Este trabajo fue posible gracias a subvenciones del NIH (EY012977), el Premio de carrera en las Ciencias Biomédicas del Fondo Burroughs Wellcome, la Fundación de Whitehall, y la Fundación Sloan (AKM).

1. Virus Manipulación y almacenamiento <ol><li> Protección adecuada y técnicas de manejo deben ser elegidos con base en el nivel de bioseguridad de los virus que se utilizarán. Estas prácticas se pueden encontrar en Seguridad de la Biotecnología en la edición de los laboratorios microbiológicos y biomédicos º 5, disponible en el sitio web de CDC ( <a href="http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm">http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm</a> ). El uso de vectores AAV está aprobado para un nivel de bioseguridad (BSL-1). Para el experimento se muestra aquí, una bata de laboratorio y guantes durante el manejo del virus, de acuerdo con los procedimientos para la manipulación de agentes BSL-1. </li><li> A fin de preservar la actividad de los virus es mejor dividirlo en pequeñas alícuotas para evitar la congelación y descongelación repetida. </li><li> Preparar un gabinete de bioseguridad (BSC) de Clase II de alícuotas de virus en la limpieza de los BSC de los objetos innecesarios y esterilización de la superficie con un 70% EtOH. Coloque un vaso que contiene el 10% de solución de cloro en la campana de recogida de residuos contaminados con AAV. También se colocan en el BSC son tubos estériles de 0,5 ml y un recipiente con hielo seco. </li><li> Descongele stock viral en el hielo fuera de la BSC. </li><li> Vortex virus y abrir el tubo dentro del BSC. </li><li> Pipeta el volumen de la alícuota que desee (por ejemplo, 5 l) en un tubo de 0,5 ml, cerrar el tubo y colocarlo en el hielo seco para congelar el flash-virus. </li><li> Cuando todos los virus ha sido alícuotas, de desechar el tubo virus y puntas de pipeta en el recipiente de residuos que contienen cloro al 10%. </li><li> Retire el contenedor de residuos de la campana y añadir cloro adicional del 10%, luego verter el cloro en el fregadero. Elimine los residuos de plástico en un contenedor de riesgo biológico como se indica por el oficial de su institución en materia de bioseguridad. </li><li> Limpie cualquier equipo o las superficies que estuvieron en contacto con el virus con lejía al 10%. Deseche los guantes. </li><li> Almacenar en alícuotas de -80 ° C congelador. </li></ol> 2. Cirugía <ol><li> Cubra el área quirúrgica con papel absorbente mesa de laboratorio. Los instrumentos quirúrgicos deben ser esterilizados y al cabo la cirugía en condiciones asépticas, de acuerdo con la bioseguridad de su institución y las directrices el uso de animales. </li><li> Elija un área adyacente a la zona de la cirugía que se dedica a la carga de las micropipetas con el virus, y se cubre con papel absorbente mesa de laboratorio. Introducir una alícuota del virus en un contenedor de hielo en esta zona, y permiten que el virus de descongelar el hielo que la cirugía se está realizando. </li><li> Configuración de un contenedor de residuos de cloro al 10% en el área de virus dedicado a tratar de deshacerse de puntas de pipeta, etc, que entran en contacto con el virus. </li><li> Tire de una micropipeta de vidrio a una Wiretrol diámetro de la punta de aproximadamente 20 micras. Coloque una pequeña gota de aceite mineral en el extremo romo de la micropipeta e inserte el émbolo cable suministrado con la micropipeta. </li><li> Asegurar la micropipeta en la abrazadera del brazo microbomba. </li><li> Se inyecta por vía subcutánea con un ratón de la buprenorfina en dosis de 0,5 mg / kg. Anestesiar el ratón con Avertin través de una inyección intraperitoneal con una dosis de 200 mg / kg y eliminar el vello de la parte superior de la cabeza con unas tijeras quirúrgicas, asegurándose de dejar un amplio margen suficiente para evitar que el pelo de entrar en la incisión. </li><li> Coloque una manta de calefacción a la base de un stereotax para mantener una temperatura corporal de 37 ° C durante toda la cirugía y seguro con el ratón en la stereotax. </li><li> Bañe a la cabeza con tres matorrales alterna de etanol y betadine para esterilizar la zona. </li><li> Coloque una gota de pomada en cada ojo Tobradex para mantener los ojos húmedos durante la cirugía. </li><li> Hacer una incisión en la línea media de la cabeza y tirar de la piel para exponer el cráneo. </li><li> Retire con cuidado la fascia del cráneo con unas pinzas de punta fina. </li><li> Busque el área a ser inyectada usando coordenadas estereotáxica y marca el cráneo con un rotulador quirúrgico. </li><li> El uso de un taladro dental con una fresa de 1,4 mm, una delgada zona del cráneo de aproximadamente 2 mm de diámetro hasta que las grietas del cráneo, que divide la zona adelgazada en varios segmentos. </li><li> A lo largo de este proceso, mantén el cráneo húmedo con aplicaciones de solución salina estéril. </li><li> Realizar una craneotomía removiendo suavemente los segmentos del cráneo diluir con extra-fino con punta de pinzas. </li></ol> 3. Preparación de la inyección <ol><li> Con un KIMWIPE coloca sobre la tapa, abra el tubo del virus de la pipeta y 1.5uL (por una inyección de 1 ul) del virus en un pequeño trozo de parafilm. </li><li> Coloque la punta de la micropipeta en la cepa viral y retirar manualmente el émbolo. Si hay dificultades de dibujo stock viral en la micropipeta, la punta puede ser ligeramente más grande que un piercing KIMWIPE con la micropipeta para romper una pequeña parte de la copa. </li><li> Parte inferior del brazo microbomba en émbolo hasta que una pequeña cantidad de virus que se disipe de la punta de la micropipeta. Eliminar esta pequeña gota con un hisopo de algodón y el aplicador de descarte en las erascontenedor e. </li><li> Aplique una gota de aceite mineral en la punta de la micropipeta para evitar la obstrucción de la micropipeta se introduce en el cerebro. </li></ol> 4. Virus de la inyección <ol><li> Uso de X e Y estereotáxica coordenadas, la posición de la micropipeta sobre el área a ser inyectada. En este experimento, el estereotáxica coordenadas que se utilizan para localizar la corteza visual primaria es de 2,7 mm posterior a bregma y 2,5 mm lateral a la línea media. Muy baja lentamente la micropipeta (a una tasa aproximada de 1mm/1minute) a la posición adecuada Z. </li><li> Introduzca los parámetros de inyección deseado en la caja SYS Micro4 controlador microbomba e iniciar la inyección. Para este experimento, un microlitro de más de 10 minutos se puede utilizar como una tasa de inyección. </li><li> Cuando se acabe la inyección, deje la pipeta para descansar de uno hasta dos minutos para evitar el flujo de salida del virus durante la extracción. Después de este período, muy lentamente quitar la micropipeta en el cerebro (tasa igual al anterior). </li><li> Sutura en el cuero cabelludo y el sello con adhesivo tisular. Inyectar el animal por vía subcutánea con buprenorfina en dosis de 0,1 mg / kg cada 8-12 horas durante las próximas 72 horas, o hasta que el animal está mostrando signos de dolor. Que el animal pueda recuperarse bajo una lámpara de calor hasta que se deambular y listo para ser devuelto a su jaula. Experimentos de imagen puede ser iniciado después de que el tiempo de incubación deseado (días o semanas después de la inyección viral). </li></ol> 5. Limpieza <ol><li> Enjuague la micropipeta con lejía al 10% y desechar en un contenedor de objetos punzantes. </li><li> Disponga de los contenedores de residuos en la misma forma que la sección 1. </li><li> Deseche el papel del laboratorio en el banco de bin riesgo biológico y limpie todas las superficies y los instrumentos que pudieran haber entrado en contacto con el virus con lejía al 10%. </li><li> Virus utilizados pueden ser congelados de nuevo, teniendo en cuenta que repetidos de congelación / deshielo causan la degradación de los virus. </li></ol> 6. Resultados representante <img src="/files/ftp_upload/2140/2140fig1.jpg" alt="Figura 1" /> Figura 1. Transducidas neurona después de la inyección de doble hebra del virus adeno-asociado serotipo 1 (S1 dsAAV) la realización de la proteína verde fluorescente (GFP) bajo el control del promotor CMV. El cuerpo de la célula, así como las dendritas proximales y distales son claramente visibles en las secciones fijas fotografiado usando un microscopio de epifluorescencia. Barra de escala = 100 micras. <img src="/files/ftp_upload/2140/2140fig2.jpg" alt="Figura 2" /> Figura 2. Etiquetado típico de una inyección de virus intracraneal en corteza visual primaria con S1 dsAAV muestra el grado de propagación viral, así como neuronas marcadas, la glía y procesos. Barra de escala = 250 micras. <img src="/files/ftp_upload/2140/2140fig3.jpg" alt="Figura 3" /> Figura 3. Etiquetado de las células en el hipocampo con S1 dsAAV. Barra de escala = 250 micras. Estas cifras son una adaptación de Lowery et al. 2009 1

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

Los protocolos de los animales y experimentales fueron aprobados por la Universidad de la Universidad de Rochester Comité Internacional sobre los Recursos (UCAR), de conformidad con la Política de PHS en Cuidado Humano y Uso de Animales de Laboratorio.

Viral mediada por la entrega de genes tiene un gran potencial para el estudio de los procesos neurológicos y el tratamiento de trastornos cerebrales 1,2,3. La gran versatilidad de esta técnica también puede ser aprovechado para marcar con fluorescencia células para obtener imágenes tanto in vitro como in vivo 4. Aquí se demuestra un procedimiento detallado para la transducción de las neuronas y la glía en la corteza visual del ratón con una doble cadena adeno-virus que ex...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
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<tr>
<td>Stoelting Mouse and Neonatal Rat Adaptor</td>
<td>&nbsp;</td>
<td>Stoelting</td>
<td>51625</td>
<td>Regular stereotax for securing animals for surgery may be substituted</td>
</tr>
<tr>
<td>Extra Fine Bonn Scissors, 8.5cm, straight tip, cutting edge 13mm</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>14084-08</td>
<td>&nbsp;</td>
<tr>
<td>Eye Dressing Forceps, 10cm, tip width 0.5mm, curved</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11152-10</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #5/45 Forceps- Dumoxel Standard Tip, 11cm, angled</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11251-35</td>
<td>Extra-fine tipped forceps for performing craniotomy</td>
</tr>
<tr>
<td>Standard Pattern Forceps, straight, 2.5mmx1.35mmtip, 12cm</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11000-12</td>
<td>&nbsp;</td>
<tr>
<td>Microtorque Control Box and Tech2000 handpiece</td>
<td>&nbsp;</td>
<td>Ram Products, Inc.</td>
<td>TECH2000ON/OFF</td>
<td>Dental drill</td>
</tr>
<tr>
<td>Micro Drill Stainless Steel Burrs 1.4mm tip diameter</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>19008-14</td>
<td>&nbsp;</td>
<tr>
<td>Wiretrol micropipettes, to deliver 1-5 Ul</td>
<td>&nbsp;</td>
<td>VWR International/ Drummond</td>
<td>5-000-1001 or 53480-287</td>
<td>&nbsp;</td>
<tr>
<td>Mineral oil</td>
<td>&nbsp;</td>
<td>VWR International</td>
<td>29447-338</td>
<td>&nbsp;</td>
<tr>
<td>Manual Micromanipulator and Tilting Base (right-handed)</td>
<td>&nbsp;</td>
<td>World Precision Instruments, Inc.</td>
<td>M3301-M3-R</td>
<td>Used for determining stereotaxic co-ordinates</td>
</tr>
<tr>
<td>UltraMicroPump (UMP3) (one) with SYS-Micro4 Controller</td>
<td>&nbsp;</td>
<td>World Precision Instruments, Inc.</td>
<td>UMP3-1</td>
<td>&nbsp;</td>
<tr>
<td>Sutures</td>
<td>&nbsp;</td>
<td>VWR International</td>
<td>95056-952</td>
<td>&nbsp;</td>
<tr>
<td>P-97 Flaming/Brown Micropipette Puller</td>
<td>&nbsp;</td>
<td>Sutter Instruments</td>
<td>P-97</td>
<td>&nbsp;</td>
<tr>
<td>Tobradex</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>Available from your institution&#x2019;s veterinary services</td>
</tr>		</tbody>
		</table>
		</div>

intracranial injection of adeno-associated viral vectors

Inyección intracraneal de vectores virales para expresar una proteína fluorescente es una técnica de etiquetado versátil para la visualización de grupos específicos de células en diferentes regiones del cerebro, tanto en vivo y en secciones de cerebro. A diferencia de la inyección de colorantes fluorescentes, etiquetado viral ofrece focalización de los tipos de células y es menos costoso y consume mucho tiempo que el establecimiento de líneas de ratones transgénicos. En esta técnica, un virus adeno-asociados (AAV) vector se inyecta por vía intracraneal mediante coordenadas estereotáxica, una micropipeta y una bomba automática para la entrega precisa de AAV en la zona deseada con el mínimo daño a los tejidos circundantes. Los parámetros de inyección se pueden adaptar a los distintos experimentos mediante el ajuste de la edad del animal a una velocidad de inyección, la ubicación de la inyección, el volumen de la inyección, la inyección, el serotipo AAV y el promotor dirige la expresión de genes. Dependiendo de las condiciones elegidas, de forma viral inducida por la expresión del transgén puede permitir la visualización de grupos de células, las células individuales o bien los procesos celulares, hasta el nivel de las espinas dendríticas. El experimento que se muestra aquí muestra una inyección de doble cadena AAV que expresa la proteína verde fluorescente para el etiquetado de las neuronas y la glía en la corteza visual primaria del ratón.

Watch this Scientific Journal Video about Intracranial Injection of Adeno-associated Viral Vectors at JoVE.com

Intracranial Injection of Adeno-associated Viral Vectors

Aquí presentamos la inyección intracraneal de vectores AAV de marcaje fluorescente de las neuronas y la glía en la corteza visual.

Inyección intracraneal de adeno-asociado vectores virales

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

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

Research

JoVE Journal

Neuroscience

47.2K vistas.  University of Rochester. Aquí presentamos la inyección intracraneal de vectores AAV de marcaje fluorescente de las neuronas y la glía en la corteza visual.

Video: Intracranial Injection of Adeno-associated Viral Vectors

Targeting of Deep Brain Structures with Microinjections for Delivery of Drugs, Viral Vectors, or Cell Transplants

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an injecting needle produces during its trajectory is a major concern especially in the mouse brain for not only the brain is small but also sometimes multiple injections are needed.  We show here a method to produce glass capillary needles with a 50-&mu;m lumen which significantly reduces the brain damage and allows a precise targeting into the rodent brain. This method allows a delivery of small volumes (from 20 to 100 nl), reduces bleeding risks, and minimizes passive diffusion of drugs into the brain parenchyma. By using different size of capillary glass tubes, or changing the needle lumen, several types of substances and cells can be injected. Microinjections with a glass capillary tube represent a significant improvement in injection techniques and deep brain targeting with minimal collateral damage in small rodents.

In this article, we show a method to make glass capillary needles with a 50-&mu;m lumen. This technique significantly reduces the brain damage, minimizes passive diffusion of drugs and allows a precise targeting into the rodent brain.

Focalización de estructuras profundas del cerebro con microinyecciones de entrega de medicamentos, vectores virales, o trasplantes de células

Microinjections into the brain parenchyma are important procedures to deliver drugs, viral vectors or cell transplants.  The brain lesion that an ...

Investigación

Neurociencias

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

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene expression in the mouse brain. Here we describe a technique first introduced by Passini and Wolfe for direct intracranial delivery of virally-encoded transgenes into the neonatal mouse brain. In its most basic form, the procedure requires only an ice bucket and a microliter syringe. However, the protocol can also be adapted for use with stereotaxic frames to improve consistency for researchers new to the technique. The method relies on the ability of adeno-associated virus (AAV) to move freely from the cerebral ventricles into the brain parenchyma while the ependymal lining is still immature during the first 12-24 hr after birth. Intraventricular injection of AAV at this age results in widespread transduction of neurons throughout the brain. Expression begins within days of injection and persists for the lifetime of the animal. Viral titer can be adjusted to control the density of transduced neurons, while co-expression of a fluorescent protein provides a vital label of transduced cells. With the rising availability of viral core facilities to provide both off-the-shelf, pre-packaged reagents and custom viral preparation, this approach offers a timely method for manipulating gene expression in the mouse brain that is fast, easy, and far less expensive than traditional germline engineering.

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

Inyección Viral Intracerebroventricular de Neonatología del ratón Cerebro de transducción neuronal persistente y generalizada

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene ...

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.

Limbal Inyección-Enfoque Subretinal de vectores virales para terapia génica en ratones epitelio pigmentario de la retina

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 ...

Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the hematogenous route, which involves infection of the endothelial cells and pericytes of the brain. The second is the intracerebroventricular (ICV) route. Once within the central nervous system (CNS), viruses may spread to the subarachnoid space, meninges, and choroid plexus via the cerebrospinal fluid. In experimental models, the earliest stages of CNS viral distribution are not well characterized, and it is unclear whether only certain cells are initially infected. Here, we have analyzed the distribution of cytomegalovirus (CMV) particles during the acute phase of infection, termed primary viremia, following ICV or intravascular (IV) injection into the neonatal mouse brain. In the ICV injection model, 5 &#181;l of murine CMV (MCMV) or fluorescent microbeads were injected into the lateral ventricle at the midpoint between the ear and eye using a 10-&#181;l syringe with a 27 G needle. In the IV injection model, a 1-ml syringe with a 35 G needle was used. A transilluminator was used to visualize the superficial temporal (facial) vein of the neonatal mouse. We infused 50 &#181;l of MCMV or fluorescent microbeads into the superficial temporal vein. Brains were harvested at different time points post-injection. MCMV genomes were detected using the in situ hybridization method. Fluorescent microbeads or green fluorescent protein expressing recombinant MCMV particles were observed by fluorescent microscopy. These techniques can be applied to many other pathogens to investigate the pathogenesis of encephalitis.

Here, we describe a simple method of intracerebroventricular and intravascular injection of viral particles or fluorescent microbeads into the neonatal mouse brain. The localization pattern of the virus and nanoparticles could be detected by microscopic evaluation or by in situ hybridization.

Intracerebroventricular y por inyección intravascular de partículas víricas y fluorescentes microperlas en el cerebro neonatal

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the ...

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.

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.

Adenovirus Subagonal Asociado 9 (AAV9) Vector Entrega en Ratones Adultos

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

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.

Métodos y consejos para la administración intravenosa del Virus Adeno-asociado a las ratas y la evaluación de la transducción de señales del sistema nervioso Central

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

Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or triple transgenic mice are generated in order to achieve selective expression of a reporter or effector gene (e.g., from the Rosa26 locus) in the desired spinal population. 2) Intraspinal injection of Cre-dependent recombinant adeno-associated virus (rAAV); here Cre-dependent AAV vectors coding for the reporter or effector gene of choice are injected into the spinal cord of mice expressing Cre recombinase in the desired neuronal subpopulation. This protocol describes how to generate Cre-dependent rAAV vectors and how to transduce neurons in the dorsal horn of the lumbar spinal cord segments L3-L5 with rAAVs. As the lumbar spinal segments L3-L5 are innervated by those peripheral sensory neurons that transmit sensory information from the hindlimbs, spontaneous behavior and responses to sensory tests applied to the hindlimb ipsilateral to the injection side can be analyzed in order to interrogate the function of the manipulated neurons in sensory processing. We provide examples of how this technique can be used to analyze genetically defined subsets of spinal neurons. The main advantages of virus-mediated transgene expression in Cre transgenic mice compared to classical reporter mouse-induced transgene expression are the following: 1) Different Cre-dependent rAAVs encoding various reporter or effector proteins can be injected into a single Cre transgenic line, thus overcoming the need to create several multiple transgenic mouse lines. 2) Intraspinal injection limits manipulation of Cre-expressing cells to the injection site and to the time after injection. The main disadvantages are: 1) Reporter gene expression from rAAVs is more variable. 2) Surgery is required to transduce the spinal neurons of interest. Which of the two methods is more appropriate depends on the neuron population and research question to be addressed.

Intraspinal injection of recombinase dependent recombinant adeno-associated virus (rAAV) can be used to manipulate any genetically labelled cell type in the spinal cord. Here we describe how to transduce neurons in the dorsal horn of the lumbar spinal cord. This technique enables functional interrogation of the manipulated neuron subtype.

Expresión de transgenes de mediada por Virus adeno-asociado en neuronas genéticamente definidas de la médula espinal

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or ...

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.

Directa de la inyección intratecal de virus Adeno-asociado recombinantes en ratones adultos

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.

Producción, purificación y Control de calidad de vectores de base de Virus Adeno-asociado

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

Convection Enhanced Delivery of Optogenetic Adeno-associated Viral Vector to the Cortex of Rhesus Macaque Under Guidance of Online MRI Images

In non-human primate (NHP) optogenetics, infecting large cortical areas with viral vectors is often a difficult and time-consuming task. Here, we demonstrate the use of magnetic resonance (MR)-guided convection enhanced delivery (CED) of optogenetic viral vectors into primary somatosensory (S1) and motor (M1) cortices of macaques to obtain efficient, widespread cortical expression of light-sensitive ion channels. Adeno-associated viral (AAV) vectors encoding the red-shifted opsin C1V1 fused to yellow fluorescent protein (EYFP) were injected into the cortex of rhesus macaques under MR-guided CED. Three months post-infusion, epifluorescent imaging confirmed large regions of optogenetic expression (&#62;130 mm2) in M1 and S1 in two macaques. Furthermore, we were able to record reliable light-evoked electrophysiology responses from the expressing areas using micro-electrocorticographic arrays. Later histological analysis and immunostaining against the reporter revealed widespread and dense optogenetic expression in M1 and S1 corresponding to the distribution indicated by epifluorescent imaging. This technique enables us to obtain expression across large areas of the cortex within a shorter period of time with minimal damage compared to the traditional techniques and can be an optimal approach for optogenetic viral delivery in large animals such as NHPs. This approach demonstrates great potential for network-level manipulation of neural circuits with cell-type specificity in animal models evolutionarily close to humans.&#160;

Here, we demonstrate magnetic resonance (MR)-guided convection enhanced delivery (CED) of viral vectors into the cortex as an efficient and simplified approach for achieving optogenetic expression across large cortical areas in the macaque brain.