Damos las gracias a Birgit Westernstroeer del Centro de Medicina Reproductiva y Andrología de la Universidad de Münster para la enseñanza de la microinyección testicular. Además, estamos agradecidos a Ursula Antkewitz y Petra Reckling para la asistencia técnica. Agradecemos a la Fundación Alemana de Investigación (DFG) para apoyar este trabajo (WE2458 / 10-1). &#160;

Todos los experimentos realizados en animales han sido aprobados por el comité de ética local (Landesamt für Landwirtschaft, Lebensmittelsicherheit und Fischerei, Mecklenburg-Vorpommern, Alemania). 1. preparación de plásmido <ol><li> Para la preparación de plásmido, utilizar kits de purificación de plásmido (ver tabla Materiales) o métodos similares con tampón de eliminación de endotoxina de manera que las reacciones inmunes del animal pueden ser evitados. Siga las instrucciones del manual. Emplear ddH2O para diluir la solución de plásmido. </li><li> Eliminar los residuos haciendo girar la solución de ADN a la velocidad máxima (20.000 xg). Luego recoger el sobrenadante. </li><li> Determinar la concentración de plásmido con un espectrofotómetro (ver materiales). </li><li> Ajustar la concentración de plásmido con ddH2O a 1.3 mg (ADN) / l. Nota: Las concentraciones más bajas reducen la eficiencia de la transfección, mientras que las concentraciones más altas pueden causar una solución de inyección demasiado viscoso. Además, ella eficacia de transfección depende del tamaño del plásmido y tiene que ser ensayada por separado. </li><li> Antes de la inyección, preparar una mezcla de solución con, por ejemplo 40 l plásmido junto con 5 l de PBS (10x) y 5 l Verde Rápido (0,5%). Fast Green es necesaria para el seguimiento del proceso de inyección. Preferiblemente, utilice pared PCR tubos delgados o Parafilm. Nota: Dependiendo del tamaño de los testículos, se necesitará un volumen de 20-50 l para cada testículo. </li></ol> 2 Preparación de la pipeta de microinyección <ol><li> Utilice los capilares de borosilicato (ver tabla Materiales) para la preparación de las pipetas de microinyección. </li><li> Tire de capilares de vidrio con un extractor capilar vertical (ver tabla Materiales). </li><li> Rompa la punta capilar con fórceps por suavemente bandas. La punta es usualmente 50-80 micras de diámetro y aproximadamente 1 cm de largo. Trate de evitar las puntas que son demasiado largos ya que estos no son lo suficientemente rígido como para penetrar en el tejido. </li><li> Por fin, sharpen la punta en unos 30 ° a 45 ° de ángulo usando un biselador micropipeta (ver tabla Materiales). </li><li> Cargar la pipeta de microinyección con la mezcla de solución inyectable (ver Preparación de plásmido). Aplicar la solución en la parte posterior del capilar de vidrio con una pequeña jeringa. Nota: Trate de evitar las burbujas de aire durante la carga, de lo contrario estos serán inyectados en los túbulos seminíferos, junto con la solución de plásmido y por lo tanto reducirá la conductividad, así como posiblemente causar daño a los tejidos. </li></ol> 3. Anestesia y Cirugía <ol><li> Realice la operación en condiciones asépticas utilizando material estéril (es decir, jeringa, aguja, instrumentos quirúrgicos, etc). Esto reducirá la infección del animal y asegurar buenas tasas de supervivencia. </li><li> Utilice ratones machos para la electroporación a una edad de 6-8 semanas. </li><li> Para inicializar tratamiento analgésico postquirúrgico, proporciona el ratón con analgésicos añadió agua potable en el día antes de la cirugía. Este culoUres una analgesia preventiva en caso de hipodipsia postquirúrgica altamente probable (consumo de agua disminuyó). Para este fin, exponer el agua potable con, por ejemplo una suspensión de ibuprofeno pediátrico (20 mg / ml). El agua de bebida medicada también debe ser suministrada en el día uno y dos de la recuperación en la concentración idéntica. Esto significa una dosis diaria de 7,5 mg / kg, válido para 5 ml / d de agua potable y el peso corporal de 25 g durante 8 semanas C57BL / 6J años de edad. Este régimen analgésico garantiza un alivio del dolor y la recuperación acelerada fundamental junto con la alta bienestar 26. </li><li> Para la preparación de la anestesia solución en el día de la cirugía de trabajo, mezclar 10% ketamina y 2% xylazin en una proporción de 1: 1 (ver tabla Materiales). </li><li> Para anestesiar al animal, se aplican 0,25 ml / 100 mg de peso corporal por vía subcutánea de solución anestésica (ketamina = 0,125 g / kg; xylazin = 0,025 g / kg). Usar un sitio de inyección entre la pelvis y extremidad a fin de evitar daños en los órganos y lesiones de los ratones. Por lo general, se necesitan 10 minhasta que el ratón está profundamente anestesiados. </li><li> Cubra los ojos de ratones con ungüento veterinario para prevenir la sequedad durante la anestesia. </li><li> Pruebe profunda arresto anestésico, que es notable por la falta total de respuesta. Para ello, simplemente pellizcar el dedo del pie del animal. </li><li> Para iniciar la cirugía, eliminar el vello abdominal con una máquina de afeitar eléctrica o similar y desinfectar el área de operación con sterilium o similar. </li><li> Hacer una incisión ventral directamente encima de las glándulas prepuciales en el centro de la zona abdominal. En ese lugar, primero tire de la piel lejos y llevar a cabo una pequeña incisión cutánea transversal de aproximadamente 8-14 mm. Luego, continuar por la capa muscular abdominal. Nota: La lesión debe ser tan pequeña como sea posible para reducir el daño al animal. </li><li> Tire hacia arriba de las almohadillas de grasa abdominales cuidadosamente para exponer el testículo adjunto y colocarlo en un impermeable cortina de papel desechable antes preparado justo al lado del lugar de la incisión (Figura 2A). </li><li> Use binoculares a find el conducto eferente como el objetivo de la inyección. El ductus eferente se identifica como la unión entre el recipiente bien testículo y el epidídimo. Se encuentra junto a la destacada arteria testicular. El conducto se extiende casi en un ángulo de 45 ° a la arteria y visiblemente entra en el epididymidis caput. Nota: Dependiendo de la edad del ratón, el ductus se suele enterrado en el tejido graso. </li><li> Utilice unas pinzas finas para despejar el conducto eferente de este tejido graso. Después de eso, coloque el ductus liberado en una tira de papel estéril para asegurar la clara visibilidad de la vía sin perjudicar entorno. </li></ol> 4. La microinyección y la Solicitud de ADN <ol><li> Conectar la pipeta de microinyección, que se carga con la solución de plásmido a la unidad de micromanipulador / inyector. </li><li> Coloque la aguja de microinyección paralelo al conducto eferente con la punta apuntando hacia la rete testis (Figura 2B). </li><li> Use unas pinzas finas ypelar el conducto sobre la pipeta de microinyección. Asegúrese de que el capilar se mantiene paralelo al conducto mientras tira de la vuelta. Nota: Este procedimiento es más conveniente que penetrar en el recipiente moviendo la aguja con el micromanipulador. </li><li> Dirija la aguja con cuidado hacia el testículo y detener al igual que penetra en la red testicular directamente debajo de la túnica albugínea (Figura 2C). Nota: En el caso de extralimitación de la rete testis, la solución de plásmido se escapará hacia el espacio intersticial. Esto conduce comúnmente al fracaso de la transfección de los túbulos seminíferos. </li><li> Para la inyección de la solución de plásmido utilizar el microinyector con los siguientes ajustes: pi: 100 hPa ti,: 0,2 seg, y pc: 0 hPa (Figura 2D). </li><li> Supervisar todo el proceso de inyección mediante la observación de los testículos estado de llenado con la ayuda de el color verde. Tenga cuidado de que el testículo sólo se llena hasta 2/3 de su volumen con la plsolución asmid (Figura 2E). Nota: Si se excede el volumen de inyección, el tejido testicular podría ser dañado. </li></ol> 5. electroporación de Testículo <ol><li> Para activar la electroporación efectiva, remoje los electrodos de pinza en PBS (1x). Esto asegura la conductancia adecuado. </li><li> Suavemente apriete el testículo entre los electrodos húmedos (Figura 2F). Medir la resistencia eléctrica con el electroporador. </li><li> Aplicar corriente a los testículos. Realizar ocho pulsos cuadrados de 40 V a los 4 sitios testículo diferentes con una duración constante de tiempo de pulso 50 ms y 950 ms tiempo de intervalo. </li></ol> 6. cierre de la herida y después de la cirugía <ol><li> Cuando se termina la electroporación, colocar el testículo de nuevo en la ubicación original. </li><li> Coser la capa muscular interna con sutura quirúrgica (ver materiales). </li><li> Cierre de la piel mediante el empleo de clips de sutura (ver tabla Materiales). Tire de la piel encima de ingenioh pinzas. Asegúrese de excluir a la capa muscular. Coloque los clips, cada uno a una distancia de no más de 5 mm. Nota: Debido a que la sutura quirúrgica puede ser tragada por el ratón, clips de sutura son favorecidos para el cierre de la lesión cutánea. </li><li> Deje que el ratón se recupere de la anestesia en las toallas de papel estéril colocados en una jaula del ratón vacío estéril, que está templado sobre una almohadilla caliente 37 ° C. La almohadilla debe garantizar el calentamiento de una mitad de la jaula de la creación de un gradiente de calor. Para el animal esto hace posible la búsqueda de la zona individual, más cómodo debido a la variedad de temperatura en la jaula. Además, el ratón cubrir con una hoja de toalla de papel estéril de modo que el estrés puede reducirse aún más. Nota: Dado que la superficie corporal del ratón es extraordinariamente alta en relación a la masa corporal, en comparación con los animales de mayor tamaño, el apoyo térmica es de particular importancia para la recuperación exitosa de los roedores. </li><li> Cuando el animal operado ha despertado completamente, transferir it para su posterior recuperación a una nueva jaula de ratón completamente equipada. No ponga el animal de nuevo a su antigua jaula o en un grupo de ratones. Asegúrese de que la jaula es tan limpio y estéril como sea posible para evitar la infección de la herida. Controle todo el proceso de recuperación. Para transferir el ratón de nuevo a las instalaciones de cuidado animal, espere hasta que se haya recuperado por completo y parece comportarse normalmente. Nota: persona adicional y estrategias post-quirúrgicas se discuten por Pritchett-Corning KR et al 6. </li></ol>

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Marten Michaelis, Alexander Sobczak, y Joachim M. Weitzel empleadas en el Instituto de Biología de la Reproducción del Instituto Leibniz de Biología Animal Farm (FBN), Dummerstorf, Alemania, declarar que no tienen intereses financieros en competencia.

La investigación en el campo de la biología reproductiva, particularmente en el área de la fertilidad masculina y la espermatogénesis, inevitablemente, se basa en los organismos vivos. Con el fin de examinar la función testicular, sin cultivo celular adecuado / sistema in vitro se ha establecido capaz de reflejar todas las cruciales cambios morfológicos de una espermatogonia diploide a un 1,2 espermatozoide maduro haploide. Por lo tanto, la generación de animales modificados g...

La espermatogénesis en mamíferos se considera que es un proceso sofisticado de células madre auto-renovación sometidos sucesivamente la mitosis, la meiosis y diferenciación con el fin de desarrollar en espermatozoides maduros haploide. Estos cambios morfológicos son orquestados por diferentes tipos de células, ya pesar de profundos intentos, todavía es imposible de imitar estos procesos en cultivo celular 1,2. Por lo tanto, la investigación sobre la espermatogénesis hasta ahora se basa en organismos vivos como modelos in vivo. En general, los estudios de la función génica se basan generalmente en animales transgénicos. Sin embargo, la generación y el mantenimiento de este tipo de modelo animal consume tiempo, costo intensiva y bastante elaborada. Esto se atribuye al proceso de mejora genética a largo requerido para generar y mantener el transgén largo de las generaciones. Además, la manipulación genética de todo el organismo por el enfoque transgénico o knockout es propenso a causar alteraciones fisiológicas cuando la orientación genes con funciones esenciales en múltiples regiones, por ejemplo, fuera de los testículos o sistémica. Además, algunos métodos de transfección transitoria se asocian con algunas desventajas cruciales. Por ejemplo, inconvenientes típicos de la transferencia de genes mediada por virus son la posible provocación de inmunorreacciones y reglamentos de seguridad adicionales, mientras que la lipofección y bombardeo de micropartículas 3 4 podría dañar el tejido y están limitadas a una cierta profundidad suficiente de células para la eficiencia de transfección. En contraste, la electroporación (EP) como otra forma común de transfección transitoria, parece constituir una técnica prometedora para permitir la transfección in vivo y análisis in vivo de genes consistente en. En general, EP se conoce como un fenómeno dinámico que depende de la tensión transmembrana local con poros en consecuencia mediadas dentro de nanosegundos. Estas lagunas se pueden mantener durante milisegundos, suficiente to conceder acceso a ADN, ARN o moléculas pequeñas 5. Cuando el voltaje aplicado es demasiado alto, el carácter generalmente transitoria de EP se contrarresta debido a la producción de calor y la inducción de permeabilización demasiado amplia con el consiguiente daño irreversible de la célula 5. Aquí, se muestra que la electroporación es un sistema de transfección eficaz y económico que es capaz de ser utilizado para la transformación genética testículo con el fin de dilucidar las características testiculares de genes in vivo. En este artículo se aborda la preparación de plásmido, microinyección por el conducto eferente y la posterior electroporación de testículos de ratón. Este procedimiento puede ser el medio de elección para lograr la transfección rápido, específico y eficaz de los túbulos seminíferos de los testículos de ratón in vivo con el fin de investigar los procesos de la espermatogénesis.

<table border="0" cellpadding="0" cellspacing="3">
	<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 &#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 contribución de video y artículo da una descripción completa de microinyección y electroporación de testículo de ratón in vivo. Esta técnica particular para la transfección de células de ratón testiculares permite el estudio de los procesos únicos en la espermatogénesis. El siguiente protocolo se centra en la transfección de células testiculares de ratón con construcciones de plásmidos. Específicamente, se utilizó el vector reportero pEGFP-C1, que expresa mayor proteína verde fluorescente (EGFP) y también el vector pDsRed2-N1 que expresa la proteína fluorescente de color rojo (DsRed2). Ambos genes indicadores codificados estaban bajo el control del promotor de citomegalovirus humano temprano inmediato (CMV). Para llevar a cabo la transferencia génica en los testículos de ratón, los constructos de plásmido reportero se inyectan en los testículos de ratones vivos. Para ello, el testículo de un animal anestesiado está expuesto y el sitio de microinyección se prepara. Nuestro lugar preferido deinyección es el conducto eferente, con la rete testis en última instancia conectado como la ruta de transporte anatómica de los espermatozoides entre el testículo y el epidídimo. De esta manera, el llenado de los túbulos seminíferos después de la microinyección se gestiona de manera excelente y controlada debido a la utilización de soluciones de ADN teñidos. Después de observar un relleno suficiente de los testículos por su estructura túbulo coloreado, el órgano se electroporación. Esto permite la transferencia de la solución de ADN en las células testiculares. Después de 3 días de incubación, se retira el testículo y se investigó bajo el microscopio para fluorescencia verde o roja, que ilustra el éxito de la transfección. En general, este protocolo puede ser empleado para la entrega de ADN o ARN construye en la vida testículo del ratón con el fin de (sobre) expresa o derribar los genes, lo que facilita el análisis de la función génica in vivo. Además, es adecuado para el estudio de construcciones informadoras o putativo Regula genelementos tory. Por lo tanto, las principales ventajas de la técnica de electroporación son un rendimiento rápido en combinación con bajo esfuerzo, así como el equipo técnico moderado y habilidades requeridas en comparación con las técnicas alternativas.

La configuración experimental para realizar la microinyección y electroporación de testículos de ratón in vivo, ya que se utiliza de acuerdo con el protocolo se ilustra en la Figura 1. Aunque es posible adquirir micropipetas fabricados industrialmente, preferimos generar nuestras propias pipetas tirando (Figura 1A ) y biselado (Figura 1B) capilares de vidrio para que satisfizo nuestras necesidades. El equipo para la microinyección y electroporación se il...

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 artículo describe la microinyección y electroporación de testículo de ratón in vivo como una técnica de transfección de células de ratón testiculares para estudiar procesos únicos de la espermatogénesis. El protocolo presentado implica etapas de preparación capilar de vidrio, microinyección a través del conducto eferente, y transfección por electroporación.

En Vivo microinyección y electroporación del mouse Testis

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

27.8K vistas.  Leibniz Institute for Farm Animal Biology (FBN).  Este artículo describe la microinyección y electroporación de testículo de ratón in vivo como una técnica de transfección de células de ratón testiculares para estudiar procesos únicos de la espermatogénesis. El protocolo presentado implica etapas de preparación capilar de vidrio, microinyección a través del conducto eferente, y transfección por electroporación. 

Video: 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. 

En la inyección intraventricular Utero y electroporación de embriones de ratón E15

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

Investigación

Biología

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.

La electroporación in vivo sola célula en el cerebro intacto desarrollo

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.

La electroporación in vivo en el útero y el ex de la expresión génica en células de ratón ganglionares retinianas

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.

La transfección de ADN de músculos esqueléticos de mamíferos con En Vivo La electroporación

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

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

Cultura primaria y electroporación plásmido del órgano de Corti murino.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

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.

En vivo de imágenes y tratamientos terapéuticos en un modelo de ratón ortotópico de cáncer de ovario

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.

La activación de la apoptosis mediante microinyección citoplasmática del 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.

La microinyección de ovocitos de ratón, Maduración y Evaluación de ploidía

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.

Germen de Trasplante de Células y Tejidos Testículos xenotrasplante en ratones

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.

En electroporación in vivo de Morpholinos en la retina de pez cebra adultos

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