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.

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		<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 &#216; 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 00</td>
			<td>Catgut 
			Markenkirchen</td>
			<td></td>
		</tr>
		<tr>
			<td>syringe, with 30G 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 ddH2O</td>
		</tr>
		<tr>
			<td>10x PBS, pH 7.4</td>
			<td>Carl Roth</td>
			<td>3957.1</td>
			<td>NaCl [1.37 M]</td>
		</tr>
		<tr>
			<td>6781.1</td>
			<td>KCl [27 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>T106.2</td>
			<td>Na2HPO4+2H2O [100 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3904.1</td>
			<td>KH2PO4 [18 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10% Ketamin</td>
			<td>Serum Werk 
			Bernburg</td>
			<td>Urotamin</td>
			<td>mix in a rate 1:1</td>
		</tr>
		<tr>
			<td>2% Xylazin</td>
			<td>Serum Werk 
			Bernburg</td>
			<td>Xylazin</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilium</td>
			<td>Bode Chemie</td>
			<td>Sterillium</td>
			<td>disinfection</td>
		</tr>
		<tr>
			<td>vet ointment</td>
			<td>S&#38;K Pharma 
			GmbH</td>
			<td>Kerato Biciron 5% 
			Augensalbe</td>
			<td>or similar to prevent 
			dryness of eyes</td>
		</tr>
		<tr>
			<td>To-Pro-3 iodide</td>
			<td>Invitrogen</td>
			<td>T3605</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

Research

JoVE Journal

Biology

In Vivo Microinjection and Electroporation of Mouse Testis

27.6K 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

Single Cell Electroporation in vivo within the Intact Developing Brain

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. The distinct advantage of this technique is that experimental manipulations may be performed on individual cells while leaving the surrounding tissue unaltered, thereby distinguishing cell-autonomous effects from those resulting from global treatments. When combined with advanced in vivo imaging techniques, SCE of fluorescent markers permits direct visualization of cellular morphology, cell growth, and intracellular events over timescales ranging from seconds to days. While this technique is used in a variety of in vivo and ex vivo preparations, we have optimized this technique for use in Xenopus laevis tadpoles. In this video article, we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the albino Xenopus tadpole. We also discuss methods to optimize yield, and show examples of live two-photon fluorescence imaging of neurons fluorescently labeled by SCE.

Single-cell electroporation (SCE) is a specialized technique allowing delivery of DNA or other macromolecules into individual cells within intact tissue, including in vivo preparations. Here we detail the procedure for SCE of a fluorescent dye or plasmid DNA into neurons within the intact brain of the Xenopus laevis tadpole.

Eletroporação única célula in vivo dentro do cérebro intacto desenvolvimento

Single-cell electroporation (SCE) is a specialized technique allowing the delivery of DNA or other macromolecules into individual cells within intact ...

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell differentiation, axon guidance, retinotopic organization and synapse formation[1]. One drawback is the inability to efficiently and reliably manipulate gene expression in RGCs in vivo, especially in the otherwise accessible murine visual pathways. Transgenic mice can be used to manipulate gene expression, but this approach is often expensive, time consuming, and can produce unwanted side effects. In chick, in ovo electroporation is used to manipulate gene expression in RGCs for examining retina and RGC development. Although similar electroporation techniques have been developed in neonatal mouse pups[2], adult rats[3], and embryonic murine retinae in vitro[4], none of these strategies allow full characterization of RGC development and axon projections in vivo. To this end, we have developed two applications of electroporation, one in utero and the other ex vivo, to specifically target embryonic murine RGCs[5, 6]. 
With in utero retinal electroporation, we can misexpress or downregulate specific genes in RGCs and follow their axon projections through the visual pathways in vivo, allowing examination of guidance decisions at intermediate targets, such as the optic chiasm, or at target regions, such as the lateral geniculate nucleus. Perturbing gene expression in a subset of RGCs in an otherwise wild-type background facilitates an understanding of gene function throughout the retinal pathway. Additionally, we have developed a companion technique for analyzing RGC axon growth in vitro. We electroporate embryonic heads ex vivo, collect and incubate the whole retina, then prepare explants from these retinae several days later. Retinal explants can be used in a variety of in vitro assays in order to examine the response of electroporated RGC axons to guidance cues or other factors. In sum, this set of techniques enhances our ability to misexpress or downregulate genes in RGCs and should greatly aid studies examining RGC development and axon projections.

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Here we present two techniques for manipulating gene expression in murine retinal ganglion cells (RGCs) by in utero and ex vivo electroporation. These techniques enable one to examine how alterations in gene expression affect RGC development, axon guidance, and functional properties.

No útero e ex eletroporação in vivo para Expressão Gênica em células ganglionares da retina do rato

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell ...

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 ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

In vivo de imagens e tratamentos terapêuticos em um modelo de camundongo ortotópico de câncer de ovário

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1. Apoptosis can be triggered when cells encounter a wide range of cytotoxic stresses. These insults initiate signaling cascades that ultimately cause the release of cytochrome c from the mitochondrial intermembrane space to the cytoplasm2. The release of cytochrome c from mitochondria is a key event that triggers the rapid activation of caspases, the key cellular proteases which ultimately execute cell death3-4.
The pathway of apoptosis is regulated at points upstream and downstream of cytochrome c release from mitochondria5. In order to study the post-mitochondrial regulation of caspase activation, many investigators have turned to direct cytoplasmic microinjection of holocytochrome c (heme-attached) protein into cells6-9. Cytochrome c is normally localized to the mitochondria where attachment of a heme group is necessary to enable it to activate apoptosis10-11. Therefore, to directly activate caspases, it is necessary to inject the holocytochrome c protein instead of its cDNA, because while the expression of cytochrome c from cDNA constructs will result in mitochondrial targeting and heme attachment, it will be sequestered from cytosolic caspases. Thus, the direct cytosolic microinjection of purified heme-attached cytochrome c protein is a useful tool to mimic mitochondrial cytochrome c release and apoptosis without the use of toxic insults which cause cellular and mitochondrial damage.
In this article, we describe a method for the microinjection of cytochrome c protein into cells, using mouse embryonic fibroblasts (MEFs) and primary sympathetic neurons as examples. While this protocol focuses on the injection of cytochrome c for investigations of apoptosis, the techniques shown here can also be easily adapted for microinjection of other proteins of interest.

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

In this protocol, we describe the direct cytoplasmic microinjection of cytochrome c protein into fibroblasts and primary sympathetic neurons. This technique allows for the introduction of cytochrome c protein into the cytoplasm of cells and mimics the release of cytochrome c from mitochondria, which occurs during apoptosis.

Ativação da apoptose por microinjeção citoplasmática de citocromo C</em

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1.  Apoptosis can be triggered when cells encounter a ...

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Avaliação do mouse microinjeção de oócitos, maturação e ploidia

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

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 ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Em Eletroporação vivo de Morpholinos na Retina Zebrafish Adulto

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Na eletroporação in vivo de Morpholinos no Adulto Regeneração Zebrafish Cauda Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...