Ce travail a été rendu possible par des subventions du NIH (EY012977), une bourse de carrière dans les sciences biomédicales de l&#39;Burroughs Wellcome Fund, la Fondation de Whitehall, et la Sloan Foundation (AKM).

1. Manipulation du virus et le stockage <ol><li> Une protection adéquate et des techniques de manipulation doivent être choisis en fonction du niveau de biosécurité du virus à être utilisé. Ces pratiques peuvent être trouvés dans la biosécurité dans les laboratoires ème édition microbiologique et biomédicale 5, disponible sur le site du CDC ( <a href="http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm">http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm</a> ). L&#39;utilisation de vecteurs AAV est approuvé pour la biosécurité de niveau 1 (BSL-1). Pour l&#39;expérience présentée ici, une blouse et des gants seront portés lors de la manipulation du virus, conformément aux procédures de traitement BSL-1 agents. </li><li> Afin de préserver l&#39;activité du virus, il est préférable de le diviser en petites portions pour éviter le gel et le dégel répétés. </li><li> Préparer une enceinte de sécurité biologique (BSC) de classe II pour aliquotage virus par le BSC compensation de tous les objets inutiles et de stérilisation de la surface avec de l&#39;EtOH 70%. Placer un bécher contenant une solution javellisée à 10% dans la hotte de la collecte des déchets contaminés AAV. Également placée dans le BSC sont stériles tubes de 0.5ml et un récipient de glace sèche. </li><li> Dégel stock viral sur la glace en dehors de la BSC. </li><li> Virus de Vortex et ouvrir le tube à l&#39;intérieur du BSC. </li><li> Pipeter le volume aliquote désirée (par exemple 5 uL) dans un tube de 0.5ml, fermer le tube et le placer dans la glace sèche pour flash-geler le virus. </li><li> Lorsque tous les virus a été aliquotés débarrasser de ce tube de virus et des embouts de pipette dans le récipient de déchets contenant Javel à 10%. </li><li> Retirez le conteneur de déchets provenant de la hotte et d&#39;ajouter l&#39;eau de Javel à 10%, puis verser l&#39;eau de javel dans l&#39;évier. Eliminer les déchets en plastique dans un container prévu comme indiqué par l&#39;agent de votre institut de biosécurité. </li><li> Nettoyer tout équipement ou des surfaces qui sont entrés en contact avec le virus de Javel à 10%. Jeter les gants. </li><li> Conserver dans aliquotes à -80 ° C au congélateur. </li></ol> 2. Chirurgie <ol><li> Couvrir la zone chirurgicale avec un papier de laboratoire banc absorbant. Instruments chirurgicaux doivent être stérilisés et de la chirurgie effectuée dans des conditions aseptiques en conformité avec la biosécurité de votre institution et les directives d&#39;utilisation d&#39;animaux. </li><li> Choisissez une zone adjacente à la zone chirurgicale qui sera dédié au chargement des micropipettes avec le virus, et le couvrir avec du papier absorbant de laboratoire banc. Placer une aliquote de virus dans un récipient de glace dans ce domaine, et permettre au virus de dégel sur la glace que la chirurgie est effectuée. </li><li> Installation d&#39;un conteneur de déchets d&#39;eau de Javel à 10% dans la zone de manipulation du virus dédié pour l&#39;élimination des embouts de pipette, etc qui viennent en contact avec le virus. </li><li> Tirez une micropipette de verre Wiretrol à un diamètre de la pointe d&#39;environ 20 microns. Placer une petite goutte d&#39;huile minérale sur la fin brutale de la micropipette et insérez le piston de fil fourni avec le micropipettes. </li><li> Fixez la micropipette dans la pince du bras micropompe. </li><li> Injecter une souris avec la buprénorphine par voie sous cutanée à la dose de 0,5 mg / kg. Anesthésier la souris avec Avertin par injection intrapéritonéale avec une dose de 200 mg / kg et enlever les poils du dessus de la tête avec des ciseaux chirurgicaux, en veillant à laisser des marges assez grandes pour empêcher les cheveux d&#39;entrer dans l&#39;incision. </li><li> Fixer une couverture chauffante à la base d&#39;un stereotax pour maintenir une température corporelle de 37 ° C toute l&#39;opération et assurer la souris dans la stereotax. </li><li> Baignez la tête avec trois gommages alternance d&#39;éthanol et bétadine pour stériliser la région. </li><li> Déposer une goutte de pommade ophtalmique Tobradex sur chaque œil à garder les yeux humides pendant la chirurgie. </li><li> Faire une incision le long de la ligne médiane de la tête et tirer la peau du dos pour exposer le crâne. </li><li> Retirez délicatement la façade du crâne à l&#39;aide à pointe fine pince. </li><li> Localisez la zone à être injecté en utilisant les coordonnées stéréotaxiques et marquer le crâne avec un stylo chirurgical. </li><li> En utilisant une fraise de dentiste avec une bavure 1.4mm, fine une zone du crâne d&#39;environ 2mm de diamètre jusqu&#39;à la fissure du crâne, en divisant la zone amincie en plusieurs segments. </li><li> Tout au long de cette procédure, garder le crâne humide avec des applications de solution saline stérile. </li><li> Effectuer une craniotomie en enlevant doucement les segments du crâne à l&#39;aide aminci extra-fine pince à pointe. </li></ol> 3. Préparation à l&#39;injection <ol><li> Avec un Kimwipe placé sur le couvercle, ouvrir le tube de virus et de la pipette 1.5uL (pour une injection 1UL) du virus sur un petit morceau de parafilm. </li><li> Placez la pointe de la micropipette dans le stock viral et manuellement retirer le piston. En cas de difficulté de dessin stock viral dans la micropipette, la pointe peut être agrandi être légèrement percer un Kimwipe à la micropipette pour casser une petite partie de la vitre. </li><li> Bras inférieur sur micropompe piston jusqu&#39;à ce qu&#39;une petite quantité de virus est dissipé depuis la pointe de la micropipette. Retirer cette petite goutte avec un coton-tige et l&#39;applicateur jeter dans étaiscontenant e. </li><li> Appliquez une goutte d&#39;huile minérale à la pointe de la micropipette pour éviter le colmatage de la micropipette est descendue dans le cerveau. </li></ol> 4. Injection du virus <ol><li> Utilisation de X et Y coordonnées stéréotaxique, la position de la micropipette sur la zone à injecter. Dans cette expérience, la stéréotaxie coordonnées utilisé pour localiser cortex visuel primaire sont postérieurs à 2,7 mm et 2,5 mm bregma latéral à la ligne médiane. Très lentement le micropipette (à un taux approximatif de 1mm/1minute) à la bonne position Z. </li><li> Entrez les paramètres d&#39;injection souhaitée dans la boîte SYS Micro4 contrôleur de micropompe et initier l&#39;injection. Pour cette expérience, 1 microlitre de plus de 10 minutes sera utilisé comme un taux d&#39;injection. </li><li> Lorsque l&#39;injection est terminée, laisser la pipette au repos pendant une à deux minutes pour empêcher l&#39;efflux de virus lors de l&#39;enlèvement. Après cette période, très lentement retirer la micropipette par le cerveau (même taux que ci-dessus). </li><li> Suture du cuir chevelu et le sceller avec de la colle tissulaire. Injecter de l&#39;animal sous-cutanée avec la buprénorphine à la dose de 0,1 mg / kg toutes les 8-12 heures au cours des 72 prochaines heures, ou aussi longtemps que l&#39;animal manifeste des signes de douleur. Permettre à l&#39;animal de retrouver sous une lampe chauffante jusqu&#39;à ce qu&#39;il soit ambulantes et prêt à être retourné à sa cage. Des expériences d&#39;imagerie peuvent être engagées après le temps d&#39;incubation désirée (jours ou semaines après l&#39;injection virale). </li></ol> 5. Nettoyage <ol><li> Rincer la micropipette avec l&#39;eau de Javel à 10% et le jeter dans un contenant pour objets pointus. </li><li> Jeter le contenant des déchets de la même manière que la section 1. </li><li> Jeter le papier dans le bac paillasse biorisques et essuyez toutes les surfaces et les instruments qui ont pu entrer en contact avec le virus de l&#39;eau de Javel à 10%. </li><li> Le virus inutilisés peuvent être congelés à nouveau, en gardant à l&#39;esprit que répétées cycles gel / dégel provoque la dégradation des virus. </li></ol> 6. Les résultats représentatifs <img src="/files/ftp_upload/2140/2140fig1.jpg" alt="Figure 1" /> Figure 1. Transduction neurone après l&#39;injection du double brin du virus adéno-associé de sérotype 1 (dsAAV S1) transportant la protéine fluorescente verte (GFP) sous contrôle du promoteur CMV. Le corps de la cellule ainsi que les dendrites proximales et distales sont clairement visibles dans les sections fixes imagée en utilisant un microscope à épifluorescence. Barre d&#39;échelle = 100 pm. <img src="/files/ftp_upload/2140/2140fig2.jpg" alt="Figure 2" /> Figure 2. Étiquetage typiques d&#39;une injection de virus intracrânienne dans le cortex visuel primaire en utilisant S1 dsAAV montrant l&#39;ampleur de la propagation virale ainsi que des neurones marqués, glie et les processus. Barre d&#39;échelle = 250 um. <img src="/files/ftp_upload/2140/2140fig3.jpg" alt="Figure 3" /> Figure 3. Étiquetage des cellules dans l&#39;hippocampe en utilisant S1 dsAAV. Barre d&#39;échelle = 250 um. Ces chiffres sont adaptés 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>

Les protocoles expérimentaux animaux et ont été approuvés par l&#39;Université de Rochester University Comité des Ressources Animales (UCAR) en conformité avec le PHS politique sur les soins aux animaux et l&#39;utilisation des animaux de laboratoire.

Livraison de gènes médiée par Virally détient un grand potentiel pour l&#39;étude des processus neurologiques et le traitement des troubles du cerveau 1,2,3. La grande polyvalence de cette technique peut aussi être exploitée pour marquer les cellules par fluorescence pour l&#39;imagerie in vitro et in vivo 4. Ici nous démontrons une procédure détaillée pour la transduction de neurones et cellules gliales dans le cortex visuel de la souris en utilisant un double brin d&#...

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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>
		</tr>
		</thead>
		<tbody>
		
<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

L&#39;injection intracrânienne de vecteurs viraux pour exprimer une protéine fluorescente est une technique de marquage polyvalent pour la visualisation des sous-ensembles spécifiques de cellules dans différentes régions du cerveau in vivo et dans les sections du cerveau. Contrairement à l&#39;injection de colorants fluorescents, l&#39;étiquetage virale offre ciblant des types cellulaires individuels et est moins coûteux et fastidieux que d&#39;établir des lignées de souris transgéniques. Dans cette technique, une viraux adéno-associés (AAV) est injecté par voie intracrânienne en utilisant les coordonnées stéréotaxique, une micropipette et une pompe automatique pour la livraison précise de l&#39;AAV à la zone désirée avec un minimum de dommages aux tissus environnants. Paramètres d&#39;injection peuvent être adaptées à des expériences individuelles en ajustant l&#39;âge des animaux à un taux d&#39;injection, l&#39;emplacement d&#39;injection, le volume d&#39;injection, d&#39;injection, AAV de sérotype et le promoteur dirigeant l&#39;expression des gènes. Selon les conditions choisies, viro-induit l&#39;expression du transgène peut permettre la visualisation des groupes de cellules, des cellules individuelles ou d&#39;une amende de processus cellulaires, jusqu&#39;au niveau des épines dendritiques. L&#39;expérience montre ici représente une injection de double-brin AAV exprimant la protéine fluorescente verte pour l&#39;étiquetage des neurones et cellules gliales dans le cortex visuel primaire de souris.

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

Intracranial Injection of Adeno-associated Viral Vectors

Nous présentons ici l&#39;injection intracrânienne de vecteurs AAV pour le marquage fluorescent des neurones et cellules gliales dans le cortex visuel.

Injection intracrânienne des vecteurs adéno-associés virale

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

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JoVE Journal

Neuroscience

47.1K vues.  University of Rochester. Nous présentons ici l'injection intracrânienne de vecteurs AAV pour le marquage fluorescent des neurones et cellules gliales dans le cortex visuel.

Vidéo: 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.

Ciblage des structures cérébrales profondes avec des microinjections de livraison des médicaments, des vecteurs viraux, ou de cellules

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

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Neurosciences

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 injection Approche-sous-rétinienne de vecteurs viraux pour la thérapie génique chez la souris épithélium pigmentaire rétinien

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 ...

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.

La livraison de vecteur 9 (AAV9) associée au virus Adeno-Associé Subpial (AAV9) chez les souris adultes

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

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.

Adeno-associated Virus-mediated Transgene Expression dans les neurones de la moelle épinière génétiquement définies

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.

Direct Injection intrathécale de Recombinant Adeno-associated virus chez la souris adulte

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.

Production, Purification et contrôle de la qualité pour les vecteurs de base de Virus Adeno-associé

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.

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

In non-human primate (NHP) optogenetics, infecting large cortical areas with viral vectors is often a difficult and time-consuming task. Here, we ...

In Vivo Direct Reprogramming of Resident Glial Cells into Interneurons by Intracerebral Injection of Viral Vectors

Converting resident glia in the brain into functional and subtype-specific neurons in vivo provides a step forward towards the development of alternative cell replacement therapies while also creating tools to study cell fate in situ. To date, it has been possible to obtain neurons via in vivo reprogramming, but the precise phenotype of these neurons or how they mature has not been analyzed in detail. In this protocol, we describe a more efficient conversion and cell-specific identification of the in vivo reprogrammed neurons, using an AAV-based viral vector system. We also provide a protocol for functional assessment of the reprogrammed cells&#8217; neuronal maturation. By injecting flip-excision (FLEX) vectors, containing the reprogramming and synapsin-driven reporter genes&#160;to specific cell types in the brain that serve as the target for cell reprogramming. This technique allows for the easy identification of newly reprogrammed neurons. Results show that the obtained reprogrammed neurons functionally mature over time, receive synaptic contacts and show electrophysiological properties of different types of interneurons. Using the transcription factors Ascl1, Lmx1a and Nurr1, the majority of the reprogrammed cells have properties of fast-spiking, parvalbumin-containing interneurons.

This protocol aims at generating directly reprogrammed interneurons in vivo, using an AAV-based viral system in the brain and a FLEX synapsin-driven GFP reporter, which allows for cell identification and further analysis in vivo.

In Vivo Reprogrammation directe des cellules gliales résidentes en interneurones par injection intracérébrale de vecteurs viraux

Converting resident glia in the brain into functional and subtype-specific neurons in vivo provides a step forward towards the development of alternative ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

Enregistrement électrocorticographique des zones du cortex cérébral manipulées à l’aide d’un virus adéno-associé ciblant la cofiline chez la souris

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...

Analyzing the Parkinson's Disease Mouse Model Induced by Adeno-associated Viral Vectors Encoding Human &#945;-Synuclein

Parkinson&#39;s disease is a neurodegenerative disorder that involves the death of the dopaminergic neurons of the nigrostriatal pathway and, consequently, the progressive loss of control of voluntary movements. This neurodegenerative process is triggered by the deposition of protein aggregates in the brain, which are mainly constituted of &#945;-synuclein. Several studies have indicated that neuroinflammation is required to develop the neurodegeneration associated with Parkinson&#39;s disease. Notably, the neuroinflammatory process involves microglial activation as well as the infiltration of peripheral T cells into the substantia nigra (SN). This work analyzes a mouse model of Parkinson&#39;s disease that recapitulates microglial activation, T-cell infiltration into the SN, the neurodegeneration of nigral dopaminergic neurons, and motor impairment. This mouse model of Parkinson&#39;s disease is induced by the stereotaxic delivery of adeno-associated viral vectors encoding the human wild-type &#945;-synuclein (AAV-h&#945;Syn) into the SN. The correct delivery of viral vectors into the SN was confirmed using control vectors encoding green fluorescent protein (GFP). Afterward, how the dose of AAV-h&#945;Syn administered in the SN affected the extent of h&#945;Syn expression, the loss of nigral dopaminergic neurons, and motor impairment were evaluated. Moreover, the dynamics of h&#945;Syn expression, microglial activation, and T-cell infiltration were determined throughout the time course of disease development. Thus, this study provides critical time points that may be useful for targeting synuclein pathology and neuroinflammation in this preclinical model of Parkinson&#39;s disease.

This work analyzes the vector dose and exposure time required to induce neuroinflammation, neurodegeneration, and motor impairment in this preclinical model of Parkinson&#39;s disease. These vectors encoding the human &#945;-synuclein are delivered into the substantia nigra&#160;to recapitulate the synuclein pathology associated with Parkinson&#39;s disease.