervous system (CNS), including gene therapy applications. AAV vectors are non-replicating, able to infect both dividing and non-dividing cells&#160;and provide long-term transgene expression. Importantly, some serotypes, such as the newly described PHP.B, can cross the blood-brain-barrier (BBB) in animal models, following systemic delivery. AAV vectors can be efficiently produced in the laboratory. However, robust and reproducible protocols are required to obtain AAV vectors with sufficient purity levels and titer values high enough for in vivo applications. This protocol describes an efficient and reproducible strategy for AAV vector production, based on an iodixanol gradient purification strategy. The iodixanol purification method is suitable for obtaining batches of high-titer AAV vectors of high purity, when compared to other purification methods. Furthermore, the protocol is generally faster than other methods currently described. In addition, a quantitative polymerase chain reaction (qPCR)-based strategy is described for a fast and accurate determination of the vector titer, as well as a silver staining method to determine the purity of the vector batch. Finally, representative results of gene delivery to the CNS, following systemic administration of AAV-PHP.B, are presented. Such results should be possible in all labs using the protocols described in this article.

Here, we describe an efficient and reproducible strategy to produce, titer, and quality-control batches of adeno-associated virus vectors. It allows the user to obtain a vector preparation with high-titer&#160;(&#8805;1 x 1013 vector genomes/mL) and a high purity, ready for in vitro or in vivo use.

Gene delivery tools based on adeno-associated viruses (AAVs) are a popular choice for the delivery of transgenes to the central nervous system (CNS), ...

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.

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 ...

Medicine

Lumbar Puncture Delivery of AAV9 in Juvenile Rats for CNS Gene Therapy

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a toxic gene, or provide a gene whose product can modulate the biology of the disease. The delivery method for the therapeutic vector can take many forms, ranging from intravenous infusion for systemic delivery to direct injection into the target tissue. For neurodegenerative disorders, it is often desirable to skew transduction towards the brain and/or spinal cord. The least invasive approach to target the entire central nervous system involves injection into the cerebrospinal fluid (CSF), allowing the therapeutic to reach a large fraction of the central nervous system. The safest approach to deliver a vector into the CSF is the lumbar intrathecal injection, where a needle is introduced into the lumbar cistern of the spinal cord. This technique, also known as a lumbar puncture, has been widely used in neonatal and adult rodents and in large animal models. While the technique is similar across species and developmental stages, subtle differences in size, structure, and elasticity of tissues surrounding the intrathecal space require accommodations in the approach. This article describes a method for performing lumbar puncture in juvenile rats to deliver an adeno-associated serotype 9 vector. Here, 25-35 &#181;L of vector were injected into the lumbar cistern, and a green fluorescent protein (GFP) reporter was used to evaluate the transduction profile resulting from each injection. The benefits and challenges of this approach are discussed.

Author Spotlight: Assessing Intrathecal Gene Therapy Efficacy in Juvenile Rats

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.

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a ...

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 ...

Subpial Adeno-associated Virus 9 (AAV9) Vector Delivery in Adult Mice

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