We thank Stephan Uphoff for discussions. 
R.C. was supported by Linacre College, Oxford University. A.P. was supported by the German Academic Exchange Service (DAAD), the German National Academic Foundation and EPSRC. M.S. was supported by the Wellcome Trust. A.N.K. was supported by a UK BBSRC grant (BB/H01795X/1), and a European Research Council Starter grant (261227).

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The authors have nothing to disclose.

Muchos parámetros se pueden variar durante la electroporación de células y adquisición de datos en función del sistema biológico de interés y la naturaleza precisa del experimento (de nivel de celda o el análisis de una sola molécula). Por ejemplo, cuando la electroporación de ADN en bacterias de 0,25 a 5 pmol de fragmentos de dsDNA marcados conduce a una baja eficiencia de la internalización, lo que permite la detección directa de una sola molécula (es decir., Sin la necesidad de photobleaching de antemano). Por encima de 5 pmol ADN de doble cadena, las células tienden a ser muy cargado, un régimen más adecuado para el análisis de una sola célula. Todos los ADNs de la etiqueta también debe ser previamente purificado en gel con el fin de eliminar cualquier rastro de colorante libre (fluoróforo que no ha reaccionado) de la solución de ADN de valores. Además, los posibles problemas con la degradación del ADN, en particular para los experimentos smFRET, pueden abordarse mediante el uso de los ADN con los ácidos nucleicos no naturales, o motivos que protegen termini-exonucleasa accesible como bucles en horquilla. Otra adjustable parámetro en la electroporación es la intensidad de campo aplicada durante la electroporación. La fuerza del campo baja (~ 1 kV / cm) dará lugar a una eficiencia de carga baja adecuada para los estudios de una sola molécula. Intensidades de campo superiores (hasta 1,8 kV / cm) se incrementará la eficiencia de carga; sin embargo, existe una correlación inversa entre la intensidad de campo y la viabilidad celular después de la electroporación (véase la Figura 4). Para referencia, una intensidad de campo normal utilizado para las bacterias y la levadura electroporación es ~ 1,5 kV / cm. La constante de tiempo, que representa la longitud de este decaimiento, es un parámetro conveniente seguir, ya que las gotas constantes de tiempo tan pronto como cualquier fenómeno de formación de arco se produce en la cubeta. En virtud de la configuración normal, la constante de tiempo debe ser mayor que 4 ms; valores más bajos conducirán a una baja eficiencia de carga o células dañadas aún no cargados. La mayoría de electroporators ofrecen otros grados de libertad (tales como &quot;truncamiento pulso&quot; o &quot;forma de impulso&quot;) que puede ser modificado para sintonizar tantocarga de las células y la viabilidad. Hemos aplicado este método para ambas bacterias y levaduras, sin embargo procedimientos similares también deben permitir la internalización de las biomoléculas marcadas en células de mamífero usando los ajustes apropiados electroporador desde su membrana es en realidad menos complejo (una sola bicapa lipídica) y desde la electroporación ya se ha utilizado con tales células 21. Cuando internalizar proteínas marcadas, todo tinte libre necesita ser removido de la solución de proteína de stock etiquetada electroporación antes. Moléculas de colorante gratuitos, debido a su menor tamaño, pueden ser internalizados preferentemente sobre las proteínas de interés, y son difíciles de discriminar durante el análisis de datos (a pesar de su difusión más rápida esperado). Como guía, para una muestra de proteína marcada orgánicamente sea adecuado para la electroporación, la cantidad de colorante libre restante debe ser inferior a 2% (detectada usando el escaneo fluorescente de un SDS-PAGE) 22. Este proceso es particularmente importante,como algunas moléculas podrían pegarse a las membranas externas de bacterias o levaduras electroporadas. A este respecto, la muestra de control negativo debe mostrar la intensidad de fluorescencia por célula claramente inferior a las células electroporadas, idealmente tan bajo como el nivel de autofluorescencia de las células vacías (células que no han sido incubadas con cualquier biomoléculas marcadas con fluorescencia ni electroporadas, Figura 2). Como con dsDNA, la eficiencia de la internalización de proteínas marcadas está vinculada a la cantidad de biomoléculas añadió a las células antes de la electroporación. Sin embargo, otros parámetros, tales como el tamaño y la carga, juegan un papel en la internalización. Las proteínas pequeñas exhiben altas eficiencias de internalización, mientras que las proteínas más grandes (hasta 98 kDa) puede ser internalizado con éxito pero con una eficiencia más baja (Figura 5) 26. El punto isoeléctrico de la proteína, interacciones potenciales con la membrana celular y otros parámetros fisicoquímicos tambiéncarga de las células influencia durante la electroporación. Como resultado, los usuarios necesitan para optimizar los experimentos de su propio sistema, a sabiendas de que una alta concentración inicial de proteína marcada (&gt; 50 micras) y darle la mejor oportunidad para la carga exitosa. La electroporación también ofrece una nueva herramienta para perturbar y analizar la función celular mediante la introducción de proteínas y otras biomoléculas en células (ya sean etiquetados o sin etiqueta). Los experimentos de ARN polimerasa T7 (Figura 5C) presenta un ejemplo de un experimento en el que podemos introducir una biomolécula que puede cambiar la expresión génica in vivo usando electroporación tal. Cuando la realización de experimentos de fluorescencia de una sola molécula, la iluminación TIR generalmente se favorece sobre otros modos de iluminación, ya que ofrece la mejor relación señal-ruido por emocionantes sólo fluoróforos dentro de una sección delgada encima de la superficie cubreobjetos (~ 100 nm). Sin embargo de imagen etiquetada biomoléculas que se difunden dentro de microorganismos vivos podrían volvercuadernillo iluminación más profunda (hasta 0,8 m para E. coli). Iluminación más profunda se consigue en el modo de Hilo, preservando al mismo tiempo una alta relación señal-ruido. Por otro lado, las imágenes de campo amplio es particularmente importante para el análisis photobleaching paso a paso, donde el usuario es la estimación del número de moléculas internalizados por photobleaching una célula cargada entero con alto poder de láser y dividiendo la intensidad inicial de fluorescencia celular por la intensidad unitario producido por una sola molécula (paso photobleaching sola, la Figura 3). También se requiere formación de imágenes de campo amplio para el seguimiento molécula de largo plazo con el fin de localizar las moléculas de difusión de interés incluso si sus trayectorias cubren todo el volumen de la celda. En este protocolo, se presenta cómo electroporación, una técnica estándar para biólogos y bioquímicos para la entrega de ácidos nucleicos en las células, constituye un método simple para la entrega de las biomoléculas fluorescentes en diversos tipos de células. Thes novedoso, técnica de alto rendimiento ofrece una herramienta única de observar moléculas marcadas en su ambiente nativo. En adición biomoléculas marcadas con fluoróforos que cubren una amplia gama de longitudes de onda, la electroporación puede entregar moléculas modificadas con muchos grupos químicos, tales como nucleótidos y aminoácidos no naturales, quelantes de metales, agentes de reticulación, y grupos introducción en jaula. Si el sistema biológico de interés no es esencial para el desarrollo de las células, el gen que codifica para la proteína diana también puede ser borrado (o derribado), asegurando que las proteínas observadas después de la internalización representan todos (o la mayoría) de la piscina intracelular de proteínas . En esencia, la electroporación puede &quot;trasplante&quot; la flexibilidad de bioconjugados in vitro en células vivas y, por tanto, beneficiar los esfuerzos de la biología sintética, la biología de sistemas, y vivo de detección de moléculas individuales en.

La mayoría de los estudios de fluorescencia dentro de las células vivas dependen de fusiones de proteína con proteínas fluorescentes (fps), tales como GFP 1. Estas etiquetas fluorescentes permiten estudios del número de copia, patrón de difusión o la localización de las proteínas implicadas en procesos tales como la expresión de genes o de transporte de membrana 2-7. MF ofrecen una alta especificidad etiquetado, aplicación fácil, y están disponibles en un gran inventario de variantes con diferentes fotofísico y propiedades químicas 1. Sin embargo, fluoróforos orgánicos siguen siendo la elección principal para los experimentos in vitro debido a su mayor fotoestabilidad (hasta 100 veces más estable que los MF) 8,9, pequeño tamaño (hasta 100 veces menor volumen que los MF) y la facilidad de etiquetado intramolecular (principalmente a través de la utilización de residuos de cisteína). Todos estos factores son particularmente importantes para una sola molécula fluorescente y FRET estudios de 10. Varios métodos de internalización combining las ventajas del etiquetado orgánico y en la detección in vivo se han introducido en la última década; sin embargo, tales métodos ya sea emplean polipéptidos relativamente grandes etiquetas (tags por ejemplo, TMP, halo, o 20 kDa SNAP) 11-14, requiere el uso de aminoácidos no naturales 15, o se limita a las células eucariotas grandes, de una sola membrana (por ejemplo. , carga por legrado, jeringa de carga, microinyección) 16-19. Este protocolo describe un novedoso, sencillo y de alto rendimiento método de internalización que las parejas las ventajas de fluoróforos orgánicos con la observación in vivo. Para desarrollar esta técnica, se adaptó el procedimiento de electroporación comúnmente utilizado para transformar las células con el plásmido de ADN 20,21 para cargar microorganismos, tales como E. coli o S. cerevisiae con biomoléculas marcadas orgánicamente. El protocolo consiste en 4 sencillos pasos: la incubación de las células con biomoléculas marcadas,la electroporación, la recuperación celular, y lavado de células para eliminar biomoléculas no internalizadas. A continuación, presentamos este protocolo de electroporación, así como los procesos de formación de imágenes de células y análisis de datos para el estudio de la fluorescencia basada en células y de una sola molécula y FRET señales. La electroporación se basa en la descarga de un campo eléctrico de alta tensión a través de una suspensión de células de baja fuerza iónica para formar poros de la membrana transitorios a través del cual biomoléculas pueden entrar en las células (Figura 1) 20,21. Así como con la transformación de bacterias o levadura con ADN de plásmido, las células tienen que estar preparados antes de la electroporación para asegurar su electrocompetency. Este procedimiento, que consta de varias etapas de lavado con agua, aumenta la permeabilidad de la membrana y disminuye la fuerza iónica de la solución de células para evitar la formación de arco en la cubeta de electroporación. En este protocolo, las células se pueden preparar como se describe a continuación (véase el protocolo: 1,1) o comprados de proveedor comercials. <img alt="Figura 1" src="/files/ftp_upload/52208/52208fig1.jpg" /> Figura 1: Representación esquemática del protocolo de internalización De izquierda a derecha:. Añadir unos pocos microlitros de biomoléculas marcadas a la alícuota de células electrocompetentes (fragmentos de ADN doblemente marcada y bacterias en este ejemplo); incubar 1 a 10 min en hielo y transferir a una cubeta de electroporación previamente enfriada; electroporar y luego añadir 0,5 a 1 ml de medio rico a las células inmediatamente después; incubar a 37 ° C (o la temperatura requerida por el organismo, por ejemplo, 29 ° C para la levadura) para permitir que las células se recuperan; realizar 5 etapas de lavado para eliminar cualquier exceso de moléculas que no son internalizados etiquetados; resuspender el sedimento final en 100-200 l de tampón PBS y pipeta de 10 l en una almohadilla de agarosa; cubrir la almohadilla con un cubreobjetos limpio y la imagen en un microscopio de fluorescencia (en el modo de campo amplio o el modo de HILO). <p clculo = &quot;jove_content&quot;&gt; células electrocompetentes se incuban con las biomoléculas marcadas justo antes de la electroporación, las cuales se pueden realizar utilizando electroporators estándar que se encuentran en la mayoría de los laboratorios de bioquímica. Inmediatamente después de la electroporación, las células se incuban en un medio rico que permite su recuperación antes del lavado (Figura 1). El exceso de biomoléculas marcadas no internalizados se retira primero por lavado en un tampón que contiene una concentración bastante alta de sal y un poco de detergente (véase el protocolo: 3,3). La presencia de sal interrumpe interacciones electrostáticas no específicas formados por biomoléculas marcadas no internalizados que de otro modo pueden pegar en la membrana externa. Del mismo modo, la presencia de detergente en el tampón de lavado interrumpe interacciones hidrófobas no específicas. Mientras que la internalización de ADN es directa (Figura 2), las precauciones deben tomarse cuando internalizar proteínas marcadas mediante electroporación. En primer lugar, la muestra de stock de proteína marcada orgánicamente todavía puede contener un pequeño porcentaje de colorante libre. Moléculas de colorante libres son mucho más pequeñas que las proteínas y por lo tanto pueden ser internalizados preferentemente. Para asegurarse de que la gran mayoría de las moléculas fluorescentes internalizadas observadas corresponden a la proteína de interés, la muestra inicial de proteína debe contener menos de ~ 2% de colorante libre (Figura 5) 22. El exceso de proteínas marcadas no internalizados también puede adherirse a la membrana celular externa después de la electroporación; este fenómeno es específico de la proteína y necesita ser comprobada para cada nueva proteína. Proponemos varias opciones que permiten la eliminación de las proteínas no internalizadas de la muestra celular cargado (Ver PROTOCOLO: 3.3.3). Finalmente, las células se resuspenden en un pequeño volumen de tampón de fosfato y se pipetearon en una almohadilla de agarosa, lo que permite su formación de imágenes en un microscopio de fluorescencia. La inmovilización en las almohadillas de agarosa es un simplE y forma eficiente de imágenes de células sobre un cubreobjetos sin dañar su integridad. La almohadilla debe contener un medio de cultivo bajo fluorescencia. Imágenes de células se puede realizar tanto en campo amplio, el total de fluorescencia de reflexión interna (TIRF) o utilizando HILO (muy inclinadas y laminado Hoja óptica) microscopía. En la configuración HILO, el rayo láser penetra más profundamente en la muestra que en TIRF, sin embargo, no se ilumina toda la muestra como para campo amplio, lo que permite una mayor relación señal-ruido 23. Dependiendo de la resolución de energía y el tiempo de láser utilizado, biomoléculas internalizados pueden ser contados (usando análisis por pasos-photobleaching, Figura 3), localizados, o seguido 24-28. Internalización de los constructos doblemente marcada con un par de FRET de fluoróforos permite la cuantificación de FRET tanto a una sola célula o los niveles de una sola molécula (Figura 6). Diferentes parámetros se pueden variardependiendo de la salida deseada y el sistema biológico estudiado. En primer lugar, la cantidad de material interiorizado por célula se puede ajustar mediante la alteración de la concentración de biomoléculas marcadas añadió a las células antes de electroporación (Figura 2). Intensidad de campo electroporación también influir tanto en la eficiencia de carga y la viabilidad celular; como se esperaba, mientras que los aumentos de la eficiencia de carga con el aumento de la intensidad de campo, la viabilidad de las células electroporadas disminuye (Figura 4A). Ambos parámetros pueden ser cuantificados mediante el registro del porcentaje de carga y división de las células después de la electroporación. Este ensayo de viabilidad junto con imágenes de fluorescencia también verifica la observación de biomoléculas internalizados en las células vivas y permite la observación continua durante varias generaciones (Figura 4B). En resumen, este protocolo permite la internalización de la etiqueta fluorescente de ADN y proteínas en moléculasE. coli o S. cerevisiae 26. Las moléculas individuales marcadas con fluoróforos orgánicos pueden ser rastreados con alta resolución espacial y temporal para escalas de tiempo de un orden de magnitud más que los MF. Finalmente, este método es compatible con widefield, TIRF y detección confocal, así como esquemas de excitación pulsada, como ALEX (alternando excitación láser 28,29).

<table border="0" cellpadding="0" cellspacing="0" width="834">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Name of Material/ Equipment</td>
			<td style="width:227px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:321px;">ElectroMax DH5-alpha Comptent cells</td>
			<td style="width:227px;">Invitrogen</td>
			<td style="width:119px;">11319-019</td>
			<td>or any other commercial or lab-mage electrocompetant bacteria or yeast.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">EZ Rich Defined Madia</td>
			<td style="width:227px;">Teknova</td>
			<td style="width:119px;">M2105</td>
			<td>low fluorescence rich media</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MicroPulser Electroporation Apparatus&#160;</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">165-2100</td>
			<td>or any classical electroporator for microorganism transformation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Certified Molecular Biology agarose</td>
			<td style="width:227px;">Biorad</td>
			<td style="width:119px;">161-3100</td>
			<td>low fluorescence agarose for agarose pad</td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:321px;">Microscope coverslips No 1.5 thickness</td>
			<td style="width:227px;">Menzel</td>
			<td style="width:119px;">BB024060SC</td>
			<td>remove background particles by heating slides in furnace at 500 &#176;C for 1h</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Single-molecule fluorescence microscope</td>
			<td style="width:227px;">Home-built</td>
			<td style="width:119px;"></td>
			<td>described in REFs</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Localization software</td>
			<td style="width:227px;">Custom-written, available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB and C++ software package that can be adapted for localization analysis.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:321px;">Tracking software</td>
			<td style="width:227px;">Available online</td>
			<td style="width:119px;"></td>
			<td>MATLAB implementation by Blair and Dufresne.&#160;</td>
		</tr>
	</tbody>
</table>

internalization and observation of fluorescent biomolecules in living microorganisms via electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Watch this Scientific Journal Video about Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation at JoVE.com

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

Internalización y Observación de fluorescente Biomoléculas en microorganismos que viven a través de electroporación

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

internalization-observation-fluorescent-biomolecules-living

Research

JoVE Journal

Biology

16.7K vistas.  University of Oxford. Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

Video: Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

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

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

Imágenes de células vivas de la F-actina Dynamics a través de Microscopía Fluorescente Speckle (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

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 ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Celular imagen Cambiar forma de vida Drosophila Los embriones

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

Control de la ubiquitina-proteasoma actividad en las células vivas utilizando un degron (dgn)-desestabilizado la proteína verde fluorescente (GFP)-basada en proteína indicadora

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific properties, such as binding affinity, agonist or antagonist behavior, or stability, relative to the native sequence. Protein design lies at the center of current advances drug design and discovery. Not only does protein design provide predictions for potentially useful drug targets, but it also enhances our understanding of the protein folding process and protein-protein interactions. Experimental methods such as directed evolution have shown success in protein design. However, such methods are restricted by the limited sequence space that can be searched tractably. In contrast, computational design strategies allow for the screening of a much larger set of sequences covering a wide variety of properties and functionality. We have developed a range of computational de novo protein design methods capable of tackling several important areas of protein design. These include the design of monomeric proteins for increased stability and complexes for increased binding affinity.
To disseminate these methods for broader use we present Protein WISDOM (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>), a tool that provides automated methods for a variety of protein design problems. Structural templates are submitted to initialize the design process. The first stage of design is an optimization sequence selection stage that aims at improving stability through minimization of potential energy in the sequence space. Selected sequences are then run through a fold specificity stage and a binding affinity stage. A rank-ordered list of the sequences for each step of the process, along with relevant designed structures, provides the user with a comprehensive quantitative assessment of the design. Here we provide the details of each design method, as well as several notable experimental successes attained through the use of the methods.

Protein WISDOM: A Workbench for In silico De novo Design of BioMolecules

We developed computational de novo protein design methods capable of tackling several important areas of protein design. To disseminate these methods we present Protein WISDOM, an online tool for protein design (<a href="http://www.proteinwisdom.org" target="_blank">http://www.proteinwisdom.org</a>). Starting from a structural template, design of monomeric proteins for increased stability and complexes for increased binding affinity can be performed.

SABIDURÍA Proteína: Un banco de trabajo para<em&gt; En silico</em&gt;<em&gt; De novo</em&gt; Diseño de biomoléculas

The aim of de novo protein design is to find the amino acid sequences that will fold into a desired 3-dimensional structure with improvements in specific ...

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 ...

Tracking Drug-induced Changes in Receptor Post-internalization Trafficking by Colocalizational Analysis

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and overall cell responsiveness to ligands and is, itself, influenced by intra- and extracellular conditions, including ligand-induced signaling. Optimized for use with monolayer-plated cultured cells, but extendable to free-floating tissue slices, this protocol uses immunolabelling and colocalizational analysis to track changes in intracellular receptor trafficking following both chronic/prolonged and acute interventions, including exogenous drug treatment. After drug treatment, cells are double-immunolabelled for the receptor and for markers for the intracellular compartments of interest. Sequential confocal microscopy is then used to capture two-channel photomicrographs of individual cells, which are subjected to computerized colocalizational analysis to yield quantitative colocalization scores. These scores are normalized to permit pooling of independent replicates prior to statistical analysis. Representative photomicrographs may also be processed to generate illustrative figures. Here, we describe a powerful and flexible technique for quantitatively assessing induced receptor trafficking.

Receptor trafficking modulates signaling and cell responsiveness to ligands and is, itself, responsive to cell conditions, including ligand-induced signaling. Here, we describe a powerful and flexible technique for quantitatively assessing drug-induced receptor trafficking using immunolabeling and colocalizational analysis.

Seguimiento de cambios inducidos por fármacos en los receptores post-internalización Trata de Análisis Colocalizational

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

-Alta resolución de imagen de lapso de tiempo y análisis automatizado de los microtúbulos en la dinámica de estar umbilical humana Las células endoteliales de vena

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. To improve the mating efficiency of B. Xylophilus and to increase the chances of mating observation, virgin adults were cultured and used for the investigation. Eggs were obtained by keeping the nematodes in water and allowing the females to lay eggs for 10 min. The second-stage juveniles (J2) were synchronized by incubating the eggs for 24 h at 25 &#176;C in the dark, and the early J4 were obtained by culturing the J2 with grey mold, Botrytis cinerea, for another 52 h. At this time point, most J4 nematodes could be clearly distinguished as being male or female using their genital morphology. The male and female J4 were collected and cultured separately in two different Petri dishes for 24 h to get virgin adult nematodes. A virgin male and a virgin female were paired in a drop of water in the well of a concave slide. The mating behavior was filmed with a video recorder under a stereomicroscope. The whole period of the mating process was 82.8 &#177;3.91 min (mean &#177;SE) and could be divided into 4 different phases: searching, contacting, copulating, and lingering. The mean minutes of duration were 21.8 &#177; 2.0, 28.0 &#177; 1.9, 25.8 &#177; 0.7 and 7.2 &#177; 0.5, respectively. Eleven sub-behaviors were described: cruising, approaching, encountering, touching, hooping, locating, attaching, ejaculating, separating, quiescence, and roaming. Interestingly, obvious intra-sexual competition was observed when one female was grouped with 3 males or one male with 3 females. This protocol is useful and valuable, not only in investigating the mating behavior of B. xylophilus, but also in acting as a reference for ethological studies of other nematodes.

Observation and Quantification of Mating Behavior in the Pinewood Nematode, Bursaphelenchus xylophilus

A protocol for investigating the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, is presented. Behavioral features of B. xylophilus are described in the mating process.

La observación y cuantificación de la conducta de apareamiento en el nematodo del pino,<em&gt; Bursaphelenchus xylophilus</em

A method for observing and quantifying the mating behavior of the pinewood nematode, Bursaphelenchus xylophilus, was established under a stereomicroscope. ...