Name: in vivo microinjection and electroporation of mouse testis
Uploaded: 2014-08-23T15:00:00.000+00:00
Duration: 8 min 39 s
Description: Watch this Scientific Journal Video about In Vivo Microinjection and Electroporation of Mouse Testis at JoVE.com

Agradecemos Birgit Westernstroeer do Centro de Medicina Reprodutiva e Andrologia da Universidade de Muenster para o ensino da microinjeção testicular. Além disso, somos gratos a Ursula Antkewitz e Petra Reckling para assistência técnica. Agradecemos a Fundação Alemã de Pesquisa (DFG) para apoiar este trabalho (WE2458 / 10-1). &#160;

Todos os experimentos com animais realizados foram aprovados pelo comitê local de ética (Landesamt für Landwirtschaft, Lebensmittelsicherheit und Fischerei, Mecklenburg-Pomerânia Ocidental, Alemanha). 1. plasmídeo Preparação <ol><li> Para a preparação de plasmídeo, utilizar kits de purificação de plasmídeo (ver tabela de materiais) ou métodos semelhantes com tampão de remoção de endotoxinas de forma que as reacções imunitárias do animal podem ser evitadas. Siga as instruções do manual. Empregar ddH2O para diluir a solução de plasmídeo. </li><li> Elimine os restos girando a solução de DNA em velocidade máxima (20.000 xg). Em seguida, recolher o sobrenadante. </li><li> Determinar a concentração de plasmídeo com um espectrofotômetro (ver materiais). </li><li> Ajustar a concentração de plasmídeo com ddH2O a 1-3 ug (ADN) / uL. Nota: as concentrações mais baixas irão reduzir a eficiência de transfecção, enquanto que as concentrações mais elevadas pode causar uma solução de injecção muito viscoso. Além disso, oa eficiência de transfecção depende do tamanho do plasmídeo e tem de ser testado individualmente. </li><li> Antes da injecção, preparar uma mistura de solução com, por exemplo, 40 ul de plasmídeo em conjunto com 5 mL de PBS (10x) e 5 ul Verde Rápido (0,5%). Fast Green é necessária para acompanhar o processo de injeção. De preferência, use parede PCR tubos finos ou Parafilm. Nota: Dependendo do tamanho do testículo, vai ser necessário um volume de 20-50 mL de cada testículo. </li></ol> 2 Preparação de microinjeção Pipeta <ol><li> Use capilares de borosilicato (ver tabela de materiais) para a preparação das pipetas de microinjeção. </li><li> Puxe capilares de vidro com um extrator capilar vertical (veja a tabela de materiais). </li><li> Quebre a ponta capilar com uma pinça por suavemente bandas. A ponta é geralmente de 50-80 fim de diâmetro e cerca de 1 cm de comprimento. Tente evitar dicas que são muito longos como estes não são rígidas o suficiente para penetrar no tecido. </li><li> Enfim, sharpen a ponta na 30 ° a 45 ° de ângulo usando um beveler micropipeta (veja a tabela de materiais). </li><li> Coloque a pipeta de microinjeção com a mistura de solução injectável (ver Preparação de plasmídeo). Aplicar a solução para o fundo do tubo capilar de vidro com uma pequena seringa. Nota: Tente evitar bolhas de ar durante o carregamento, caso contrário estes serão injetados nos túbulos seminíferos, juntamente com a solução de plasmídeo e, portanto, irá reduzir a condutividade, bem como, eventualmente, causar danos aos tecidos. </li></ol> 3 Anestesia e Cirurgia <ol><li> Executar a operação sob condições assépticas, utilizando materiais estéreis (ou seja, seringa, agulha, instrumentos cirúrgicos, etc). Isto irá reduzir a infecção do animal e garantir boas taxas de sobrevivência. </li><li> Use camundongos machos para eletroporação em uma idade de 6-8 semanas. </li><li> Para inicializar o tratamento analgésico pós-cirúrgico, fornecer o mouse com analgésicos adicionado água potável no dia anterior à cirurgia. Este burroUres a analgesia preventiva em caso de hypodipsia postsurgical altamente provável (a ingestão de água diminuída). Para este fim, expor a água potável, com por exemplo uma suspensão de ibuprofeno pediátrico (20 mg / ml). A água para beber medicada também deve ser fornecida em um dia e dois de recuperação na concentração idêntica. Isto significa que uma dose diária de 7,5 mg / kg, a validade de 5 ml / d de água potável e de 25 g de peso corporal durante 8 semanas, com idade C57BL / 6J. Este regime analgésico garante um alívio da dor fundamental e recuperação acelerada junto com elevado bem-estar 26. </li><li> Para a preparação da solução de anestesia no dia da cirurgia de trabalho, misturar 10% cetamina e xilazina a 2% em uma proporção de 1: 1 (ver tabela Materiais). </li><li> Para anestesiar o animal, aplicam-se 0,25 ml / 100 mg de peso corporal por via subcutânea da solução anestésica (cetamina = 0,125 g / kg; Xilazina = 0,025 g / kg). Utilize um local de injecção entre pélvica e nos membros, a fim de evitar danos a órgãos e lesões dos camundongos. Geralmente, são necessários 10 minaté que o mouse está profundamente anestesiados. </li><li> Cubra os olhos do rato com vet pomada para evitar o ressecamento durante a anestesia. </li><li> Teste a parada anestésico, o que é perceptível por uma total falta de resposta. Para esse efeito, basta apertar o dedo do pé do animal. </li><li> Para iniciar a operação, remover o cabelo abdominal com um barbeador elétrico ou similar e desinfectar a área de operação com sterilium ou similar. </li><li> Fazer uma incisão ventral directamente acima das glândulas prepucial no centro da área abdominal. Naquele lugar, primeiro puxe a pele longe e realizar um pequeno corte cutânea transversal de cerca de 8-14 mm. Em seguida, continuar ao longo da camada muscular abdominal. Nota: A lesão deve ser tão pequeno quanto possível para reduzir os efeitos nocivos para o animal. </li><li> Puxe para cima as bolsas de gordura abdominal cuidadosamente para expor o testículo em anexo e coloque-o em uma cortina impermeável preparado antes descartável de papel ao lado do local da incisão (Figura 2A). </li><li> Use binóculos para find o ducto eferente como o alvo da injeção. O eferente ductus é identificada como a junção navio tênue entre o testículo e epidídimo. Ele está localizado ao lado da artéria testicular proeminente. O canal funciona quase em um ângulo de 45 ° para a artéria e visivelmente entra no epidídimo caput. Nota: Dependendo da idade do mouse, o canal é geralmente enterrada no tecido adiposo. </li><li> Use uma pinça fina para limpar o ducto eferente deste tecido adiposo. Depois disso, coloque o ductus lançado em uma tira de papel estéril para assegurar a clara visibilidade do ducto sem prejudicar arredores. </li></ol> 4. microinjeção e Aplicação DNA <ol><li> Ligue o pipeta microinjecção, que é carregado com a solução de plasmídeo para a unidade micromanipulador / injector. </li><li> Inserir a agulha de microinjecção paralelo à conduta eferente com a ponta voltada para a rede testicular (Figura 2B). </li><li> Use uma pinça fina etiras da conduta através da pipeta micro-injecção. Certifique-se de que o capilar é mantido paralelo ao ducto enquanto a puxa para cima. Nota: Este procedimento é mais conveniente do que penetrar no navio, movendo a agulha com a micromanipulador. </li><li> Direcione a agulha cuidadosamente para o testículo e parar assim como penetra na rete testis diretamente abaixo da túnica albugínea (Figura 2C). Nota: No caso de overreaching rete testis, a solução plasmídeo vai vazar para o espaço intersticial. Isso geralmente leva ao fracasso de transfecção dos túbulos seminíferos. </li><li> Para a injecção da solução de plasmídeo utilizar o microinjetor com seguintes configurações: pi: 100 hPa ti,: 0,2 seg, e pc: 0 hPa (Figura 2D). </li><li> Acompanhar todo o processo de injecção por meio da observação do estado de enchimento do testículo com a ajuda da cor verde. Tome cuidado para que o testículo só é cheio até dois terços do seu volume com o plsolução asmid (Figura 2E). Nota: Se o volume de injeção for excedido, o tecido testicular podem ser prejudicados. </li></ol> 5. Eletroporação de Testículo <ol><li> Para habilitar eletroporação eficaz, mergulhe os eletrodos pinça em PBS (1x). Isto assegura condutância adequada. </li><li> Suavemente aperte o testículo entre os eletrodos molhados (Figura 2F). Meça a resistência elétrica com a electroporator. </li><li> Aplique corrente ao testículo. Execute oito pulsos quadrados de 40 V em 4 locais diferentes testículo com uma duração constante de 50 ms e tempo de pulso de 950 ms tempo de intervalo. </li></ol> 6. Wound Encerramento e Pós-cirurgia <ol><li> Quando eletroporação estiver concluída, coloque o testículo de volta ao local original. </li><li> Costure a camada muscular interna com sutura cirúrgica (ver materiais). </li><li> Feche a pele através do emprego de grampos de sutura (ver tabela de materiais). Retire a pele para cima sagacidadeh pinças. Certifique-se de excluir a camada muscular. Coloque os clipes, cada um a uma distância de não mais de 5 mm. Nota: Por causa de sutura cirúrgica pode ser engolido pelo mouse, clipes de sutura são favorecidos para fechar a lesão de pele. </li><li> Permitir que o rato a recuperar da anestesia sobre toalhas de papel estéreis colocados numa gaiola do rato estéril vazio, que é temperado numa almofada quente 37 ° C. A almofada deve assegurar o aquecimento de um meio da gaiola criando um gradiente de calor. Para o animal torna-se possível procurar a área individual, mais confortável, devido à variedade de temperatura na gaiola. Além disso, cubra o mouse com uma folha de papel toalha estéril para que o estresse pode ser ainda mais reduzida. Nota: Como a superfície do corpo do rato é extraordinariamente alta em relação à massa corporal, quando comparados com animais de maior porte, o apoio térmica é de particular importância para a recuperação bem sucedida de roedores. </li><li> Quando o animal tem operado totalmente desperto, transferir it para posterior recuperação para uma nova gaiola do rato completamente equipada. Não coloque o animal de volta para sua gaiola de idade ou para um grupo de camundongos. Certifique-se que a gaiola é limpa e estéril quanto possível para prevenir a infecção da ferida. Acompanhar todo o processo de recuperação. Para transferir o mouse de volta para a unidade de cuidados animal, espere até que ele se recuperou totalmente e parece comportar-se normalmente. Nota: estratégias pós-cirúrgicos per- adicionais e são discutidos por Pritchett-Corning KR et al 6. </li></ol>

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Marten Michaelis, Alexander Sobczak, e Joachim M. Weitzel empregados no Instituto de Biologia Reprodutiva, do Instituto Leibniz de Biologia Animal Farm (FBN), Dummerstorf, Alemanha, declaram que não têm interesses financeiros concorrentes.

A investigação no campo da biologia reprodutiva, em especial na área da fertilidade masculina e, inevitavelmente, a espermatogénese depende de organismos vivos. A fim de examinar a função testicular, sem / sistema de cultura de células in vitro adequada foi estabelecida capaz de reflectir as alterações morfológicas importantes de um diplóide spermatogonium para um 1,2 espermatozóide maduro haplóide. Assim, a geração de animais geneticamente modificados é muitas vezes ...

Espermatogénese mamíferos é considerado um processo sofisticado de células estaminais auto-renovação submetidos sucessivamente a mitose, meiose e a diferenciação, a fim de se desenvolver em espermatozóides maduros haplóide. Estas alterações morfológicas são orquestradas por diferentes tipos de células e, apesar das tentativas profundas, ainda não é possível para imitar esses processos de cultura de células de 1,2. Portanto, as pesquisas sobre a espermatogênese até agora depende de organismos vivos como modelos in vivo. Em geral, os estudos de função gênica são geralmente baseados em animais transgênicos. No entanto, gerar e manter este tipo de modelo animal é demorado, dispendioso e muito elaborado. Isto é atribuído ao processo de criação de comprimento necessária para a geração e manutenção do transgene ao longo de gerações. Além disso, a manipulação genética de todo o organismo pela abordagem transgênicos ou knockout é propenso a causar danos fisiológicos ao direcionamento genes com funções essenciais em várias regiões, como por exemplo, fora do testículo ou sistemicamente. Além disso, alguns métodos de transfecção transiente estão associados com alguns inconvenientes fundamentais. Por exemplo, desvantagens típicas de transferência de genes mediada por vírus são a provocação de possíveis reacções imunológicas e regulamentos de segurança adicionais, ao passo que a lipofecção 3 e 4 bombardeamento com micropartículas pode danificar o tecido e são limitados a uma certa profundidade da célula suficiente para a eficiência de transfecção. Em contraste, electroporação (EP) como outra forma comum de transfecção transiente, parece constituir uma técnica promissora para permitir na transfecção in vivo e na análise de genes in vivo consistentes. Em geral, a EP é referido como um fenómeno dinâmico que depende da tensão transmembranar local com poros consequentemente mediadas dentro nanossegundos. Essas lacunas podem ser mantidos por milésimos de segundo, suficiente to conceder acesso ao DNA, RNA ou pequenas moléculas 5. Quando a tensão aplicada for muito elevada, o carácter geralmente transiente de PE é contrariada, devido à produção de calor e de permeabilização de indução muito abrangente, com consequente dano irreversível da célula 5. Aqui, mostramos que a electroporação é um sistema de transfecção mais eficaz e económica, que é capaz de ser utilizado para a transformação genética do testículo, de modo a elucidar as características de genes in vivo testiculares. Este artigo aborda preparação plasmídeo, microinjeção via o ducto eferente ea eletroporação posterior de rato testículo. Este procedimento pode ser o meio de escolha para atingir a transfecção rápida, específica e eficaz de túbulos seminíferos de testículo do rato in vivo, a fim de investigar os processos de espermatogénese.

<table><tbody><tr><td>Centrifuge</td><td>Sigma</td><td>1-15 PK</td><td></td></tr><tr><td>Spectrophotometer</td><td>Nanodrop</td><td>ND-1000</td><td></td></tr><tr><td>Micropipette puller</td><td>Narishige</td><td>PC-10</td><td>vertical capillary puller</td></tr><tr><td>Microinjection capillaries</td><td>Clark</td><td>GC100-10</td><td>borosilicate standard wall</td></tr><tr><td>Micropipette beveler</td><td>Bachofer</td><td>Typ 462</td><td>rotating disk beveler</td></tr><tr><td>Electroporator</td><td>Nepagene</td><td>CUY 21EDIT</td><td>square wave electroporator</td></tr><tr><td>Tweezer electrodes</td><td>Nepagene</td><td>CUY 650P5</td><td>5 mm &amp;Oslash; disk electrodes</td></tr><tr><td>Stereo microscope</td><td>Zeiss</td><td>Stemi 2000-C</td><td></td></tr><tr><td>Cold light source</td><td>Zeiss</td><td>KL 1500LCD</td><td></td></tr><tr><td>Microinjection pump</td><td>Eppendorf</td><td>Femtojet</td><td></td></tr><tr><td>Micromanipulator</td><td>homemade</td><td></td><td>XYZ cross table</td></tr><tr><td>Surgical instruments</td><td>FST</td><td></td><td>scissors, forceps, needle holder</td></tr><tr><td>Fine forceps</td><td>FST</td><td>Dumont #5, #7</td><td>efferent ducts preparation</td></tr><tr><td>Michel clip applying stapler</td><td>FST</td><td>Michel, 12029-12</td><td></td></tr><tr><td>Michel suture clips</td><td>FST</td><td>Michel, 12040-01</td><td>7.5 x 1.75 mm</td></tr><tr><td>Surgical suture</td><td>Catgut, Markneukirchen, Germany</td><td>Catgut 00</td><td></td></tr><tr><td>Syringe with 30 G needle</td><td>B. Braun</td><td>Omnican 40</td><td>to load micropipette and for anesthesia</td></tr><tr><td>Plasmid isolation kit</td><td>Promega</td><td>Cat. # A2495</td><td>plasmid Midiprep</td></tr><tr><td>Plasmid pEGFP-C1</td><td>Clontech</td><td>Cat. #6084-1</td><td>CMV-promoter + EGFP</td></tr><tr><td>Plasmid pDsRed2-N1</td><td>Clontech</td><td>Cat. #6084-1</td><td>CMV-promoter + DsRed2</td></tr><tr><td>Fast Green dye</td><td>Sigma</td><td>F7258-25G</td><td>for dilution in ddH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>10% Ketamine</td><td>Serum Werk, Bernburg, Germany</td><td>Urotamin</td><td>mix in a rate 1:1 with xylazine</td></tr><tr><td>2% Xylazine</td><td>Serum Werk, Bernburg, Germany</td><td>Xylazin</td><td>mix in a rate 1:1 with ketamine</td></tr><tr><td>Sterilium</td><td>Bode Chemie</td><td>Sterillium</td><td>disinfection</td></tr><tr><td>Vet ointment</td><td>S&amp;amp;K Pharma, GmbH</td><td>Kerato Biciron 5%, Augensalbe</td><td>opthalmic ointment to prevent eye dryness</td></tr><tr><td>To-Pro-3 iodide</td><td>Invitrogen</td><td>T3605</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;10x PBS, pH 7.4&lt;/strong&gt;</td></tr><tr><td>1.37 M NaCl</td><td>Carl Roth</td><td>3957.1</td><td></td></tr><tr><td>Ibuprofen, Dolormin</td><td>Johnson &amp;amp; Johnson Consumer Health Care Germany</td><td>01094902</td><td>analgesic pediatric solution (NSAID) for postsurgery pain relief</td></tr><tr><td>27 mM KCl</td><td>Carl Roth</td><td>6781.1</td><td></td></tr><tr><td>100 mM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Carl Roth</td><td>T106.2</td><td></td></tr><tr><td>18 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Carl Roth</td><td>3904.1</td><td></td></tr></tbody></table>

in vivo microinjection and electroporation of mouse testis

Esta contribuição de vídeo e artigo apresenta uma descrição abrangente de microinjeção e eletroporação de testículo do rato in vivo. Esta técnica de transfecção particular para células testiculares de mouse permite o estudo de processos exclusivos na espermatogênese. O protocolo a seguir concentra-se em transfecção de células de testículo de rato, com as construções de plasmídeos. Especificamente, utilizou-se o vector repórter pEGFP-C1, que expressa a proteína verde fluorescente melhorada (EGFP) e também o vector pDsRed2 N1-expressar a proteína fluorescente vermelha (DsRed2). Ambos os genes repórter foram codificados sob o controlo do promotor de citomegalovírus humano imediato precoce (CMV). Para realizar a transferência de genes em testículos de rato, as construções de plasmídeos repórter são injectados em testículos de ratos vivos. Para esse efeito, o testículo de um animal anestesiado é exposta e o local de microinjecção é preparado. O nosso lugar preferido deinjeção é o ducto eferente, com a rede testicular, em última instância conectada como a rota de transporte anatômica dos espermatozóides entre o testículo e epidídimo. Desta forma, o enchimento dos túbulos seminíferos após microinjecção é excelentemente gerido e controlado, devido à utilização de soluções de ADN coradas. Depois de observar um enchimento suficiente do testículo por sua estrutura tubular de cor, o órgão é electroporado. Isto permite a transferência da solução de ADN para dentro das células dos testículos. Na sequência de 3 dias de incubação, os testículos são removidos e investigada sob o microscópio de fluorescência verde ou vermelho, ilustrando o sucesso da transfecção. Geralmente, este protocolo pode ser utilizado para a entrega de DNA- ou RNA- constrói na vida do mouse testis, a fim de (mais) expressar ou derrubar genes, facilitando na análise da função dos genes vivo. Além disso, é adequado para o estudo de construções repórter ou regula gene putativoelementos Conservador. Assim, as principais vantagens da técnica de eletroporação é rápido desempenho em combinação com baixo esforço, bem como o equipamento técnico moderado e as habilidades necessárias em relação a técnicas alternativas.

A configuração experimental para a realização de micro-injecção e de electroporação de testículo do rato in vivo, uma vez que é usado de acordo com o protocolo ilustrado na Figura 1. Embora seja possível para adquirir as micropipetas de fabrico industrial, que preferido para gerar os nossos próprios pipetas puxando (Figura 1A e) chanfradura (Figura 1B) capilares de vidro para se adequarem às necessidades. O equipamento para a microinjecção e elec...

Watch this Scientific Journal Video about In Vivo Microinjection and Electroporation of Mouse Testis at JoVE.com

In Vivo Microinjection and Electroporation of Mouse Testis

Este artigo descreve a microinjecção e a electroporação de testículo do rato in vivo, como uma técnica para a transfecção de células de testículo de rato para estudar os processos exclusivos da espermatogénese. O protocolo apresentado envolve os passos de preparação de capilar de vidro, através da conduta de microinjecção eferente, e transfecção por electroporação.

In Vivo microinjeção e Eletroporação de testículos de camundongo

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

in-vivo-microinjection-and-electroporation-of-mouse-testis

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

In Vivo Microinjection and Electroporation of Mouse Testis

27.8K visualizações.  Leibniz Institute for Farm Animal Biology (FBN).  Este artigo descreve a microinjecção e a electroporação de testículo do rato in vivo, como uma técnica para a transfecção de células de testículo de rato para estudar os processos exclusivos da espermatogénese. O protocolo apresentado envolve os passos de preparação de capilar de vidro, através da conduta de microinjecção eferente, e transfecção por electroporação. 

Vídeo: In Vivo Microinjection and Electroporation of Mouse Testis

In Utero Intraventricular Injection and Electroporation of E15 Mouse Embryos

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors lining the walls of the lateral ventricle. This technique is now widely used to study the processes involved in corticogenesis by over-expressing or knocking down genes and observing the effects on cellular proliferation, migration, and differentiation. In comparison to traditional knockout strategies, in-utero electroporation provides a rapid means to manipulate a population of cells during a specific temporal window. In this video protocol we outline the experimental methodology for preparing mice for surgery, exposing the uterine horns through laporatomy, injecting DNA into the lateral ventricles of the developing embryo, electroporating DNA into the progenitors lining the lateral wall, and caring for animals post-surgery. Our laboratory uses this protocol for surgeries on E13-E16 mice, however, it is most commonly performed at E15, as shown in this video. 

In Utero injeção intraventricular e eletroporação de embriões de camundongos E15

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors ...

Biologia

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

Transfecção de células de camundongo por Ganglionares da Retina In vivo Eletroporação

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Neurociência

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

Em Eletroporação vivo de Retina do rato em desenvolvimento

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance molecules, local gene transfer or knock- out is required. Gene targeting knock-in or knock-out into local regions is possible to perform with combination with a specific CRE line, which is laborious, costly, and time consuming. Therefore, a simple transfection method, an in utero electroporation technique, which can be performed with short time, will be handy to test the possible function of candidate genes prior to the generation of transgenic animals 1,2. In addition to this, in utero electroporation targets areas of the brain where no specific CRE line exists, and will limit embryonic lethality 3,4. Here, we present a method of in utero electroporation combining two different types of electrodes for simple and convenient gene transfer into target areas of the developing brain. First, a unique holding method of embryos using an optic fiber optic light cable will make small embryos (from E9.5) visible for targeted DNA solution injection into ventricles and needle type electrodes insertion to the targeted brain area 5,6. The patterning of the brain such as cortical area occur at early embryonic stage, therefore, these early electroporation from E9.5 make a big contribution to understand entire area patterning event. Second, the precise shape of a capillary prevents uterine damage by making holes by insertion of the capillary. Furthermore, the precise shape of the needle electrodes are created with tungsten and platinum wire and sharpened using sand paper and insulated with nail polish 7, a method which is described in great detail in this protocol. This unique technique allows transfection of plasmid DNA into restricted areas of the brain and will enable small embryos to be electroporated. This will help to, open a new window for many scientists who are working on cell differentiation, cell migration, axon guidance in very early embryonic stage. Moreover, this technique will allow scientists to transfect plasmid DNA into deep parts of the developing brain such as thalamus and hypothalamus, where not many region-specific CRE lines exist for gain of function (GOF) or loss of function (LOF) analyses.

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

A gene transfer method into the developing mouse brain is described by using a unique surgical method and special shape of electrodes. This unique technique allows transfection of plasmid DNA temporally and spatially, which will aid many neuroscientists in studying brain development.

Mouse No Utero Eletroporação: Transfecção Gênica Controlled Spatiotemporal

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance ...

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

Transferência de genes para a orelha do camundongo Desenvolvimento Inner por<em&gt; In Vivo</em&gt; Eletroporação

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Um método In Vivo para estudar a integridade de barreira de sangue-testículo de rato

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

Biologia do Desenvolvimento

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

Transfecção de DNA de mamíferos usando músculos esqueléticos In Vivo Eletroporação

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

Germline genetically engineered mouse models (G-GEMMs) have provided valuable insight into in vivo gene function in development, homeostasis, and disease. However, the time and cost associated with colony creation and maintenance are high. Recent advances in CRISPR-mediated genome editing have allowed the generation of somatic GEMMs (S-GEMMs) by directly targeting the cell/tissue/organ of interest.
The oviduct, or fallopian tube in humans, is considered the tissue-of-origin of the most common ovarian cancer, high-grade serous ovarian carcinomas (HGSCs). HGSCs initiate in the region of the fallopian tube distal to the uterus, located adjacent to the ovary, but not the proximal fallopian tube. However, traditional mouse models of HGSC target the entire oviduct, and thus do not recapitulate the human condition. We present a method of DNA, RNA, or ribonucleoprotein (RNP) solution microinjection into the oviduct lumen and in vivo electroporation to target mucosal epithelial cells in restricted regions along the oviduct. There are several advantages of this method for cancer modeling, such as 1) high adaptability in targeting the area/tissue/organ and region of electroporation, 2) high flexibility in targeted cell types (cellular pliancy) when used in combination with specific promoters for Cas9 expression, 3) high flexibility in the number of electroporated cells (relatively low frequency), 4) no specific mouse line is required (immunocompetent disease modeling), 5) high flexibility in gene mutation combination, and 6) possibility of tracking electroporated cells when used in combination with a Cre reporter line. Thus, this cost-effective method recapitulates human cancer initiation.

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

This protocol describes microinjection and in vivo electroporation for regionally restricted CRISPR-mediated genome editing in the mouse oviduct.

Modelos de camundongos com engenharia genômica somática usando microinjeção e eletroporação in vivo

Germline genetically engineered mouse models (G-GEMMs) have provided valuable insight into in vivo gene function in development, homeostasis, and disease. ...

Pesquisa do câncer

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies showed that microinjection of germ cells from fertile donor mice into the seminiferous tubules of infertile recipient mice results in donor-derived spermatogenesis and sperm production by the recipient animal2. The use of donor males carrying the bacterial &beta;-galactosidase gene allowed identification of donor-derived spermatogenesis and transmission of the donor haplotype to the offspring by recipient animals1. Surprisingly, after transplantation into the lumen of the seminiferous tubules, transplanted germ cells were able to move from the luminal compartment to the basement membrane where spermatogonia are located3. It is generally accepted that only SSCs are able to colonize the niche and re-establish spermatogenesis in the recipient testis. Therefore, germ cell transplantation provides a functional approach to study the stem cell niche in the testis and to characterize putative spermatogonial stem cells. To date, germ cell transplantation is used to elucidate basic stem cell biology, to produce transgenic animals through genetic manipulation of germ cells prior to transplantation4,5, to study Sertoli cell-germ cell interaction6,7, SSC homing and colonization3,8, as well as SSC self-renewal and differentiation9,10.
Germ cell transplantation is also feasible in large species11. In these, the main applications are preservation of fertility, dissemination of elite genetics in animal populations, and generation of transgenic animals as the study of spermatogenesis and SSC biology with this technique is logistically more difficult and expensive than in rodents. Transplantation of germ cells from large species into the seminiferous tubules of mice results in colonization of donor cells and spermatogonial expansion, but not in their full differentiation presumably due to incompatibility of the recipient somatic cell compartment with the germ cells from phylogenetically distant species12. An alternative approach is transplantation of germ cells from large species together with their surrounding somatic compartment. We first reported in 2002, that small fragments of testis tissue from immature males transplanted under the dorsal skin of immunodeficient mice are able to survive and undergo full development with the production of fertilization competent sperm13. Since then testis tissue xenografting has been shown to be successful in many species and emerged as a valuable alternative to study testis development and spermatogenesis of large animals in mice14.

Protocols for germ cell transplantation and testis tissue xenografting are described. Germ cell transplantation results in donor-derived spermatogenesis in recipient testes and represents a functional reconstitution assay for identification of spermatogonial stem cells (SSCs). Testis tissue xenografting reproduces testis development and spermatogenesis of various donor species in recipient mice.

Transplante de células germinativas e Tecidos Testículo xeno em Ratos

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies ...

Novel Method of Plasmid DNA Delivery to Mouse Bladder Urothelium by Electroporation

Genetically engineered mouse models (GEMMs) are extremely valuable in revealing novel biological insights into the initiation and progression mechanisms of human diseases such as cancer. Transgenic and conditional knockout mice have been frequently used for gene overexpression or ablation in specific tissues or cell types in vivo. However, generating germline mouse models can be time-consuming and costly. Recent advancements in gene editing technologies and the feasibility of delivering DNA plasmids by viral infection have enabled rapid generation of non-germline autochthonous mouse cancer models for several organs. The bladder is an organ that has been difficult for viral vectors to access, due to the presence of a glycosaminoglycan layer covering the urothelium. Here, we describe a novel method developed in lab for efficient delivery of DNA plasmids into the mouse bladder urothelium in vivo. Through intravesical instillation of pCAG-GFP DNA plasmid and electroporation of surgically exposed bladder, we show that the DNA plasmid can be delivered specifically into the bladder urothelial cells for transient expression. Our method provides a fast and convenient way for overexpression and knockdown of genes in the mouse bladder, and can be applied to building GEMMs of bladder cancer and other urological diseases.

We describe a novel method for the delivery of DNA plasmid into the urothelial cells of mouse bladder in vivo through urethra catheterization and electroporation. It offers a fast and convenient way for generating autochthonous mouse models of bladder diseases.

Novo método de entrega de DNA do plasmídeo para Mouse bexiga Urotélio por eletroporação

Genetically engineered mouse models (GEMMs) are extremely valuable in revealing novel biological insights into the initiation and progression mechanisms ...