KO con el apoyo de una beca de la Asociación Americana del Corazón.

Antes de empezar Corazones adultos <ol><li> Recién preparar hemolinfa artificial (IA) solución que contiene 108 mm de Na +, 5 mM K +, 2 mM Ca 2 +, 8 mm de MgCl2, 1 mM NaH 2 PO 4, 4 mm de NaHCO3, sacarosa 10 mM, trehalosa 5 mM, HEPES 5 mM (pH 7.1, todos los reactivos de Sigma Chemicals). La sacarosa y la trehalosa, debe añadirse a la AH de las soluciones madre refrigerados antes de su uso con el fin de prevenir la contaminación bacteriana. </li><li> Traiga AH a temperatura ambiente y oxigenar la solución por vía aérea-burbujas durante al menos 15 min. </li><li> Tire de varios capilares finos (por ejemplo, capilares de vidrio, 100 ul, VWR) necesarios para la eliminación de grasa. </li></ol> Corazones larvas <ol><li> Sylgard recubierto cubreobjetos: con un palillo de dientes, la transferencia de unos 50-70μl de solución Sylgard en un cubreobjetos 22x22mm. Deje que el endurecimiento Sylgard a 65 ° C durante la noche. </li><li> Preparar el Broadie y tampón de Bate (135 NaCl, KCl 5 mM, 4 mM MgCl 2, 2 mM CaCl 2, TES 5mM, sacarosa 36 mm;. PH 7,15 Ver Broadie y Bate, 1993). </li><li> Prepare las herramientas necesarias para aplicar el pegamento histoacryl. En primer lugar, tirar de varios capilares finos (por ejemplo, la Ciencia Productos GB100T8P). Conecte una punta de la pipeta 1,5 ml con un tubo de plástico delgado (Tygon, 1 / 32 &quot;) mediante la inserción de la pequeña en el tubo. Insertar un tubo capilar con su amplia apertura hacia el otro extremo. De carga del capilar con pegamento Histoacryl por inmersión de la capilaridad en una gota de pegamento y una succión suave en la punta de plástico. Cuando se trabaja con el pegamento que uno debe tener un vaso capilar segunda mano para quitar exceso de pegamento en la punta de la cola capilar. La cola es un líquido hasta que se entra en contacto con solución salina amortiguada donde se endurecerá en un par de segundos. Sin embargo, por lo general sigue siendo de plástico el tiempo suficiente para ser removido de la punta de la cola capilar o si bien se aplica a la muestra. </li></ol> Semi-intacta Drosophila corazón de la mosca adulta <ol><li> Las moscas adultas de Drosophila se anestesian con siesta Fly (Carolina Biological Supply Co.) por 2-5 minutos. Se debe tener cuidado de no dejar que las moscas de la siesta para volar largas exposiciones. La exposición a corto plazo al frío también puede ser utilizado para anestesiar a las moscas de CO 2, pero no es recomendable ya que tiene más efectos duraderos sobre la frecuencia cardiaca y rítmica. </li><li> Moscas anestesiadas se colocan, cara dorsal hacia abajo, en una placa de Petri revestida con una capa fina de vaselina. La cutícula hidrófoba en las alas y el cuerpo de forma reversible &quot;pegarse&quot; a la gelatina, pero la marcha se puede colocar si es necesario. Esto es especialmente importante durante las manipulaciones posteriores. </li><li> Una reducción inicial se realiza mediante un par de tijeras curvas de primavera (Roboz # 5611). Las cuchillas de tijera se colocan debajo de las piernas en ángulo hacia abajo y hacia la superficie dorsal del tórax cerca del cuello. La cabeza, el cordón nervioso ventral, las piernas se retiran con un solo corte. La preparación se sumerge en un hemolinfa oxigenada, artificial (IA) solución. </li><li> Con la tijera de primavera el extremo posterior del abdomen (compuesto de segmentos abdominales 7 y 8) se elimina con un solo corte. Esto proporciona un acceso lateral para realizar cortes a lo largo de ambos bordes del abdomen y sirve para cortar la conexión entre el intestino posterior y la cutícula abdominal. El colgajo libre de la cutícula ventral de abdomen, luego se retira. </li><li> En los pasos 3 y 4 de las conexiones anteriores y posteriores de la tripa se cortan para los órganos abdominales se encuentran ahora en su lugar gracias tracheols. El intestino y otros órganos abdominales se pueden quitar generalmente como una masa única con las pinzas de joyería (Roboz, # 55) y suave tirón. La eliminación de los órganos internos revela el tubo de corazón que late todavía en la cutícula dorsal rodeada de cuerpos grasos. </li><li> Los cuerpos de grasa son muy opaco en el microscopio de luz en consecuencia a menudo es necesario para eliminar parte de esta grasa con el fin de ver el tubo de corazón claramente. Esto se logra mediante una técnica de liposucción utilizando capilares de vidrio finamente elaborado. Los capilares se hacen usando una pipeta estándar extractor (Por ejemplo, Sutter P-97 con una pipeta asistente de cámara) y se inserta en un tubo de plástico (Tygon, 1 / 16 &quot;) conectado a una fuente de vacío. Este sistema se utiliza para succionar el exceso de grasa alrededor del tubo cardíaco. Fuerza de succión es proporcional al diámetro de la punta para pipetas con puntas superiores a 40 micras de diámetro, no debe ser utilizado. El cuidado extremo se debe tomar para evitar el contacto con el propio corazón y por la parte posterior del corazón es muy frágil que la región debe evitarse por completo. </li><li> El adulto tubo de Drosophila corazón quedará al descubierto y deben ser golpes. Si es necesario el corazón cutícula y adjunto se puede colocar con cuidado, levantando la cutícula de la vaselina y cuidadosamente apisonado de vuelta, de nuevo evitando cualquier contacto con el tejido del corazón. En tsu punto de vista la solución del baño de la preparación debe ser reemplazado por AHL fresco. Esta preparación se debe permitir que se equilibre durante 20-30 minutos con la oxigenación antes de cualquier manipulación. Si la preparación se ha de mantener durante más de 60 minutos que deben ser preservados con AHL fresco. </li></ol> Inmovilización de las larvas de Drosophila para la grabación óptica <ol><li> Coloque una gota de pegamento Histoacryl cerca de cada esquina del cubreobjetos Sylgard. Añadir 50μl de tampón B &amp; B a las gotas de pegamento que se endurece al instante. Llenar el espacio entre los puntos de pegado con B &amp; B - el pegamento se mantenga la memoria intermedia en la diapositiva. </li><li> Tome un vagabundo larva L3 y colocarlo en un pañuelo de papel mojado en hielo durante 2 min. Esto retrasará su movimiento. A continuación, traslado a la larva en el búfer y orientarlo con el dorso hacia arriba (este es el lado de la larva que carece de los cinturones de dentículo). Aplicar rápidamente un poco de pegamento entre la región de la cabeza y el Sylgard. El pegamento se endurece muy rápidamente y evitar que la cabeza de la larva que se mueva. </li><li> El uso de pinzas con cuidado agarra al animal por los espiráculos posteriores y suavemente estirar. Fijar la larva en esta posición mediante la aplicación de la cola a ambos lados de la larva en la mitad posterior. Para reducir aún más el movimiento de la cutícula de los músculos de la pared del cuerpo, agregue más pegamento en los flancos del animal. La larva se ha fijado ahora en esta posición. El cubreobjetos se puede colocar en una pequeña placa de Petri llena de AHL o PBS. Latidos del corazón ahora se puede grabar sin la interferencia de los movimientos de la pared del cuerpo. </li></ol> Los resultados representativos: El semi-intacta corazón adulto de Drosophila se latir rítmicamente durante horas después de la disección cuando se mantiene en el AH fresca y oxigenada (ver datos suplementarios, Ocorr, et al., 2007). Un ejemplo representativo se muestra en la película 1. Corazones desde el 3 de instar de la larva y las moscas jóvenes (1 - 3 semanas después de la eclosión) generalmente presentan un ritmo regular del corazón a un ritmo de entre 1 - 3 Hz. Las moscas más de tres semanas por lo general son más arrítmico, sin embargo, estos corazones son todavía capaces de vencer de forma espontánea durante horas en AHL oxigenada. Corazones en moscas adultas que se encuentren dañados como consecuencia del procedimiento de disección suelen mostrar regiones localizadas de la constricción extrema que no pueden relajarse. Las tasas de latidos del corazón más lento que 1 Hz se observan rara vez en las moscas de menores de 3 semanas de edad y, en consecuencia de estas preparaciones se consideran dañados y se descartan. <a href="http://www.jove.com/files/ftp_upload/1425/MOVIE1.avi">Haga clic aquí para descargar la película 1.</a> - A una semana de vida volar tipo salvaje adulto (cepa de laboratorio w 1118) que muestra el tubo expuesto corazón abdominal (anterior a la izquierda). Tenga en cuenta las contracciones regulares y rítmicos. Zonas opacas en la esquina superior derecha están los restantes órganos de la grasa abdominal, células redondas a cada lado del corazón son las células del pericardio.

<ol>
	<li>Ocorr, K., Reeves, N., Wessells, R. J., Fink, M., Chen, H. -. S. V., Akasaka, T., Yasuda, S., Metzger, J., Giles, W., Posakony, J. W., Bodmer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=KCNQ+potassium+channel+mutations+cause+cardiac+arrhythmias+in+Drosophila+that+mimic+the+effects+of+aging.">KCNQ potassium channel mutations cause cardiac arrhythmias in Drosophila that mimic the effects of aging.</a> Proc Natl Acad Sci U S A. 104, 3943-3948 (2007).</li><li>Broadie, K. S., Bate, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+embryonic+neuromuscular+synapse+of+Drosophila+melanogaster.">Development of the embryonic neuromuscular synapse of Drosophila melanogaster.</a> J Neurosci. 13, 144-166 (1993).</li></ol>

El modelo de Drosophila ha demostrado ser una potente herramienta genética que se ha utilizado para tratar una variedad de problemas científicos que van desde el desarrollo embrionario para el aprendizaje y la memoria. Recientemente, este organismo modelo versátil se ha utilizado para investigar la genética de la función cardíaca en la marcha. Un número de intentos de cuantificar la fisiología del corazón en adultos de Drosophila se han basado en observaciones realizadas en las moscas intacta a través de la cu...

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<td>Micro Dissecting Spring Scissors (curved)</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5611</td>
<td>Good for gross cuts (Step 3)</td>
</tr>
<tr>
<td>Micro Dissecting Spring Scissors (straight)</td>
<td>&nbsp;</td>
<td>Roboz</td>
<td>RS-5620</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #55 forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11295-51</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #5 forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11295-10</td>
<td>&nbsp;</td>
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<td>Glass Capillaries, 100ul</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>53432-921</td>
<td>&nbsp;</td>
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<td>Glass Capillaries, fine</td>
<td>&nbsp;</td>
<td>Science Products</td>
<td>GB100T8P</td>
<td>&nbsp;</td>
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<td>Pipette Puller</td>
<td>&nbsp;</td>
<td>Sutter Equipment</td>
<td>P-97</td>
<td>Both horizontal and vertical pipette pullers will work</td>
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<tr>
<td>Plastic tubing</td>
<td>&nbsp;</td>
<td>Tygon R-3603</td>
<td>1/16&#x201D; inner diameter</td>
<td>for 100&mu;l capillaries</td>
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<td>Plastic tubing</td>
<td>&nbsp;</td>
<td>Tygon R-3603</td>
<td>1/32&#x201D; inner diameter</td>
<td>for small capillaries</td>
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<tr>
<td>Histoacryl&reg; tissue adhesive</td>
<td>&nbsp;</td>
<td>B.Braun, Melsungen</td>
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visualizing the beating heart in drosophila

La<em&gt; Drosophila</emCorazón&gt; ha surgido recientemente como un buen modelo de sistema para el examen de los mecanismos genéticos, celulares y moleculares que subyacen a la función en el corazón de miogénica. Un paso clave en el examen de la función del corazón en la mosca es encontrar una manera de acceder al corazón de una manera que preserva su función miogénica al tiempo que permite el órgano corazón que late a ser observados y registrados. Dos métodos diferentes para observar y registrar los latidos del corazón, tanto en larva y adulto<em&gt; Drosophila</em&gt; A continuación se describen. Nuestros semi-intacta la preparación con las moscas adultas permite una visualización clara del corazón abdominal sin la interferencia de la cutícula pigmentada y que cubre los cuerpos grasos. Para grabar el corazón late larval es necesario inmovilizar a la larva, lo que minimiza los movimientos del cuerpo lo que reduce los movimientos de la pared del corazón que no están asociadas con las contracciones del miocardio. Nuestras metodologías de producción de un adulto estable y preparaciones de larvas del corazón que puede vencer a la hora a un ritmo de 3.1 Hz.

Watch this Scientific Journal Video about Visualizing the Beating Heart in Drosophila at JoVE.com

Visualizing the Beating Heart in Drosophila

Técnica necesaria para la visualización de los latidos del corazón en adultos y larvas<em&gt; Drosophila</em&gt; Se presentan. Cada etapa de la vida requiere de una metodología diferente.

Visualizar el corazón latiendo en Drosophila</em

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

visualizing-the-beating-heart-in-drosophila

Research

JoVE Journal

Biology

Visualizing the Beating Heart in Drosophila

18.6K vistas.  The Sanford Burnham Institute for Medical Research. Técnica necesaria para la visualización de los latidos del corazón en adultos y larvas Drosophila Se presentan. Cada etapa de la vida requiere de una metodología diferente.

Video: Visualizing the Beating Heart in Drosophila

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Visualizar el vivo Drosophila Gliales-unión neuromuscular con tintes fluorescentes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Investigación

Biología

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

Marcaje fluorescente de Drosophila Estructuras del Corazón

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

Monitorización de la función del corazón en las larvas Drosophila melanogaster Para los estudios fisiológicos

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

Preparación de embriones para Microscopía Electrónica de la Drosophila Tubo cardíaco embrionario

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Quantification of &gamma;H2AX Foci in Response to Ionising Radiation

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA lesions with respect to survival and preservation of genomic integrity. An early response to the induction of DSBs is phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX1. Following induction of DSBs, H2AX is rapidly phosphorylated by the phosphatidyl-inosito 3-kinase (PIKK) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Typically, only a few base-pairs (bp) are implicated in a DSB, however, there is significant signal amplification, given the importance of chromatin modifications in DNA damage signalling and repair. Phosphorylation of H2AX mediated predominantly by ATM spreads to adjacent areas of chromatin, affecting approximately 0.03% of total cellular H2AX per DSB2,3. This corresponds to phosphorylation of approximately 2000 H2AX molecules spanning ~2 Mbp regions of chromatin surrounding the site of the DSB and results in the formation of discrete &gamma;H2AX foci which can be easily visualized and quantitated by immunofluorescence microscopy2. The loss of &gamma;H2AX at DSB reflects repair, however, there is some controversy as to what defines complete repair of DSBs; it has been proposed that rejoining of both strands of DNA is adequate however, it has also been suggested that re-instatement of the original chromatin state of compaction is necessary4-8. The disappearence of &gamma;H2AX involves at least in part, dephosphorylation by phosphatases, phosphatase 2A and phosphatase 4C5,6. Further, removal of &gamma;H2AX by redistribution involving histone exchange with H2A.Z has been implicated7,8. Importantly, the quantitative analysis of &gamma;H2AX foci has led to a wide range of applications in medical and nuclear research. Here, we demonstrate the most commonly used immunofluorescence method for evaluation of initial DNA damage by detection and quantitation of &gamma;H2AX foci in &gamma;-irradiated adherent human keratinocytes9.

Quantification of &amp;#947;H2AX Foci in Response to Ionising Radiation

Quantification of DNA double-strand streaks using &gamma;H2AX formation as a molecular marker has become an invaluable tool in radiation biology. Here we demonstrate the use of an immunofluorescence assay for quantification of &gamma;H2AX foci after exposure of cells to radiation.

La cuantificación de γH2AX focos en respuesta a la radiación ionizante

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA ...

Quantitation of &gamma;H2AX Foci in Tissue Samples

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) processes or by exogenous factors, particularly ionizing radiation and radiomimetic compounds. Phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX, is an early response to DNA double-strand breaks1. This phosphorylation event is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Overall, DSB induction results in the formation of discrete nuclear &gamma;H2AX foci which can be easily detected and quantitated by immunofluorescence microscopy2. Given the unique specificity and sensitivity of this marker, analysis of &gamma;H2AX foci has led to a wide range of applications in biomedical research, particularly in radiation biology and nuclear medicine. The quantitation of &gamma;H2AX foci has been most widely investigated in cell culture systems in the context of ionizing radiation-induced DSBs. Apart from cellular radiosensitivity, immunofluorescence based assays have also been used to evaluate the efficacy of radiation-modifying compounds. In addition, &gamma;H2AX has been used as a molecular marker to examine the efficacy of various DSB-inducing compounds and is recently being heralded as important marker of ageing and disease, particularly cancer3. Further, immunofluorescence-based methods have been adapted to suit detection and quantitation of &gamma;H2AX foci ex vivo and in vivo4,5. Here, we demonstrate a typical immunofluorescence method for detection and quantitation of &gamma;H2AX foci in mouse tissues.

Quantitation of &amp;#947;H2AX Foci in Tissue Samples

Quantitation of DNA double-strand breaks on the basis of &gamma;H2AX foci has become an invaluable tool, particularly in radiation biology, for the evaluation of tissue radiosensitivity and effects of radiation modifying compounds. Here we demonstrate the use of an immunofluorescence assay for quantitation of &gamma;H2AX foci in tissue samples.

La cuantificación de γH2AX focos en muestras de tejido

DNA double-strand breaks (DSBs) are particularly lethal and genotoxic lesions, that can arise either by endogenous (physiological or pathological) ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &amp;#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

Evaluación de la distribución espacial de la radiación ionizante γH2AX siguientes

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Clonogenic Assay: Adherent Cells

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. Apart from documenting the method, the initial landmark study generated the first radiation-dose response curve for X-ray irradiated mammalian (HeLa) cells in culture1. Basically, the clonogenic assay enables an assessment of the differences in reproductive viability (capacity of cells to produce progeny; i.e. a single cell to form a colony of 50 or more cells) between control untreated cells and cells that have undergone various treatments such as exposure to ionising radiation, various chemical compounds (e.g. cytotoxic agents) or in other cases genetic manipulation. The assay has become the most widely accepted technique in radiation biology and has been widely used for evaluating the radiation sensitivity of different cell lines. Further, the clonogenic assay is commonly used for monitoring the efficacy of radiation modifying compounds and for determining the effects of cytotoxic agents and other anti-cancer therapeutics on colony forming ability, in different cell lines. A typical clonogenic survival experiment using adherent cells lines involves three distinct components, 1) treatment of the cell monolayer in tissue culture flasks, 2) preparation of single cell suspensions and plating an appropriate number of cells in petri dishes and 3) fixing and staining colonies following a relevant incubation period, which could range from 1-3 weeks, depending on the cell line. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cell lines with the use of an immortalized human keratinocyte cell line (FEP-1811)2. Also, our aims are to describe common features of clonogenic assays including calculation of the plating efficiency and survival fractions after exposure of cells to radiation, and to exemplify modification of radiation-response with the use of a natural antioxidant formulation.

The applicability of the clonogenic assay for evaluating reproductive viability has been established for more than 50 years. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cells.

Ensayo clonigénicas: las células adherentes

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

Imágenes de Apoptotic Liquidación de células vivas durante<em&gt; Drosophila</em&gt; Embriogénesis

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

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 ...