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

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

in vivo microinjection and electroporation of mouse testis

Esta 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

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

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

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

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

La transfección de células de ratón del ganglio de la retina por In vivo La electroporación

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

Neurociencias

In vivo Electroporation of Developing Mouse Retina

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

In vivo Electroporation of Developing Mouse Retina

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

En electroporación in vivo de la retina del ratón en desarrollo

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

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

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

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

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

Del ratón en el útero de electroporación: Transefection controlada Gene espacio-temporal

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

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

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

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

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

Transferencia de genes a la oreja del ratón en desarrollo por el interior<em&gt; En Vivo</em&gt; La electroporación

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

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

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

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

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

Un método In Vivo para el estudio de integridad de barrera Hematotesticular ratón

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

Biología del Desarrollo

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 ...

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

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

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

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

Modelos de ratón de ingeniería genómica somática utilizando microinyección in vivo y electroporación

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

Investigación sobre el cáncer

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 ...

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

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

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

Novedoso método de plásmido ADN entrega al urotelio de la vejiga de ratón por electroporación

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