Los autores agradecen a Sharmin Hossain para el crecimiento de películas de lapso de tiempo la captura de inmaduros neurona y Dunfield Derek para las imágenes de microscopía electrónica de SCE consejos micromicropipette.

Instalación del equipo: <ol><li> Los equipos eléctricos requeridos para esta técnica son un estimulador, un osciloscopio y un soporte para micropipeta equipado con un hilo de plata que se inserta en la micropipeta. Por lo general el uso de un axón instrumentos Axoporator 800A estimulador, sin embargo otros estimuladores comunes, tales como los instrumentos de hierba SD9 estimuladores también son adecuados para una sola célula de electroporación. El estimulador se conecta a la headstage, que conecta con el electrodo de hilo de plata que se coloca dentro de la micropipeta de vidrio que contiene una solución interna llevar a cabo. El otro cable del estimulador se conecta a la entrada del osciloscopio, que también recibe información desde la base externa de renacuajo. La toma de tierra externa renacuajo es más que un hilo de plata que permanece en contacto con el renacuajo electrotonic durante la electroporación. El circuito se completa una vez que la micropipeta se pone en contacto con el renacuajo. </li><li> Electroporación sola célula requiere un microscopio con una buena distancia de trabajo (por lo menos 5 cm), ya que esto da margen de maniobra y permite que la micropipeta para ser llevado en un ángulo de unos 30-45 grados. </li></ol> Fabricación de pipetas: Preparación de pipetas adecuada es un paso crítico en este protocolo. La dificultad radica en el equilibrio de una punta estrecha (menos de 1μm de diámetro) con uno que no se rompa fácilmente al perforar el tejido. También hemos encontrado que las puntas con un ángulo de inclinación de más de 10 grados es preferible. <ol><li> Prepare una micropipeta de vidrio tirado con un diámetro de la punta de menos de 1μm, y un ángulo de inclinación de la punta superior a 10 grados. <ol><li> Vidrio de borosilicato con un filamento interno con un diámetro exterior de 1,5 mm de diámetro interior de 0,75 mm y es muy adecuado para este propósito. <ol><li> El vidrio relativamente gruesa proporciona un grado de rigidez de la punta. </li><li> El filamento interno es importante en la elaboración de nuevo lleno de líquido hacia la punta de la micropipeta. </li></ol></li></ol></li><li> Fabricación de consejos adecuados a menudo se requieren varias modificaciones sobre la base de los resultados observados. Diferentes tejidos puede requerir una punta con una forma ligeramente diferente o diámetro de la punta. </li></ol> Una sola célula de electroporación protocolo: Para los nuevos usuarios, es recomendable comenzar con los tintes fluorescentes dextrano, ya que, a diferencia de las proteínas codificadas genéticamente fluorescente, que se puede observar inmediatamente en epifluorescencia. El uso de colorantes fluorescentes permite al usuario ver directamente la ubicación de la punta de la micropipeta dentro del tejido, y sobre la electroporación ofrece información instantánea sobre si el equipo ha sido configurado correctamente y si la punta de la micropipeta es apropiado. <ol><li> Rellene micropipeta con una solución de compuesto que se electroporated ya sea mediante el uso de una jeringa de carga o de back-llenado. <ol><li> Si se utiliza el ADN plásmido, que las muestras están libres de endotoxinas, y en una concentración entre 1-2μg/ml, preparado en agua. </li><li> La concentración de los tintes fluorescentes dextrano debe ser determinada empíricamente. A menudo se utilizan tintes fluorescentes diluido en agua a aproximadamente 300μM. </li><li> Por lo general, los volúmenes de 0,5-1μl son suficientes. </li><li> Antes de cargar, en pocas palabras centrífuga soluciones a alta velocidad para reducir la cantidad de restos de partículas arrastrados a la micropipeta, lo que puede conducir a la obstrucción. </li><li> Las burbujas de aire dentro de la punta a menudo afectan el flujo de corriente y deben ser desalojados por la micropipeta agitando vigorosamente antes de montar en el headstage. </li></ol></li><li> Inserte micropipeta en el soporte o headstage montado en un micromanipulador de 3 ejes. Asegúrese de que el electrodo de alambre de plata se inserta en micropipeta, y está en contacto con electrotonic la solución interna. </li><li> Anestesiar a los renacuajos por inmersión en un 0,02% MS-222 (3-amino-éster etílico del ácido), preparado en el medio de la crianza de renacuajos (solución de Steinberg, pH 7,4). <ol><li> Normalmente se usan etapa 44-48 albino renacuajos Xenopus laevis en nuestros experimentos. </li><li> Los renacuajos son por lo general completamente anestesiado después de aproximadamente 5 minutos. </li><li> Varios renacuajos se puede anestesiar al mismo tiempo para reducir el tiempo de espera, sin embargo, evitar la exposición a MS-222 para períodos más allá de una hora. </li></ol></li><li> La transferencia de un renacuajo anestesiados solo a la cámara de electroporación utilizando una micropipeta de transferencia de plástico. <ol><li> La cámara de electroporación se puede hacer fácilmente en casa por tallar una cavidad en forma de renacuajo en una pequeña Sylgard ® bloque de silicona, y la inserción de un cable de tierra de plata en la cavidad de tal manera que se pondrá en contacto con electrotonic el renacuajo en la cámara. </li></ol></li><li> Lado la posición de renacuajo dorsal usando un pincel humedecido. </li><li> Baje la micropipeta hasta que esté en contacto con la piel, justo al lado de la región de interés. <ol><li> Orientación regiones con cuerpos celulares densamente aumentará en gran medida las posibilidades de éxito de la electroporación.Por lo general el objetivo de techo óptico renacuajo. </li><li> Traiga la micropipeta en un ángulo de 30 a 45 grados. Menos profundas que las trayectorias de perforación de la piel más difícil, mientras que los ángulos más pronunciada a menudo obstruyen una de vista. </li></ol></li><li> La reducción continua de la micropipeta inicialmente hoyuelo en la piel, para después pasar por dentro del tejido subyacente. <ol><li> Las células superficiales son los preferidos para imágenes in vivo, y puede ser objetivo de asegurar que la punta de la micropipeta se reduce poco a poco en formación de hoyuelos en la piel. </li></ol></li><li> Uno debe vigilar constantemente la resistencia micropipeta una vez colocado en el tejido. Si se utiliza un Axoporator 800A (Axon Instruments), las resistencias de aproximadamente 10 40MΩ son óptimas. La resistencia es un buen indicador de si el diámetro de la punta es apropiada para una sola célula de electroporación. <ol><li> Altas resistencias micropipeta (&gt; 100MΩ) son indicativos de consejos obstruido o micropipetas con puntas que son demasiado estrechas. </li><li> Bajas resistencias micropipeta (&lt;10MΩ) son indicativos de un diámetro de la punta que es demasiado grande, a menudo el resultado de una punta rota. </li><li> Si se utiliza un estimulador que no proporciona una medida de resistencia directa, la resistencia micropipeta indirectamente se puede medir observando la amplitud de los pulsos de tensión medida por el osciloscopio (véase más adelante). </li></ol></li><li> Aplicar tren de tensión. <ol><li> De ADN plásmido, trenes de pulsos encontrado para ser más eficaces consisten en negativo pulsos de onda cuadrada de 1 ms de duración, entregado a 300 Hz con una duración de tren de pulsos de 500 ms. <ol><li> La tensión del pulso se determina empíricamente de tal manera que la amplitud de los pulsos medido en el osciloscopio está entre 0,75 y 1.5μA. </li></ol></li><li> Para lámparas fluorescentes tintes dextrano, positivos impulsos de onda cuadrada de la duración de 300μs se entregan con una frecuencia de 300 Hz con una duración de tren de pulsos de 10 ms. <ol><li> Al igual que con el protocolo de ADN, la tensión del pulso se determina empíricamente de tal manera que la amplitud de los impulsos medidos en el osciloscopio es de entre 0,75 y 1.5μA. </li><li> Ajustes adicionales a los parámetros del estímulo puede hacerse con base en la observación de los resultados de la electroporación en epifluorescencia. <ol><li> Si las células electroporated aparecer tenue, los parámetros del pulso debe aumentar gradualmente hasta que las células aparecen más brillantes. </li><li> Por otro lado, si los grupos de células están etiquetados, los parámetros del pulso debe ser reducido hasta que las células individuales están etiquetados. </li></ol></li></ol></li></ol></li><li> Retraer micropipeta, vuelva a insertar en un sitio diferente dentro del tejido, y aplicar tren de tensión. <ol><li> Para mejorar el rendimiento, por lo general electroporar varios sitios con cada hemisferio del renacuajo Tectum óptico con un espacio suficiente para asegurar que las células marcadas no se superpongan. </li></ol></li><li> Transferir el renacuajo electroporated en un recipiente con una solución de la crianza de Steinberg. <ol><li> Renacuajos rápido se recuperará de la anestesia en pocos minutos. </li></ol></li><li> La micropipeta mismo puede ser utilizado para numerosos renacuajos. Es importante observar constantemente la amplitud de los pulsos observados en el osciloscopio. <ol><li> Si la amplitud del pulso sigue siendo baja después de aumentar significativamente la intensidad del pulso este es probablemente el resultado de la punta está obstruida, a menudo se ve cuando electroporating muchos renacuajos en una sola sesión. <ol><li> Un método para desalojar a una obstrucción punta de menores es la aplicación de un pulso positivo solo después de insertar la micropipeta en el tejido. </li><li> Si la obstrucción no puede ser desplazado, o la obstrucción a menudo se repite, reemplace la micropipeta. </li></ol></li><li> Si la amplitud del pulso sigue siendo muy grande después de disminuir significativamente los parámetros de pulso, esto es una señal de que la punta se ha roto y necesita ser reemplazado. </li></ol></li></ol> La detección de células con éxito electroporated: <ol><li> Después de la electroporación, los renacuajos son examinados para detectar células marcadas con un microscopio de epifluorescencia vertical. </li><li> En el caso de los tintes fluorescentes, los renacuajos pueden ser examinados después de tan poco como 30 minutos después de la electroporación. <ol><li> Hemos encontrado, sin embargo, que los intervalos más largos están asociados con menores niveles de fluorescencia de fondo. </li></ol></li><li> Para genéticamente codificados proteínas fluorescentes, la detección se realiza normalmente al menos 12 horas después de la electroporación. <ol><li> Las células que expresan proteínas fluorescentes seguirá cada vez más brillante con el tiempo, por lo tanto, las células oscuras pueden ser re-examinados después de un intervalo adicional de 12 a 24 horas. </li></ol></li><li> Los renacuajos son anestesiados como se describió anteriormente, en una cámara Sylgard ® y cubreobjetos. Renacuajos individuales se examinan con epifluorescencia. <ol><li> Tenga en cuenta que la exposición excesiva a la luz de epifluorescencia dará lugar a la fototoxicidad que pueden matar a la célula marcada. Por lo tanto, minimizar la cantidad de tiempo que las células están expuestas a la luz fluorescente. </li></ol></li><li> Volver renacuajos a un recipiente con una solución de la crianza de Steinberg. </li></ol>

<ol>
	<li>Bestman, J. E., Ewald, R. C., Chiu, S. L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+single-cell+electroporation+for+transfer+of+DNA+and+macromolecules.">In vivo single-cell electroporation for transfer of DNA and macromolecules.</a> Nature Protocols. 1, 1267-1272 (2006).</li><li>Dunfield, D., Haas, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+electroporation.">Single cell electroporation.</a> Encyclopedia of Neuroscience (4th Ed.). , .</li><li>Haas, K., Jensen, K., Sin, W. C., Foa, L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+electroporation+in+Xenopus+tadpoles+in+vivo--from+single+cells+to+the+entire+brain.">Targeted electroporation in Xenopus tadpoles in vivo--from single cells to the entire brain.</a> Differentiation. 70, 148-154 (2002).</li><li>Haas, K., Sin, W. C., Javaherian, A., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+electroporation+for+gene+transfer+in+vivo.">Single-cell electroporation for gene transfer in vivo.</a> Neuron. 29, 583-591 (2001).</li></ol>

Una sola célula de electroporación (SCE) es una poderosa herramienta para determinar la función del gen y la realización de la manipulación genética dirigida. La transparencia de los renacuajos albinos y la accesibilidad del cerebro hacen que este sistema modelo ideal para la visualización de crecimiento neuronal y eventos intracelulares en un organismo vivo. SCE permite la visualización del crecimiento de una sola neurona, y para llevar a cabo por células autónomas manipulaciones dentro de un cerebro de otra manera inalterada. Si bien este artículo de vídeo muestra el procedimiento de una sola célula de electroporación en los renacuajos Xenopus laevis, esta técnica ha sido utilizada en otros organismos, y también se ha utilizado en rodajas de hipocampo y cultivos disociados de la célula.

single cell electroporation in vivo within the intact developing brain

Una sola célula de electroporación (SCE) es una técnica especializada que permite la entrega de ADN u otras macromoléculas en las células individuales en el tejido intacto, incluso en los preparativos vivo. La ventaja de esta técnica es que la manipulación experimental se puede realizar en las células individuales, dejando el tejido circundante sin alterar, por lo tanto distinguir células autónomas efectos de los resultantes de los tratamientos globales. Cuando se combina con avanzadas técnicas de imagen en vivo, SCE de marcadores fluorescentes permite la visualización directa de la morfología celular, el crecimiento celular, y eventos intracelulares en escalas de tiempo que van desde segundos a días. Si bien esta técnica se utiliza en una variedad de preparaciones en vivo y ex vivo, hemos optimizado esta técnica para su uso en los renacuajos Xenopus laevis. En este artículo de vídeo, se detalla el procedimiento para la SCE de un colorante fluorescente o de ADN plásmido en las neuronas en el cerebro intacto de los albinos renacuajos Xenopus. También se discuten los métodos para optimizar el rendimiento, y mostrar ejemplos de vida de dos fotones de imágenes de fluorescencia de las neuronas marcados con fluorescencia por SCE.

Watch this Scientific Journal Video about Single Cell Electroporation in vivo within the Intact Developing Brain at JoVE.com

Single Cell Electroporation in vivo within the Intact Developing Brain

Una sola célula de electroporación (SCE) es una técnica especializada que permite la entrega de ADN u otras macromoléculas en las células individuales en el tejido intacto, incluso en los preparativos vivo. Aquí detallamos el procedimiento para la SCE de un colorante fluorescente o de ADN plásmido en las neuronas en el cerebro intacto de los renacuajos Xenopus laevis.

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 ...

single-cell-electroporation-in-vivo-within-the-intact-developing-brain

Nanyang Technological University

Research

JoVE Journal

Biology

13.2K vistas.  University of British Columbia - UBC. Una sola célula de electroporación (SCE) es una técnica especializada que permite la entrega de ADN u otras macromoléculas en las células individuales en el tejido intacto, incluso en los preparativos vivo. Aquí detallamos el procedimiento para la SCE de un colorante fluorescente o de ADN plásmido en las neuronas en el cerebro intacto de los renacuajos Xenopus laevis.

Video: Single Cell Electroporation in vivo within the Intact Developing Brain

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

Electroporación de micobacterias

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

Investigación

Biología

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 ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Una sola célula grabaciones de succión de los conos del ratón

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Imágenes in vivo de larvas de Drosophila intactos a nivel sub-celular Resolución

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

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 ...

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

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

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

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

En electroporación in vivo de Morpholinos en la regeneración de adultos de pez cebra Aleta de cola

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

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Imagen confocal de Soltero mitocondriales superóxido Parpadea en corazón intacto o<em&gt; En Vivo</em

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Evaluación de calcio en Sparks intactas las fibras musculares esqueléticas

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

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 ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.