Este trabalho só foi possível por concessões do NIH (EY012977), um Prémio Carreira na de Ciências Biomédicas da Burroughs Wellcome Fund, a Fundação Whitehall, e da Fundação Sloan (AKM).

1. Manuseio e Armazenamento de vírus <ol><li> Uma protecção adequada e técnicas de manejo devem ser escolhidos com base no nível de biossegurança do vírus para ser usado. Estas práticas podem ser encontrados em Biossegurança em Microbiologia e Biomédica ª edição Laboratories 5, disponível no site do CDC ( <a href="http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm">http://www.cdc.gov/od/OHS/biosfty/bmbl5/bmbl5toc.htm</a> ). O uso de vetores AAV é aprovado para o Nível de Biossegurança 1 (NB-1). Para o experimento mostrado aqui, um jaleco e luvas vai ser usado durante o manuseamento do vírus, de acordo com os procedimentos para a manipulação de NBS-1 agentes. </li><li> Para preservar a atividade do vírus é melhor dividi-lo em pequenas alíquotas para evitar o congelamento e descongelamento repetidos. </li><li> Prepare um Gabinete de Biossegurança (BSC) Classe II de alíquotas de vírus por limpar o BSC de quaisquer objetos desnecessários e esterilizar a superfície com EtOH 70%. Coloque um copo contendo solução de água sanitária a 10% no capô para a recolha de resíduos contaminados AAV. Também colocado no BSC são estéreis tubos de 0,5 ml e um recipiente de gelo seco. </li><li> Descongelar estoque viral no gelo fora do BSC. </li><li> Vortex vírus e abrir o tubo dentro do BSC. </li><li> Pipetar volume da alíquota desejada (por exemplo, 5 mL) em um tubo de 0,5 ml, fechar o tubo e colocá-lo no gelo seco para flash-freeze o vírus. </li><li> Quando todos os vírus foi aliquotado, alienar o tubo de vírus e ponteiras no contentor de resíduos contendo lixívia a 10%. </li><li> Remova o recipiente de resíduos da capa e adicionar água sanitária a 10% adicionais, em seguida, despeje a lixívia na pia. Descarte de resíduos plásticos em um recipiente biohazard conforme instruído pelo oficial de seu instituto de biossegurança. </li><li> Limpar qualquer equipamento ou superfícies que entraram em contato com o vírus com água sanitária a 10%. Descarte as luvas. </li><li> Alíquotas armazenar em freezer -80 C °. </li></ol> 2. Cirurgia <ol><li> Cobrir a área cirúrgica com papel absorvente laboratório banco. Instrumentos cirúrgicos devem ser esterilizados e cirurgia feita sob condições assépticas, de acordo com a biossegurança de sua instituição e as diretrizes de uso animal. </li><li> Escolha uma área adjacente à área cirúrgica, que será dedicado a carregar o micropipetas com o vírus, e cobri-lo com papel absorvente laboratório banco. Transferir uma parte alíquota do vírus em um recipiente de gelo nesta área e permitir que o vírus para descongelar em gelo como a cirurgia está sendo realizada. </li><li> Configuração de um recipiente de resíduos de lixívia a 10% na área dedicada vírus manuseio para a eliminação de pontas de pipeta, etc, que entram em contato com o vírus. </li><li> Puxe uma micropipeta de vidro Wiretrol a um diâmetro da ponta de aproximadamente 20 microns. Coloque uma pequena gota de óleo mineral na extremidade sem corte da micropipeta e inserir o êmbolo de arame fornecido com o micropipetas. </li><li> Assegurar a micropipeta no grampo do braço micropump. </li><li> Injetar um rato por via subcutânea com buprenorfina na dose de 0,5 mg / kg. Anestesiar o rato com Avertin via intraperitoneal com uma dose de 200 mg / kg e remover o cabelo do topo da cabeça com uma tesoura cirúrgica, certificando-se de deixar margens grande o suficiente para impedir que o cabelo de entrar na incisão. </li><li> Anexar uma manta de aquecimento à base de um stereotax para manter uma temperatura corporal de 37 ° C durante toda a cirurgia e segura o mouse na stereotax. </li><li> Banhar a cabeça com três esfrega alternadas de etanol e betadine para esterilizar a área. </li><li> Coloque uma gota de TOBRADEX pomada em cada olho para manter os olhos úmidos durante a cirurgia. </li><li> Faça uma incisão abaixo da linha média da cabeça e puxe a pele de volta para expor o crânio. </li><li> Remova cuidadosamente a fáscia do crânio usando uma pinça de ponta fina. </li><li> Localizar a área a ser injetado usando coordenadas estereotáxica e marcar o crânio com uma caneta cirúrgica. </li><li> Usando uma broca de dentista com um 1,4 milímetros rebarba, uma área fina do crânio de aproximadamente dois milímetros de diâmetro, até as rachaduras do crânio, dividindo a área diluído em vários segmentos. </li><li> Todo esse procedimento, manter o crânio úmido com aplicações de soro fisiológico estéril. </li><li> Realizar uma craniotomia delicadamente removendo os segmentos crânio diluído com extra-fina com ponta fórceps. </li></ol> 3. Preparação de injeção <ol><li> Com um KimWipe colocado sobre a tampa, abra o tubo de vírus e de pipeta 1.5uL (para uma injeção 1uL) do vírus em um pequeno pedaço de parafilme. </li><li> Coloque a ponta da micropipeta para o estoque viral e retirar manualmente o êmbolo. Se a dificuldade é experimentada desenho estoque viral para a micropipeta, a ponta pode ser ligeiramente aumentada ser uma perfuração KimWipe com a micropipeta para quebrar uma pequena parte do vidro. </li><li> Braço inferior micropump em êmbolo até uma pequena quantidade de vírus é dissipada a partir da ponta da micropipeta. Remover esta pequena gota com um cotonete e aplicador de descarte em erascontainer e. </li><li> Aplique uma gota de óleo mineral para a ponta da micropipeta para evitar o entupimento como a micropipeta é abaixado no cérebro. </li></ol> 4. Injeção de vírus <ol><li> Usando o X e Y estereotáxica coordenadas, a posição da micropipeta sobre a área a ser injetada. Neste experimento, os estereotáxica coordenadas utilizadas para localizar córtex visual primário é posterior ao bregma 2,7 milímetro e 2,5 mm lateral à linha média. Muito lentamente o micropipeta (a uma taxa aproximada de 1mm/1minute) para a posição Z adequada. </li><li> Digite os parâmetros de injeção desejada na caixa de controlador de SYS MICRO4 micropump e iniciar a injeção. Para esta experiência, 1 microlitro mais de 10 minutos será usado como uma taxa de injeção. </li><li> Quando a injeção é terminar, deixe a pipeta para descansar por 1-2 minutos para evitar o efluxo do vírus durante a remoção. Após este período, muito lentamente remover a micropipeta do cérebro (mesma taxa acima). </li><li> Sutura no couro cabeludo e selá-lo com cola de tecido. Injetar o animal por via subcutânea com buprenorfina na dose de 0,1 mg / kg a cada 8-12 horas nas próximas 72 horas, ou enquanto o animal está exibindo sinais de dor. Permitir que o animal se recuperar sob uma lâmpada de calor até que esteja deambulando e pronto para ser devolvido à sua gaiola. Experimentos com imagens pode ser iniciada após o tempo de incubação desejado (dias a semanas após a injeção viral). </li></ol> 5. Limpeza <ol><li> Lavar a micropipeta com lixívia a 10% e descartá-lo em um recipiente adequado. </li><li> Dispor de recipiente de resíduos da mesma maneira como Seção 1. </li><li> Descarte o papel bancada do laboratório em bin de risco biológico e limpar todas as superfícies e instrumentos que possam ter estado em contacto com o vírus com água sanitária a 10%. </li><li> Vírus não utilizados podem ser congelados novamente, tendo em mente que repetidos de congelamento / descongelamento causar degradação do vírus. </li></ol> 6. Resultados representante <img src="/files/ftp_upload/2140/2140fig1.jpg" alt="Figura 1" /> Figura 1. Transduzidas neurônio após a injeção de double-stranded adeno-associated virus sorotipo 1 (dsAAV S1) carregando proteína verde fluorescente (GFP) sob controle do promotor CMV. O corpo da célula, bem como dendritos proximais e distais são claramente visíveis nas seções fixas fotografada usando um microscópio de epifluorescência. Barra de escala = 100 mm. <img src="/files/ftp_upload/2140/2140fig2.jpg" alt="Figura 2" /> Figura 2. Labeling típico de uma injeção do vírus intracraniana em córtex visual primário usando S1 dsAAV mostrando a extensão da disseminação viral, assim como neurônios rotuladas, glia e processos. Barra de escala = 250 mM. <img src="/files/ftp_upload/2140/2140fig3.jpg" alt="Figura 3" /> Figura 3. Rotulagem de células no hipocampo usando S1 dsAAV. Barra de escala = 250 mM. Estes números são adaptados 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>

Os protocolos experimentais animais e foram aprovados pela Universidade de Rochester University Comissão de Recursos Animais (UCAR), em conformidade com o PHS Política de Assistência Humanitária e Uso de Animais de Laboratório.

Virally mediada entrega do gene tem um grande potencial para o estudo de processos neurológicos e tratamento de distúrbios cerebrais 1,2,3. A grande versatilidade desta técnica também pode ser explorada para fluorescente rótulo células para geração de imagens tanto in vitro como in vivo 4. Aqui demonstramos um procedimento detalhado para a transdução de neurônios e células gliais no córtex visual do rato usando um double-stranded do vírus adeno-associação expressando a proteína verde fluorescente melhorada. Embora esta técnica é relativamente simples, há uma série de detalhes técnicos que precisam ser considerados. Um fator importante é a injeção induzida por dano ao tecido - portanto, é crucial ter muita cautela durante a cirurgia e inserção / remoção da micropipeta. A cirurgia não pode resultar na alteração das estruturas a ser trabalhada. Além disso, depois de carregar o vírus para a micropipeta, muito cuidado deve ser tomado para limpar as bolhas de ar na ponta da pipeta e garantir o volume adequado é injetado. Redução do micropipeta um pouco além da meta Z-coordenar, em seguida, retirá-la na posição adequada pode evitar vírus de transbordar para fora do local da injeção. Ao escolher um tempo de incubação, permitindo que o tempo adequado para a expressão do gene viral deve ser considerada. Isso vai variar de acordo com o vírus utilizado. A resposta imune limitada pode ser obtida pela injeção. Em nossa experiência, essa inflamação é geralmente restrito à faixa de agulha e pode ser evitada por imagem longe da vista de injeção (aproximadamente 50 μms ou mais). Por último, se as seções do cérebro estão a ser montados em lâminas, o uso de um meio de montagem Antifade é sugerido para manter a fluorescência. Injeção intracraniana de vetores virais tem várias vantagens técnicas em relação às técnicas rotulagem outros. Através do uso de coordenadas estereotáxica e volume injetado a modulação, etiqueta fluorescente pode ser precisamente localizado em uma área de interesse. A quantidade de transdução pode ser alterado, ajustando o volume de injeção, a título de vírus ou o tempo de sobrevivência, permitindo a visualização de qualquer grupo de células ou células individuais. Além disso, o uso de vírus diferentes (por exemplo lentivírus, herpes vírus adenovírus,) também pode modular o tempo e quantidade de proteína fluorescente expressa, bem como os tipos de células-alvo. No geral, esta técnica pode fornecer para a rotulagem muito versátil de elementos diferentes do cérebro para a imagem latente.

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		<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

Injeção intracraniana de vetores virais projetadas para expressar uma proteína fluorescente é uma técnica de rotulagem versátil para visualização de subgrupos específicos de células em diferentes regiões do cérebro in vivo e em secções do cérebro. Ao contrário da injeção de corantes fluorescentes, rotulagem viral oferece segmentação de tipos de células individuais e é menos caro e demorado do que estabelecer linhas de ratinhos transgénicos. Nesta técnica, um vetor adeno-associado (AAV) viral é injetado intracraniana usando coordenadas estereotáxica, uma micropipeta e uma bomba automatizada para entrega preciso de AAV para a área desejada com o mínimo dano ao tecido circundante. Parâmetros de injeção pode ser adaptado para experimentos individuais, ajustando a idade do animal em injeção, injeção de localização, volume de injeção, a taxa de injeção, sorotipo AAV eo promotor dirigindo a expressão do gene. Dependendo das condições de escolhido, virally induzida a expressão do transgene pode permitir a visualização de grupos de células, as células individuais ou multa processos celulares, até o nível das espinhas dendríticas. O experimento mostrado aqui representa uma injeção de double-stranded AAV expressando a proteína fluorescente verde para a rotulagem de neurônios e células gliais no córtex visual primário do mouse.

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

Intracranial Injection of Adeno-associated Viral Vectors

Aqui apresentamos a injeção intracraniana de vetores de AAV para a rotulagem fluorescentes de neurônios e células gliais no córtex visual.

Injeção intracraniana da adeno-associado vetores virais

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 visualizações.  University of Rochester. Aqui apresentamos a injeção intracraniana de vetores de AAV para a rotulagem fluorescentes de neurônios e células gliais no córtex visual.

Vídeo: 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.

Segmentação de estruturas cerebrais profundas com Microinjeções para entrega de drogas, vetores virais, ou Transplantes de Células

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

Neurociência

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.

Injection Viral intracerebroventricular do Neonatal do rato do cérebro para a transdução neuronal persistente e 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.

Limbo Abordagem-Subretinal injeção de vetores virais para terapia genética em ratos 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 ...

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 intravascular A injeção de Viral partículas e fluorescentes microesferas no cérebro 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.

Vedação subjugada Adeno-associada do vírus 9 (AAV9) em ratos 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 e dicas para administração intravenosa de vírus Adeno-associado aos ratos e avaliação do sistema nervoso Central transdução

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.

Expressão do Transgene de mediada por vírus adeno-associado em geneticamente definidos neurônios da medula espinhal

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.

Injeção intratecal de vírus Adeno-associado recombinantes em ratos 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.

Produção, purificação e controle de qualidade para vetores de baseados em vírus Adeno-associado

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.