Questo lavoro è stato reso possibile da finanziamenti del NIH (EY012977), un Premio alla Carriera nel Scienze Biomediche dal Fondo Burroughs Wellcome, la Fondazione Whitehall, e la Fondazione Sloan (AKM).

1. Virus Manipolazione e stoccaggio <ol><li> Adeguata protezione e le tecniche di manipolazione devono essere scelti in base al livello di biosicurezza del virus da utilizzare. Queste pratiche possono essere trovati in biosicurezza in microbiologiche e Biomedica Laboratori 5 ° edizione, disponibile sul sito del 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;uso di vettori AAV è approvato per la Biosicurezza Livello 1 (BSL-1). Per l&#39;esperimento mostrato qui, un camice e guanti da indossare durante la manipolazione del virus, secondo le procedure per la gestione BSL-1 agenti. </li><li> Per preservare l&#39;attività del virus è meglio dividerlo in piccole aliquote per evitare congelamenti e scongelamenti ripetuti. </li><li> Preparare una biosicurezza Cabinet (BSC) Classe II per aliquoting virus deselezionando la BSC di tutti gli oggetti inutili e sterilizzare la superficie con il 70% EtOH. Posizionare un bicchiere contenente una soluzione di candeggina al 10% nel cappuccio per la raccolta di AAV-rifiuti contaminati. Anche messo in BSC sono sterili tubi da 0,5 ml e un contenitore di ghiaccio secco. </li><li> Disgelo magazzino virale sul ghiaccio al di fuori del BSC. </li><li> Vortex virus e aprire il tubo all&#39;interno del BSC. </li><li> Pipettare il volume aliquota desiderata (ad esempio 5 mL) in una provetta da 0,5 ml, chiudere il tubo e posizionarlo nel ghiaccio secco di flash-congelare il virus. </li><li> Quando tutti i virus è stato aliquotati, smaltire il tubo di virus e puntali nel contenitore dei rifiuti contenente il 10% di candeggina. </li><li> Rimuovere il contenitore dal cofano e aggiungere ulteriori candeggina al 10%, poi versare la candeggina nel lavandino. Smaltire i rifiuti in plastica in un contenitore rischio biologico secondo le istruzioni del funzionario biosicurezza del vostro Istituto. </li><li> Pulire le attrezzature e superfici che è venuto a contatto con il virus con il 10% di candeggina. Gettare i guanti. </li><li> Aliquote da conservare in freezer -80 ° C. </li></ol> 2. Chirurgia <ol><li> Coprire l&#39;area chirurgica con carta assorbente da banco di laboratorio. Strumenti chirurgici devono essere sterilizzati e chirurgia effettuata in condizioni asettiche secondo biosicurezza del vostro istituto e le linee guida sull&#39;uso degli animali. </li><li> Scegliere una zona adiacente all&#39;area chirurgica che sarà dedicata al caricamento dei micropipette con il virus, e coprire con carta assorbente da banco di laboratorio. Inserire un aliquota del virus in un contenitore di ghiaccio in questo settore, e permettono al virus di disgelo sul ghiaccio, come l&#39;intervento è in corso. </li><li> Installazione di un contenitore dei rifiuti del 10% candeggina nell&#39;area dedicata trattamento del virus per lo smaltimento di puntali, ecc che vengono a contatto con il virus. </li><li> Tirare una micropipetta Wiretrol vetro a un diametro punta di circa 20 micron. Mettere una piccola goccia di olio minerale sulla punta smussata della micropipetta e inserire lo stantuffo cavo fornito con il micropipette. </li><li> Fissare la micropipetta nel morsetto del braccio micropompa. </li><li> Iniettare un mouse con buprenorfina per via sottocutanea alla dose di 0,5 mg / kg. Anestetizzare il mouse con Avertin tramite iniezione intraperitoneale con una dose di 200 mg / kg e togliere i capelli dalla parte superiore della testa con le forbici chirurgiche, avendo cura di lasciare margini abbastanza grandi per evitare che i capelli di entrare l&#39;incisione. </li><li> Fissare una coperta riscaldamento alla base di un stereotax di mantenere una temperatura corporea di 37 ° C durante tutto l&#39;intervento e fissare il mouse nella stereotax. </li><li> Bagnano la testa con tre scrub alternati di etanolo e betadine per sterilizzare la zona. </li><li> Mettere una goccia di unguento oculare Tobradex su ciascun occhio a tenere gli occhi umidi durante l&#39;intervento. </li><li> Fare un&#39;incisione lungo la linea mediana della testa e tirare la pelle torna a esporre il cranio. </li><li> Rimuovere con attenzione la fascia dal cranio con punta fine pinze. </li><li> Individuare l&#39;area per essere iniettato usando le coordinate stereotassica e segnare il cranio con una penna chirurgica. </li><li> Utilizzando un trapano per l&#39;odontoiatria a 1,4 millimetri radica, sottile una zona del cranio circa 2mm di diametro fino a quando le crepe del cranio, dividendo l&#39;area diluito in diversi segmenti. </li><li> In tutta questa procedura, tenere il cranio umido con le applicazioni di soluzione fisiologica sterile. </li><li> Eseguire una craniotomia con delicatezza eliminando i segmenti cranio diluito con pinze a punta extra-fine. </li></ol> 3. Iniezione di preparazione <ol><li> Con un KimWipe posto sopra il coperchio, aprire il tubo da virus e pipetta 1.5uL (per una iniezione 1UL) di virus su un piccolo pezzo di parafilm. </li><li> Posizionare la punta della micropipetta in azioni virali e ritirare manualmente lo stantuffo. In caso difficoltà disegno magazzino virale nel micropipetta, la punta può essere allargato leggermente pungente essere uno KimWipe con la micropipetta per rompere una piccola parte del vetro. </li><li> Micropompa braccio inferiore sul stantuffo fino a quando una piccola quantità di virus è dissipato dalla punta della micropipetta. Rimuovere questa piccola goccia con un cotton-fioc e applicatore scartare in ericontenitore e. </li><li> Applicare una goccia di olio minerale per la punta della micropipetta per evitare l&#39;intasamento come la micropipetta è abbassato nel cervello. </li></ol> 4. Virus iniezione <ol><li> Utilizzo di X e Y stereotassica coordinate, la posizione del micropipetta sopra la zona da iniettare. In questo esperimento, il stereotassica coordinate utilizzato per individuare corteccia visiva primaria sono posteriori al bregma 2,7 millimetri e 2,5 mm laterale alla linea mediana. Molto lentamente abbassare la micropipetta (ad un tasso di circa 1mm/1minute) nella posizione corretta Z. </li><li> Immettere i parametri di iniezione desiderato nel controller SYS Micro4 casella micropompa e avviare l&#39;iniezione. Per questo esperimento, 1 microlitro in 10 minuti sarà utilizzato come un tasso di iniezione. </li><li> Quando l&#39;iniezione è terminata, lasciare la pipetta riposare per 1-2 minuti per evitare di efflusso del virus durante la rimozione. Dopo questo periodo, molto lentamente la micropipetta rimuovere dal cervello (tasso come sopra). </li><li> Sutura al cuoio capelluto e sigillare con la colla dei tessuti. Iniettare l&#39;animale con buprenorfina per via sottocutanea alla dose di 0,1 mg / kg ogni 8-12 ore per i prossimi 72 ore, o fino a quando l&#39;animale è mostrano segni di dolore. Permettere agli animali di recuperare sotto una lampada di calore fino a quando non è deambulazione e pronto per essere restituito alla sua gabbia. Esperimenti di imaging possono essere avviate dopo il tempo di incubazione desiderato (giorni o settimane dopo l&#39;iniezione virale). </li></ol> 5. Pulizia <ol><li> Sciacquare la micropipetta con il 10% di candeggina e smaltirlo in un apposito contenitore. </li><li> Smaltire i contenitori dei rifiuti nello stesso modo come Sezione 1. </li><li> Eliminare la carta banco di laboratorio in bin rischio biologico e pulire tutte le superfici e gli strumenti che possano essere venuti in contatto con il virus con il 10% di candeggina. </li><li> Virus non utilizzati possono essere ricongelato, tenendo presente che ripetuti cicli gelo / disgelo causa la degradazione del virus. </li></ol> 6. Rappresentante Risultati <img src="/files/ftp_upload/2140/2140fig1.jpg" alt="Figura 1" /> Figura 1. Trasdotte neurone dopo l&#39;iniezione del doppio filamento del virus adeno-associato sierotipo 1 (dsAAV S1) portando la proteina fluorescente verde (GFP) sotto il controllo del promotore CMV. Il corpo della cellula e dendriti prossimale e distale sono chiaramente visibili in sezioni fisse fotografato con un microscopio epifluorescente. Barra di scala = 100 micron. <img src="/files/ftp_upload/2140/2140fig2.jpg" alt="Figura 2" /> Figura 2. Etichettatura tipici di una iniezione di virus intracranica in corteccia visiva primaria con S1 dsAAV indicando la misura della propagazione virale così come neuroni marcati, glia e processi. Barra di scala = 250 micron. <img src="/files/ftp_upload/2140/2140fig3.jpg" alt="Figura 3" /> Figura 3. Etichettatura delle cellule dell&#39;ippocampo con S1 dsAAV. Barra di scala = 250 micron. Queste cifre sono adattate da 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>

I protocolli di sperimentazione animale e sono stati approvati dalla University of Rochester University commissione Animal Resources (UCAR) in conformità con la Politica PHS sulla cura umanitaria ed uso di animali da laboratorio.

Virale mediata consegna del gene ha un grande potenziale per lo studio di processi neurologici e trattamento dei disturbi cerebrali 1,2,3. La grande versatilità di questa tecnica può anche essere sfruttato per un&#39;etichetta fluorescente cellule per l&#39;imaging sia in vitro che in vivo 4. Qui mostriamo una procedura dettagliata per la trasduzione di neuroni e glia nella corteccia visiva del mouse utilizzando un doppio filamento del virus adeno-associazione che esprime maggiore proteina verde fluorescente. Mentre questa tecnica è relativamente semplice, ci sono una serie di dettagli tecnici che devono essere prese in considerazione. Un fattore importante è l&#39;iniezione indotto danni tissutali - perciò è fondamentale usare molta cautela durante l&#39;intervento e l&#39;inserimento / rimozione del micropipetta. Un intervento chirurgico non può comportare l&#39;alterazione delle strutture da acquisire. Inoltre, dopo il caricamento del virus nella micropipetta, grande cura deve essere presa per eliminare eventuali bolle d&#39;aria sulla punta della pipetta e garantire il volume corretto viene iniettato. Abbassamento del micropipetta leggermente oltre l&#39;obiettivo coordinata Z poi ritirarsi nella posizione corretta può prevenire virus traboccante fuori dal sito di iniezione. Quando si sceglie un tempo di incubazione, che permette un tempo adeguato per l&#39;espressione genica virale dovrebbe essere considerato. Questa varierà in base al virus utilizzato. Una risposta immunitaria limitata può essere suscitata da iniezione. Nella nostra esperienza, questa infiammazione è generalmente limitato al percorso dell&#39;ago e si può evitare di imaging lontano dalla vista di iniezione (circa 50 μms o più). Infine, se le sezioni del cervello devono essere montati su vetrini, l&#39;uso di un mezzo di montaggio Antifade è suggerita per mantenere la fluorescenza. Intracranica iniezione di vettori virali ha diversi vantaggi tecnici rispetto alle tecniche di etichettatura altri. Attraverso l&#39;uso di coordinate stereotassiche e modulando il volume iniettato, etichetta fluorescenti possono essere localizzate con precisione ad una zona di interesse. La quantità di trasduzione può essere modificato regolando il volume di iniezione, il titolo del virus o il tempo di sopravvivenza, consentendo la visualizzazione di entrambi i gruppi di cellule o di singole celle. Inoltre, l&#39;utilizzo di virus diversi (ad esempio lentivirus, herpes virus, l&#39;adenovirus) può anche modulare i tempi e le quantità di proteine ​​fluorescenti espressa così come i tipi di cellule bersaglio. Nel complesso, questa tecnica può fornire per l&#39;etichettatura molto versatile di elementi differenti del cervello per l&#39;imaging.

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<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

Intracranica iniezione di vettori virali ingegnerizzati per esprimere una proteina fluorescente è una tecnica di etichettatura versatile per la visualizzazione di sottoinsiemi di cellule nelle differenti regioni del cervello sia in vivo che in sezioni cerebrali. A differenza della iniezione di coloranti fluorescenti, etichettatura virale offre targeting di tipi di cellule individuali ed è meno costoso e richiede tempo che stabilire linee di topi transgenici. In questa tecnica, un virale adeno-associato (AAV) vettore viene iniettato intracranially usando le coordinate stereotassica, una micropipetta e una pompa automatica per la consegna precisa AAV alla zona desiderata con minimo danno al tessuto circostante. Parametri di iniezione può essere adattata alle singole sperimentazioni regolando l&#39;età degli animali a velocità di iniezione, la posizione di iniezione, il volume di iniezione, di iniezione, AAV sierotipo e il promotore di guida l&#39;espressione genica. A seconda delle condizioni scelte, virale indotta l&#39;espressione del transgene può permettere la visualizzazione di gruppi di cellule, cellule singole o multa processi cellulari, fino al livello delle spine dendritiche. L&#39;esperimento qui rappresenta una iniezione di AAV a doppio filamento che esprimono la proteina fluorescente verde per l&#39;etichettatura dei neuroni e glia nella corteccia visiva primaria del mouse.

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

Intracranial Injection of Adeno-associated Viral Vectors

Qui vi presentiamo l&#39;iniezione intracranica di vettori AAV per la marcatura fluorescente di neuroni e glia nella corteccia visiva.

Iniezione intracranica di adeno-associato vettori virali

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

47.2K Views.  University of Rochester. Qui vi presentiamo l'iniezione intracranica di vettori AAV per la marcatura fluorescente di neuroni e glia nella corteccia visiva.

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.

Targeting di strutture cerebrali profonde con microiniezioni di consegna dei farmaci, vettori virali, o trapianti di cellule

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

Ricerca

Neuroscienze

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.

Intracerebroventricular Iniezione virale del cervello neonatale mouse per persistente e diffusa trasduzione neuronale

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.

Limbare Approccio-sottoretinico iniezione di vettori virali per la terapia genica nei topi retina epitelio pigmentato

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 e intravascolare Iniezione di Viral particelle e fluorescenti microperle nel cervello neonatale

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.

Subito Adeno associato Virus 9 (AAV9) Consegna vettoriale nei topi adulti

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.

Metodi e consigli per la somministrazione endovenosa di Virus Adeno-associato ai ratti e valutazione di trasduzione del sistema nervoso centrale

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.

Espressione del Transgene di mediata da Virus adeno-associato in neuroni geneticamente definiti del midollo spinale

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.

Diretto iniezione intratecale di ricombinanti Virus Adeno-associato in topi adulti

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.

Produzione, purificazione e controllo di qualità per i vettori basati su Virus Adeno-associato

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.