Esta obra fue financiada por los Institutos Nacionales de Salud subvenciones NS047726, NS072027 y AR052646.

Los métodos de este estudio se realizaron en ética de acuerdo con la Escuela Feinberg de la Universidad Northwestern de Medicina Institucional Cuidado de Animales y el empleo Comisión (IACUC) directrices aprobadas. Se hicieron todos los esfuerzos para minimizar el sufrimiento. 1. Procedimiento experimental in vivo electroporación del Flexor digitorum brevis (FDB) Muscle Bundle de Ratón <ol><li> Plásmidos de diseño para la expresión in vivo usando un promotor conocido para expresar en células de mamífero (por ejemplo, citomegalovirus, CMV) o en el músculo (por ejemplo, Músculo creatina quinasa, MCK) 6,7. Plásmidos más grandes pueden expresar en los niveles inferiores. El promotor CMV se ha utilizado ampliamente 6,7. </li><li> Purificar plásmidos de E. coli utilizando endotoxinas condiciones libres por las instrucciones del fabricante. Disolver el plásmido en, tampón TE libre de endotoxina estéril a 2-5 mg de plásmido / l. </li><li> Diluir y alícuota del plásmido a un concentration de 20 g de ADN en 10 l de endotoxina estéril libre de TE (2 g / l) tampón por inyección en un tubo de microcentrífuga estéril. Preparar una alícuota por inyección. </li><li> Diluir el stock de hialuronidasa a 1x estéril con salina tamponada con fosfato sin calcio y magnesio (PBS - / -) dando una concentración final de 8 unidades. Alícuota de 10 l de hialuronidasa diluida por inyección en un tubo de microcentrífuga estéril. </li><li> Anestesiar el ratón usando 2,5% isoflurano hasta sedado y no responde a la cola o pellizco del dedo del pie. Colocar el animal en posición supina sobre una almohadilla térmica o una mesa de calentamiento para mantener la temperatura corporal normal. </li><li> Limpiar la almohadilla de la pata del ratón con una aplicación alternada de un lavado de povidona yodada y alcohol tres veces usando una técnica de preparación se origina en el sitio de la aguja y se irradia hacia el exterior. Esta técnica mantiene el sitio de la aguja en una condición aséptica, minimizando introducción patógeno. </li><li> Inyectar la almohadilla plantar del XXe ratón con 10 l de la solución de hialuronidasa 1x (2 mg / ml = 8 unidades) entre la piel y el músculo usando una jeringa de 1 ml estéril. La hialuronidasa digiere componentes de la matriz extracelular para permitir la entrada más eficiente de plásmido en miofibras. Introduzca en la base del talón y hacer avanzar la aguja hacia los dedos de los pies. Suelte lentamente el líquido que se retrae la aguja. El sitio de la inyección bajo la piel se expandirá y comenzar a girar rosa. </li><li> Repita los pasos 1,6 y 1,7 en el pie contralateral. </li><li> Desconecte la anestesia y colocar el ratón de nuevo en la jaula. Deje que el ratón para recuperarse totalmente de la anestesia. </li><li> Espere 2 horas. </li><li> Repita los procedimientos de esterilización anestesiar y pies, pasos 1.5 y 1.6. </li><li> Inyectar el ADN de plásmido se diluyó en la almohadilla de la pata de la misma manera como la hialuronidasa. Asegúrese de que el volumen máximo es de 20 l o menos. Para expresión dual plásmido inyectar 20 g de cada plásmido con un volumen máximo de 20 y# 181; l o menos. </li><li> Repetir la inyección de plásmido en la almohadilla de la pata contralateral o usar el pie contralateral para inyecciones de control. </li><li> Al pie que fue inyectado inicialmente, levantar la piel lejos del músculo subrayando con unas pinzas finas y colocar una aguja de 27 G a través de las bolas de la piel cerca de los dedos de los pies en una posición perpendicular a la línea de sanar-dedo del pie. Tome un segundo G aguja estéril 27 y colocarla horizontalmente a través de la piel en el talón. Coloque agujas paralelas entre sí ~ 1 cm de separación. </li><li> El uso de pinzas de cocodrilo, electrodos de fijación a las agujas paralelas asegurándose de que las agujas no entren en contacto entre sí. El uso de la plastilina puede asegurar las abrazaderas en su lugar. </li><li> Conectar los electrodos a un estimulador eléctrico. </li><li> Electroporar el músculo mediante la aplicación de 20 pulsos, 20 ms de duración / cada uno, a 1 Hz, a 100 V / cm. Los dedos de los pies pueden contraerse con la estimulación. </li><li> Repita el procedimiento de estimulación en el pie contralateral si es necesario. </li><li>Retire las agujas. Limpie la almohadilla plantar con matorrales povidona yodada. </li><li> Apague la anestesia. Devuelva el ratón a la jaula de la recuperación y la actividad monitor. Si se llevó a cabo el procedimiento como se esperaba, el animal debe recuperar la deambulación normal dentro de 30 minutos y muestran ninguna diferencia en la deambulación o la actividad a nivel de comportamiento pre-inyección. Por lo tanto, no se necesitan analgésicos después del procedimiento. Después de la recuperación, devuelva el animal para el vivero. Nota: Expresión de proteínas se puede ensayar ya a los 48 horas después de la electroporación. Sin embargo, para permitir la recuperación más completa después de la electroporación, se prefiere el aislamiento y estudio de las miofibras 7 - 14 días después de la electroporación 10. La expresión puede ensayarse tanto tiempo como 4 semanas después de la electroporación. </li></ol> 2. Procedimiento Experimental para el aislamiento de los (FDB) Fibras Flexor Brevis <ol><li> Colagenasa deshielo. Para cada paquete FDB preparar dos pozos en una placa de cultivo tisular de 12 pocillos. En un pozo añadir 1 ml de pre-calentado Eagles Modificado de Dulbecco los medios de comunicación más albúmina de suero bovino (DMEM + BSA) y en el otro bien añadir 1 ml de solución de Ringer pre-calentado. Además, añadir frescas 10 ml de solución Ringers 1x a los 35 mm plato Sylgard. </li><li> Sacrificio del animal a través de CO 2 o la inhalación de gases anestésicos, seguido por dislocación cervical u otro método para asegurar la muerte. </li><li> Retire las patas traseras por encima de la articulación del tobillo usando una hoja de afeitar limpia y sumergir en solución 1x timbres en los 35 mm plato Sylgard. </li><li> Pin un pie apretado, único hacia arriba, hacia el plato Sylgard con 30 agujas G a través de cada una de las puntas de los pies y 5 en el tobillo. </li><li> Comenzando en el tobillo, cortar la piel hacia arriba el lado del pie a lo largo de la línea del cabello hacia los dedos, con cuidado de no cortar el músculo debajo de la piel. Repetir en el otro lado del pie. </li><li> Sosteniendo la piel en la base del tobillo con unas pinzas finas, cortar la piel a través del talón. </li><li> El levantamiento de la pielbatir con unas pinzas, señalar sus tijeras hacia la piel para no nick del músculo. Cortar el tejido conectivo subyacente, eliminando de forma eficaz la piel. </li><li> Para quitar el haz muscular FDB cortó el tendón grande en el talón para liberar el tendón del hueso. Sosteniendo el tendón con una pinza, diseccionar el paquete FDB del tendón grande blanco retirando cuidadosamente cualquier tejido conectivo unido a la FDB con unas tijeras. Si se realiza correctamente el paquete levantará fácilmente fuera de los tendones subyacentes revelan tendones blancas brillantes que llevan a los dígitos individuales. Cortar la FDB en los tendones cercanos a los dedos de los pies liberar el paquete desde el pie. </li><li> Coloque todo el paquete muscular en el pozo que contiene DMEM + BSA. </li><li> Repita con el pie contralateral. </li><li> Una vez que todos los músculos de FDB son aisladas, añadir 100 l de colagenasa a cada pocillo que contiene DMEM + BSA y el músculo. </li><li> Compruebe el éxito de la electroporación en un microscopio fluorescente invertido antes de la digestión. Si properl hechoy, arriba de 90% de eficiencia de transfección se puede lograr. </li><li> Incubar la placa en una humidificado, 37 ° C incubadora a 10% de CO 2 durante aproximadamente 1 hr. El tiempo de incubación puede variar dependiendo del modelo de la enfermedad y el fondo y el lote de colagenasa. </li><li> Después de 1 hora, transferir cuidadosamente el paquete FDB a la solución de Ringer único pozo que contiene. </li><li> Se tritura el paquete FDB en el pozo con la solución de Ringers utilizando una pipeta de 1 ml. Cortar el extremo de la punta de pipeta con una hoja de afeitar limpia de tal manera que el haz puede pasar fácilmente a través de la punta de pipeta. Se tritura suavemente el músculo (~ 15 veces) permitiendo que el haz pase a arriba y hacia abajo en paralelo a la punta. Fibras intactas se caerán en la solución de Ringer. Cuando las fibras no caen fácilmente el paquete, colocar el músculo de nuevo en solución de colagenasa. </li><li> Placa las fibras aisladas en el plato ~ 500 l por placa. </li><li> Permitir que el músculo para digerir en el pozo durante 15 min. </li><li> Repita los pasos 2.15a 2.17 hasta que el paquete disminuye de tamaño y la mayoría de las fibras se disocia del haz (por lo general un total de 2.3 veces). Una vez disociado el músculo pasa fácilmente a través de una punta de pipeta de 1 ml. </li><li> Deje que las fibras se adhieren al plato durante un mínimo de 15 min. Solución de Ringer fresco puede ser añadido a la placa para diluir la colagenasa residual o cambiado en función de las limitaciones de tiempo. Nota: Las fibras están ahora listos para la imagen. </li></ol> 3. Procedimiento experimental para la visualización de daño de la membrana inducida por láser utilizando microscopía confocal <ol><li> Fibras de carga inmediatamente antes de la obtención de imágenes con 300 l de 2,5 M FM 1-43 del FM 4-64 colorante en solución de Ringer. Generalmente, las fibras de la imagen 15 min a 2 hr después del aislamiento. Sin embargo, las fibras se pueden visualizar hasta después de aislamiento 24 horas. </li><li> Colocar el plato de fondo de cristal en un microscopio confocal equipado con un láser UV y un objetivo 60X / 63X. </li><li> Localiza una fibra. Asegúrese de que la fibra se une firmementeal plato tocando la base del microscopio. Excluir fibras que fueron dañados en el proceso de disociación. Fibras dañadas pueden contener vesículas de membrana, los altos niveles de captación de tinte FM, rizo o doblar intensamente. </li><li> Para inducir daño de la membrana sobre la fibra, posicionar el sarcolema en el centro de la ventana de imagen con un claro campo de núcleos. Concéntrese en el sarcolema utilizando el canal de colorante FM tal que el sarcolema es extremadamente agudo. </li><li> Coloque la cruz ROI en el sarcolema utilizando el canal de FM 4-64. </li><li> Irradiar un punto de 1 píxel en una potencia del 80% durante 3 s (~ 350 nm x 350 nm área). Colorante FM entrará en la celda en el sitio de la lesión. Potencia y el tiempo se pueden ajustar para aumentar o disminuir el área de insulto. </li><li> Capturar imágenes de la fibra antes del daño, en el momento del daño, cada 2 seg post-daño, entonces cada 10 seg durante un máximo de 5 min. Timing se puede ajustar dependiendo de la necesidad. Las imágenes individuales se pueden convertir en películas a intervalos en Imagen J. </li><li> Repita los pasos3.3 a 3.7 en un máximo de 3 fibras por plato. </li></ol>

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El uso de electroporación para estudiar las proteínas etiquetadas con fluorescencia in vivo es ideal para el estudio de los músculos debido a la ausencia de modelos de líneas celulares adecuadas que reproducen fielmente toda la estructura celular del músculo. Este protocolo describe la utilización de la electroporación y la microscopía confocal de alta resolución para proporcionar imágenes detalladas de la localización de proteínas y la translocación después de la herida láser. Este método puede...

Miofibras individuales son células sincitiales grandes, altamente organizados. Hay unos cuantos modelos de cultivo celular de los músculos; Sin embargo, estos modelos tienen como principal limitación que no diferencian plenamente en miofibras maduras. Por ejemplo, las líneas celulares C2C12 y L6 se derivan de ratones y ratas, respectivamente a 1-4. En condiciones de privación de suero, las células mononucleares &quot;-mioblastos como&quot; dejan la proliferación, se someten a la retirada del ciclo celular, y entran en el programa miogénica formando células multinucleadas con sarcómeros, conocidos como miotubos. Con las condiciones de cultivo prolongados, miotubos pueden exhibir propiedades contráctiles y &quot;contracción&quot; en la cultura. Líneas celulares humanas han también ahora han establecido 5. Además de estas líneas de células inmortalizadas, mioblastos mononucleares se pueden aislar a partir de músculo y en las condiciones similares de privación de suero formará miotubos. Estas líneas celulares y cultivos de mioblastos primarios son altamente utilizarful, ya que pueden ser transfectadas con plásmidos o transducidas con virus y se usaron para estudiar los procesos biológicos celulares básicas. Sin embargo, estas células, incluso cuando inducida para formar miotubos, carecen de muchas de las características sobresalientes de organización músculo maduro. Específicamente, miotubos son mucho menores que las miofibras maduras individuales y carecen de la forma normal de miofibras. Críticamente, miotubos carecen transversales (T-) túbulos, la red membranosa necesaria para una eficiente Ca 2 + en libertad en todo el sarcoplasma. Un método alternativo para mioblastos primarios o líneas de células miogénicas implica el uso de miofibras maduras. La transgénesis se puede emplear para establecer la expresión de proteínas etiquetadas, pero este método es costoso y consume tiempo. In vivo electroporación de músculo de ratón se ha convertido en un método preferido por su velocidad y fiabilidad 6-10. Los métodos para la electroporación in vivo y el aislamiento eficiente de miofibras hahe sido optimizado para el músculo flexor corto de los dedos 6 ratón (FDB). Los métodos se pueden completar fácilmente y son mínimamente invasivo para inducir la expresión in vivo a partir de plásmidos. Este enfoque está ahora combinada con métodos de imagen de alta resolución de imagen, incluyendo después de la interrupción del laser del 6,7 sarcolema. La combinación de colorantes y expresión de proteínas marcadas con fluorescencia fluorescentes puede ser usado para monitorear los procesos biológicos celulares en miofibras maduras.

<table>
	<tbody>
		<tr>
			<td>DMEM+A2:D32D42A2:D31AA2:D29</td>
			<td>Life Technologies</td>
			<td>11995-073</td>
			<td>Dissection Media 
			Dilution: 50ml +100g BSA</td>
		</tr>
		<tr>
			<td>Collagenase, Type II</td>
			<td>Life Technologies</td>
			<td>17101-015</td>
			<td>Fiber Digestion 
			Dilution: 160mg / 4ml DMEM (stock 40mg/ml aliquot)</td>
		</tr>
		<tr>
			<td>Falcon 12-well dishes</td>
			<td>Fisher Scientific</td>
			<td>877229</td>
			<td>Fiber Digestion</td>
		</tr>
		<tr>
			<td>Ringers Solution</td>
			<td></td>
			<td></td>
			<td>Fiber Digestion and Imaging 
			Dilution: 146 mM NaCl 
			5 mM&#160;&#160; KCl 
			2 mM&#160;&#160; CaCl2 
			1 mM&#160;&#160; MCl2 
			10mM HEPES 
			pH7.4 in H2O</td>
		</tr>
		<tr>
			<td>1ml Pipettes</td>
			<td>Axygen</td>
			<td>T-1000-C-R</td>
			<td>Digestion</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Razor Blades</td>
			<td>Personna</td>
			<td>94-120-71</td>
			<td>Digestion</td>
		</tr>
		<tr>
			<td>Precision Glide 30G needle</td>
			<td>BD Biosciences</td>
			<td>305128</td>
			<td>Dissection</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>1x PBS (-/-)</td>
			<td>Life Technologies</td>
			<td>14190-250</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Sylgard 184</td>
			<td>Fisher Scientific</td>
			<td>50-366-794</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>FST by Dumont</td>
			<td>#5/45</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Roboz</td>
			<td>RS-4913</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>World Precision Instruments</td>
			<td>500260</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A7906</td>
			<td>Dissection media 
			Dilution: 100mg / 50ml DMEM</td>
		</tr>
		<tr>
			<td>Falcon 60mm dishes</td>
			<td>Fisher Scientific</td>
			<td>08772B</td>
			<td>Dissection- used to hold sylgard</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Clay</td>
			<td></td>
			<td></td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Stimulator&#160;</td>
			<td>Grass</td>
			<td>s88x</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Clamps</td>
			<td>Radio Shack</td>
			<td>270-356</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Triadine</td>
			<td>Aplicare</td>
			<td>82-220</td>
			<td>Electroporation</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Precision Glide 27G needle</td>
			<td>BD Biosciences</td>
			<td>305136</td>
			<td>Electroporation</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>Simulator</td>
			<td>Grass</td>
			<td>SIU-V</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Gauze</td>
			<td>Covidien</td>
			<td>2146</td>
			<td>Electroporation</td>
		</tr>
		<tr>
			<td>Hyaluronidase</td>
			<td>Sigma</td>
			<td>H4272</td>
			<td>Injection 
			Dilution: Add 160ul PBS (1X) to 40&#181;l 5x stock</td>
		</tr>
		<tr>
			<td>1cc U-100 Insulin Syringe (28G1/2)&#160;</td>
			<td>BD Biosciences</td>
			<td>329420</td>
			<td>Injection</td>
		</tr>
		<tr>
			<td>Eppendorf Tubes</td>
			<td>Fisher Scientific</td>
			<td>02-682-550</td>
			<td>Injection</td>
		</tr>
		<tr>
			<td>35mm glass bottom Microwell dish</td>
			<td>MaTek</td>
			<td>P35G-1.5-14-C</td>
			<td>Microscopy</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>FM 4-64</td>
			<td>Life Technologies</td>
			<td>T-13320</td>
			<td>Microscopy 
			Dilution: Final 2.5 ug/ml</td>
		</tr>
		<tr>
		</tr>
		<tr>
			<td>FM 1-43</td>
			<td>Life Technologies</td>
			<td>T-35356</td>
			<td>Microscopy 
			Dilution: Final 2.5 ug/ml</td>
		</tr>
		<tr>
			<td>Endotoxinfree Plasmid Maxi Kit</td>
			<td>Qiagen</td>
			<td>12362</td>
			<td>Plasmid Purification</td>
		</tr>
		<tr>
		</tr>
	</tbody>
</table>

dna electroporation, isolation and imaging of myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

La electroporación es una técnica mínimamente invasiva que introduce de manera eficiente plásmido de ADN purificado en el flexor digitorum brevis (FDB) haz muscular (Figura 1A). Siete días después de la transfección con fluorescencia de proteínas marcadas se visualizaron en miofibras aisladas (Figura 1B). Fibras aisladas se sembraron a continuación, y se prepararon para formación de imágenes en un microscopio confocal. Las fibras pueden ser so...

Watch this Scientific Journal Video about DNA Electroporation, Isolation and Imaging of Myofibers at JoVE.com

DNA Electroporation, Isolation and Imaging of Myofibers

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

ADN electroporación, aislamiento e Imagen de miofibras

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...

dna-electroporation-isolation-and-imaging-of-myofibers

Research

JoVE Journal

Biology

12.1K vistas.  Northwestern University. This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure. 

Video: DNA Electroporation, Isolation and Imaging of Myofibers

Isolation of Genomic DNA from Mouse Tails

El aislamiento de ADN genómico de colas de ratón

Investigación

Biología

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 ...

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 ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

De dos y tres dimensiones imágenes de células vivas de las proteínas de respuesta al daño del ADN

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Aislamiento y cultivo de fibras individuales y sus células satélite del músculo esquelético adulto

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

Análisis de embriones y larvas de pez cebra esqueléticas miofibras de Preparativos disociadas

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

Combinando Manipulación sola molécula y de imagen para el estudio de las interacciones proteína-ADN

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

Imágenes en tiempo real de Soltero Engineered ARN transcritos en células vivas utilizando Ratiometric bimoleculares Balizas

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Reduced-gravity Environment Hardware Demonstrations of a Prototype Miniaturized Flow Cytometer and Companion Microfluidic Mixing Technology

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If humans are to travel beyond low Earth orbit, a transition towards space-ready, point-of-care (POC) testing is required. Such testing needs to be comprehensive, easy to perform in a reduced-gravity environment, and unaffected by the stresses of launch and spaceflight. Countless POC devices have been developed to mimic laboratory scale counterparts, but most have narrow applications and few have demonstrable use in an in-flight, reduced-gravity environment. In fact, demonstrations of biomedical diagnostics in reduced gravity are limited altogether, making component choice and certain logistical challenges difficult to approach when seeking to test new technology. To help fill the void, we are presenting a modular method for the construction and operation of a prototype blood diagnostic device and its associated parabolic flight test rig that meet the standards for flight-testing onboard a parabolic flight, reduced-gravity aircraft. The method first focuses on rig assembly for in-flight, reduced-gravity testing of a flow cytometer and a companion microfluidic mixing chip. Components are adaptable to other designs and some custom components, such as a microvolume sample loader and the micromixer may be of particular interest. The method then shifts focus to flight preparation, by offering guidelines and suggestions to prepare for a successful flight test with regard to user training, development of a standard operating procedure (SOP), and other issues. Finally, in-flight experimental procedures specific to our demonstrations are described.

Spaceflight blood diagnostics need innovation. Few demonstrations have been published illustrating in-flight, reduced-gravity health diagnostic technology. Here we present a method for construction and operation of a parabolic flight test rig for a prototype point-of-care flow-cytometry design, with components and preparation strategies adaptable to other setups.

Reducción de gravedad Demostraciones entorno de hardware de un prototipo miniaturizado citómetro de flujo y Compañero Microfluidos Tecnología de mezcla

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If ...

Amplification, Next-generation Sequencing, and Genomic DNA Mapping of Retroviral Integration Sites

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of diverse libraries of retroviral integration sites using ligation-mediated PCR (LM-PCR) amplification and next-generation sequencing (NGS), (2) mapping the genomic location of each virus-host junction using BEDTools, and (3) analyzing the data for statistical relevance. Genomic DNA extracted from infected cells is fragmented by digestion with restriction enzymes or by sonication. After suitable DNA end-repair, double-stranded linkers are ligated onto the DNA ends, and semi-nested PCR is conducted using primers complementary to both the long terminal repeat (LTR) end of the virus and the ligated linker DNA. The PCR primers carry sequences required for DNA clustering during NGS, negating the requirement for separate adapter ligation. Quality control (QC) is conducted to assess DNA fragment size distribution and adapter DNA incorporation prior to NGS. Sequence output files are filtered for LTR-containing reads, and the sequences defining the LTR and the linker are cropped away. Trimmed host cell sequences are mapped to a reference genome using BLAT and are filtered for minimally 97% identity to a unique point in the reference genome. Unique integration sites are scrutinized for adjacent nucleotide (nt) sequence and distribution relative to various genomic features. Using this protocol, integration site libraries of high complexity can be constructed from genomic DNA in three days. The entire protocol that encompasses exogenous viral infection of susceptible tissue culture cells to integration site analysis can therefore be conducted in approximately one to two weeks. Recent applications of this technology pertain to longitudinal analysis of integration sites from HIV-infected patients.

We describe a protocol for amplifying retroviral integration sites from the genomic DNA of infected cells, sequencing the amplified virus-host junctions, and then mapping these sequences to a reference genome. We also describe techniques to quantify the distribution of integration sites relative to various genomic annotations using BEDTools.

Amplificación, secuenciación de próxima generación, y Genómica Mapeo de ADN retrovirales sitios de integración

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of ...