Agradecemos a Polan Santos y Darren Obbard para las ilustraciones de Drosophila en la Fig. 2a, Marcelo Jacobs-Lorena para el uso del insectario JHMRI. Este trabajo fue apoyado por la beca de la Fundación de Investigación de Ciencias de la Vida (HLT), Comité de Becas de la Universidad de Hong Kong AOE / B-07/99 (MCF), y el NIH subvenciones NS096677, NS037402 y NS083373 (JMH). Ho Lam Tang es una Fundación Shurl y Kay Curci miembro de la Fundación de Investigación de Ciencias de la Vida.

1. Preparación de las moscas CaspaseTracker <ol><li> Para preparar CaspaseTracker (DQVD) vuela de experimentos, realice esta cruz: UBI-CaspaseTracker x G-TRACE (UAS-RFP; UAS-FLP; Ubi&gt; Stop&gt; GFP-nls), mediante la transferencia de 7-10 hembra virgen (o macho) las moscas que lleva el sustrato de la caspasa-biosensor mCD8 DIAP1-Gal4 impulsada por el promotor de ubiquitina 35 junto con el mismo número de hombres (o mujeres) moscas G-TRACE, que tienen el segundo cromosoma equilibrador CyO para evitar la letalidad de la combinación homocigótica del G- TRACE (UAS-RFP, UAS-FLP, y Ubi&gt; Stop&gt; GFP-nls) 34. Lugar vuela en un vial de alimentos frescos con pasta de levadura fresca como fuente de proteínas. </li><li> Incubar los archivos a 18 ° C durante 5 a 7 días (máximo 2 semanas) y luego retire el padre vuela desde el vial para evitar el hacinamiento con la nueva progenie, y la creación de nuevos viales de cría. </li><li> Continuar la incubación a 18 ° C hasta que la progenie vuela eclose. Seleccione la no CyO [no-cURLy ala] progenie del genotipo correcto de CaspaseTracker, que son transgénicos para los elementos CaspaseTracker y G-35 TRACE. </li><li> Al mismo tiempo, generar control parental W118 moscas no transgénicas y la caspasa-insensibles (DQVA) vuela en paralelo para verificar RFP CaspaseTracker específica y la fluorescencia de GFP. </li><li> Realizar el genotipado por PCR para confirmar los archivos con CaspaseTracker utilizando los cebadores: 5&#39;-TCCCCCGGGCTGCAGGAATTC, 3&#39;-TGGAATTGGGGTACGTCTAGA, producir un producto de 3897 pb. Para la genotipificación los loci G-Trace, utilice los siguientes cebadores para GFP (474 ​​pb), 5&#39;-GAC CAC TTC TTC AAG TCC GCC ATG CCC G, 3&#39;-CTT GTA CAG CTC GTC CAT GAG GCC GAT AGT C; para la solicitud de propuestas (258 pb), 5&#39;-GGC TGC TTC ATC TAC AAG AAG GTG TTC ATC GG, 3&#39;-GAT GTC GTC CAG CTT GGA CAC GTA GTA GTA GC; y para Flpase (655 pb), 5&#39;-CCACCTAAGGTGCTTGTTCGTCAGTTTGTGG, 3&#39;-CCG TAC TAA CGC TTG TTG TCT TCT CTG TCA C. Para la genotipificación de Gal4 en el vector pUWR (554 pb), 5&#39;-GAA GCA CAC CTT CGC ATC TCG CAGTCA CGC, 3&#39;-TGG AAT TGG GGT ACG TCT AGA. </li></ol> 2. Preparación del tejido, tinción y montaje <ol><li> Para las disecciones de moscas, evitar que los tejidos disecados se peguen a las puntas de pipeta de plástico y tubos de centrífuga, puntas de la capa y tubos con albúmina de suero bovino al 1% (BSA) disueltos en agua. </li><li> Dissect vuela sobre un cojín de silicona con fórceps. <ol><li> Para evitar daños en la punta de pinzas sobre una superficie dura cuando se realiza la disección del tejido, realice la disección en una superficie de silicio. Para la fabricación de placas de disección de silicona, fundir el silicio en el kit comercial de acuerdo con las instrucciones del fabricante (Lista de Materiales). Después de fundir la silicona, añadir de 3 a 4 ml a una placa de cultivo tisular de diámetro 60 mm. platos de silicio pueden ser reutilizados. </li><li> Anestesiar a una mosca con CO 2, y la transfiere a una placa de silicona con 1 ml de PBS frío. Introducir el CO 2 a un vial que contiene la marcha usando una pistola de aire, y luego transferir la mosca t anestesiadasOA cojín que tiene un suministro de CO2. </li><li> Use un par de pinzas para sujetar la cabeza de la mosca, y un segundo par de pinzas para tirar del tórax en la dirección opuesta para desconectar la cabeza mosca del cuerpo que cubre sin dañar la conexión entre la cabeza y el intestino anterior. </li><li> Use 2 pares de pinzas para quitar las alas y las patas del tórax. </li><li> Use un par de pinzas para sostener el tórax, y otro par de pinzas para tirar del abdomen para separarlo del tórax de nuevo sin dañar la conexión entre el intestino anterior y del intestino medio. </li><li> Diseccionar los órganos internos, incluyendo cerebro, intestino, tubos de Malpighi y los ovarios como se ha demostrado anteriormente. 39-41 </li><li> Retire todas las piezas de la cutícula y los cuerpos grasos con unas pinzas, ya que estos tejidos producen fuertes autofluorescencia. La paciencia y la práctica se requieren para preparar un sistema de órgano completo, como se muestra. </li></ol></li><li> Transferir los tejidos disecados a 1,5 mL tubos de centrífuga, y fijar tejidos wiTH 0,5 ml de paraformaldehído al 4% en PBS (solución salina tamponada con fosfato) a TA durante 20 a 30 minutos con rotación en la oscuridad para evitar RFP blanqueo y GFP en los tejidos. Evitar la fijación prolongada que puede comprometer los resultados de tinción posteriores. </li><li> Retire el paraformaldehído por pipeteo suave y lavar 3 veces con 0,5 ml de PBST (PBS + 0,1% de Triton X-100) a TA. </li><li> Permeabilizar los tejidos con 0,5 ml de PBST a 4 ° C durante la noche con agitación suave. incubaciones cortas pueden causar permeabilización y el compromiso de tinción resultados incompletos. <ol><li> Si es necesario para reducir la tinción de fondo, considere la incubación de los tejidos en 0,5 ml de PBST con 1% de BSA, en lugar de PBS. </li></ol></li><li> Lavar los tejidos 3 veces con 0,5 ml de PBST. </li><li> Para la tinción de los núcleos y la actina filamentosa (F-actina), aplicar 0,5 ml de PBST con 10 mg / ml de Hoechst 33342 tinte nuclear azul y 0,3 M Alexa Fluor mancha 633 faloidina F-actina y se incuba de forma simultánea con los tejidos durante 1 h a TA wITH rotación suave en la oscuridad. Evitar las manchas prolongado ya que esto aumentará el fondo. <ol><li> Para la inmunotinción, incubar fija y permeabilizadas tejidos con 0,1% Triton X-100 en 0,5 ml de PBS durante la noche a 4 ° C, tinción con dilución 1: 100 de anticuerpo anti-ELAV o con dilución 1: 200 de anti-caspasa-3 durante la noche a 4 ° C (300 l). Siga por el correspondiente anticuerpo secundario diluido 1: 100 y se incuba durante 1-3 h a TA. Ver Lista de Materiales para obtener información anticuerpo específico. </li></ol></li><li> Lavar los tejidos 3 veces, cada una durante 5 minutos en 0,5 ml de PBST con rotación suave. </li><li> Con una pipeta, retire todo el PBST, y luego añadir 200 l anti-blanqueador de un medio de montaje (ver materiales) para cubrir completamente los tejidos durante 1-3 horas a temperatura ambiente, o 4 ° C durante la noche. tejidos óptimas que han absorbido plenamente el agente de montaje se hundirá hasta el fondo del tubo. </li><li> Pre-limpiar el portaobjetos de vidrio con agua o etanol al 70% y transferir los tejidos con el agente de montaje a los glass de diapositivas. </li><li> Con cuidado, coloque una hoja de cubierta de vidrio sobre los tejidos. Aplique vaselina en el portaobjetos de vidrio y la hoja de la cubierta para evitar la destrucción del tejido por compresión excesiva. Eliminar los agentes de montaje adicional con papel de seda. </li><li> Sellar la hoja de la cubierta mediante la aplicación de esmalte de uñas a lo largo de los bordes de la hoja de la cubierta para evitar la fuga de agentes de montaje de los tejidos. </li></ol> 3. Microscopía Confocal <ol><li> Utilizar un microscopio de epi-fluorescencia confocal invertido. Sin embargo, microscopios arriba-derecha también se pueden utilizar. Para tejidos de imagen utilizando suelo de baldosas, mantener estable a temperatura ambiente para evitar la deriva de enfoque y el desplazamiento de plano xy debido a la expansión y contracción térmica de los componentes. sistemas de control ambiental y sistemas de compensación de deriva enfoque ayudan a evitar este problema debido a la pérdida de termo-equilibrio del sistema de microscopio. </li><li> Colocar el portaobjetos en la platina del microscopio. Tome algunas imágenes de prueba usando baja resolución (125 x 125 píxeles) de unad determinar el número adecuado de azulejos necesarios para cubrir toda la región de interés (ROI). </li><li> Establecer el sistema de barrido microscopio para capturar imágenes con una resolución de escaneo tales como 500 x 500 píxeles. Seleccione el objetivo tales como 20X NA 0,8 o un 1,4 NA objetivo 63X Plan-Apocromático para el análisis de tejidos a través de cubreobjetos de vidrio, de manera que cada una de las imágenes (azulejos) se solapa en un 20% en todos los lados. </li><li> Establecer la secuencia de imágenes de largo al más corto longitudes de onda de excitación, como la luz de longitud de onda larga provoca menor de foto-blanqueo. La secuencia del más largo al más corto de longitud de onda de excitación es de: (i) 633 nm para faloidina F-actina y para el anticuerpo a caspasas activadas, (ii) 561 nm para RFP, (iii) 488 nm para GFP, y (iv) 405 nm para Hoechst tinción nuclear. </li><li> Durante estas pruebas, minimizar la exposición de las células a excitación láser fluorescente durante el proceso de obtención de imágenes para evitar la foto-blanqueo mediante la reducción de la intensidad del láser para obtener imágenes de alta calidad de los tejidos. </li><li> Despues de que yomagos se recogen, fusionar las imágenes utilizando el software de microscopio. </li></ol>

The authors have nothing to disclose.

A continuación se ilustra la construcción y el funcionamiento interno de CaspaseTracker que facilitan la detección de la actividad de caspasa basal generalizada en los tejidos sanos. Los pasos críticos para la detección de la actividad de caspasa no apoptótica in vivo son: 1) la generación de moscas con el transgén biosensor, 2) verificar la función reportero-caspasa específico con controles apropiados, 3) la práctica de técnicas de disección para observar todos los sistemas de órganos internos de...

Las caspasas son proteasas de cisteína que median la muerte celular por apoptosis mediante la escisión de muchas proteínas intracelulares después de residuos de aspartato clave. Por ejemplo, caspasas iniciadoras activan las caspasas efectoras, nucleasas de ADN derepress, componentes del citoesqueleto escinden y alteran la composición de lípidos de las membranas celulares para desmantelar rápidamente las células y estimular su reconocimiento y la inmersión por las células que disponen de los cadáveres célula vecina. 1-4 Se estima que miles de millones de células mueren por día en el cuerpo humano, y la apoptosis es un mecanismo importante de la muerte de células tumorales inducida por la quimioterapia. 5 un conjunto diferente de las caspasas puede causar la muerte celular por procesos no apoptóticas distintas para estimular la inmunidad innata. 6 por lo tanto, mayoría de las investigaciones sobre las caspasas se ha centrado en sus funciones a favor de la muerte. Curiosamente, las primeras pruebas en el campo reveló que los mismos caspasas responsables de promover la muerte de las células también tienen no la muerte funciones. Estudios pioneros han demostrado que las caspasas están implicadas en diversas funciones celulares en las células sanas, incluyendo la regulación de la proliferación celular y la migración durante la embriogénesis. 7-9 se requieren caspasas para la individualización spermatid en Drosophila 10,11, para el bloqueo de una vía de la muerte celular necroptotic alternativa en ratones 12,13, y para el procesamiento de microARN en C. . elegans 14,15 En quizás los más longevos células, las neuronas, las caspasas y otra maquinaria apoptótica están implicados en la regulación de la actividad neuronal mediante la poda de terminaciones sinápticas, un proceso que se cree que es esencial para el fortalecimiento de otras sinapsis para el aprendizaje y la memoria 16-. 18 es posible que las caspasas facilitar la poda sináptica por un tipo de mini-apoptosis de las proyecciones neuronales diminutas sin muerte de células enteras. 19 sin embargo, las caspasas pueden tener funciones alternativas no relacionadas con eventos de apoptosis similares. 20,21 papel Duals en la vida y la muerte no son exclusivos de las caspasas; BCL-2 y proteínas de la familia del citocromo c tienen papeles en la energética celular en las células sanas, pero también son parte de la ruta apoptótica núcleo que se activa por muchos tipos de estrés celular. 22-25 Aunque no está comprobado, parece lógico que la evolución ha vinculado día -Jobs hasta la muerte, puestos de trabajo en las mismas moléculas para asegurar la eliminación oportuna de las células no aptas o no deseables. En la actualidad, los mecanismos moleculares de la actividad de caspasa no apoptótico no se entienden, y el grado de actividad de la caspasa no apoptótico durante el desarrollo embrionario y en los tejidos adultos también no se conoce. Un reto importante es la dificultad de distinguir día-trabajos desde death-puestos de trabajo de las caspasas. En contraste con la apoptosis y pyroptosis, cuando la actividad de caspasa se amplifica por una cascada proteolítica, se espera que los trabajos diurnos de las caspasas se produzca a niveles mucho más bajos de actividad enzimática, probablemente por debajo de la detección por muchos tecn disponiblesgías. Antes del trabajo presentado aquí, otros desarrollaron una variedad de biosensores de caspasa para diferentes propósitos. Los biosensores de SCAT (por ejemplo, ECFP-DEVD-Venus) detectar rápidamente la actividad de caspasa en tiempo real en células cultivadas y tejidos animales utilizando FRET. 26,27 Tras la escisión de la caspasa, la fracción GFP dirigido al núcleo de Apoliner (mCD8-RFP-DQVD- nucGFP) sufre relocalización subcelular dentro de minutos cuando su membrana de sujeción plasma es escindido por las caspasas. 28 del mismo modo, ApoAlert-pCaspase3-sensor (NES-DEVD-YFP-NLS) relocaliza desde el citosol al núcleo tras la escisión de la caspasa. 29,30 Más recientemente, el cromóforo en iCasper fue diseñado inteligentemente a emitir fluorescencia cuando se escinde por caspasas, lo que permite la detección de la actividad de biosensor en tiempo real en las neuronas de embriones de Drosophila, pero principalmente en asociación con la muerte celular de desarrollo. muerte 31 dependiente de caspasa de las neuronas olfativas durante el envejecimiento estaba demonstclasificado por inmuno-detección de la forma de la caspasa-escindido de biosensores CPV (por ejemplo, mCD8-PARP-Venus). 32,33 Es importante destacar que, se detectó la forma activada de la caspasa-3 en ausencia de muerte celular por inmunotinción sensible en espinas de neuronas cultivadas, y en el soma usando la fluorescencia dependiente de caspasa del colorante indicador CellEvent nuclear, pero se encontraron dificultades debido a la foto-toxicidad, aunque la muerte celular se retrasa hasta después de la eliminación columna vertebral. 19 por lo tanto, se necesitan nuevos biosensores caspasas para detectar y realizar un seguimiento de las células con actividad de caspasa basal in vivo. Para superar estas dificultades, hemos generado un nuevo biosensor de color caspasa doble, designado CaspaseTracker. Esta estrategia combina una versión modificada del biosensor Apoliner Drosophila caspasa-sensible 28 con el sistema FRT recombinasa Drosophila G-TRACE 34 para etiquetar y rastrear células en vivo de forma permanente. &lt;sup&gt; 35 El sistema G-TRACE activado-Gal4 permite que los niveles muy bajos de caspasas para activar CaspaseTracker, lo que resulta en la expresión de RFP en el citoplasma y la expresión de GFP dirigido al núcleo permanente en cualquier célula que nunca ha experimentado la actividad de caspasa. 35 Este sistema puede etiquetar las células durante toda la vida en animales enteros utilizando Drosophila melanogaster, un sistema modelo tratable y ampliamente utilizado para el estudio de las caspasas y la muerte celular. 36-38

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			<td height="21" style="height:21px;width:273px;">CONSUMABLES AND REAGENTS</td>
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			<td style="width:225px;">H-1000</td>
			<td>Mounting medium</td>
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			<td>Dumont tweezer biology grade, stainless steel</td>
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			<td height="21" style="height:21px;">Hanging Drop Slides</td>
			<td style="width:229px;">Fisher Scientific</td>
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			<td>Glass slides</td>
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			<td style="width:225px;">Antibody Registry ID:&#160; AB_528218&#160;</td>
			<td>Stain for Drosophla pan-neuronal ELAV</td>
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			<td height="21" style="height:21px;width:273px;">Cleaved caspase-3 (Asp175) antibody</td>
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			<td>Stain for active fragment of caspase-3</td>
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			<td>to preserve fluorophores&#160;</td>
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			<td>to preserve fluorophores&#160;</td>
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			<td style="width:229px;">Dow Corning&#160;</td>
			<td style="width:225px;">Product code: 0001023934</td>
			<td>for dissection plates</td>
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			<td height="21" style="height:21px;width:273px;">EQUIPMENT</td>
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			<td height="21" style="height:21px;">LSM780 confocal microscope</td>
			<td style="width:229px;">Carl Zeiss</td>
			<td style="width:225px;">N/A</td>
			<td>Imaging</td>
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			<td height="21" style="height:21px;width:273px;">Carl Zeiss Stereomicroscope Stemi 2000&#160;</td>
			<td style="width:229px;">Carl Zeiss</td>
			<td style="width:225px;">N/A</td>
			<td>Drosophila dissection</td>
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			<td height="42" style="height:42px;width:273px;">AmScope Fiber Optic Dual Gooseneck Microscope Illuminator, 150W</td>
			<td style="width:229px;">AmScope</td>
			<td style="width:225px;">WBM99316&#160;</td>
			<td>Light source</td>
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in vivo biosensor tracks non-apoptotic caspase activity in drosophila

Caspases are the key mediators of apoptotic cell death via their proteolytic activity. When caspases are activated in cells to levels detectable by available technologies, apoptosis is generally assumed to occur shortly thereafter. Caspases can cleave many functional and structural components to cause rapid and complete cell destruction within a few minutes. However, accumulating evidence indicates that in normal healthy cells the same caspases have other functions, presumably at lower enzymatic levels. Studies of non-apoptotic caspase activity have been hampered by difficulties with detecting low levels of caspase activity and with tracking ultimate cell fate in vivo. Here, we illustrate the use of an ultrasensitive caspase reporter, CaspaseTracker, which permanently labels cells that have experienced caspase activity in whole animals. This in vivo dual color CaspaseTracker biosensor for Drosophila melanogaster transiently expresses red fluorescent protein (RFP) to indicate recent or on-going caspase activity, and permanently expresses green fluorescent protein (GFP) in cells that have experienced caspase activity at any time in the past yet did not die. Importantly, this caspase-dependent in vivo biosensor readily reveals the presence of non-apoptotic caspase activity in the tissues of organ systems throughout the adult fly. This is demonstrated using whole mount dissections of individual flies to detect biosensor activity in healthy cells throughout the brain, gut, malpighian tubules, cardia, ovary ducts and other tissues. CaspaseTracker detects non-apoptotic caspase activity in long-lived cells, as biosensor activity is detected in adult neurons and in other tissues at least 10 days after caspase activation. This biosensor serves as an important tool to uncover the roles and molecular mechanisms of non-apoptotic caspase activity in live animals.

Existen dos componentes clave que permiten CaspaseTracker para detectar la actividad de la caspasa en las células sanas normales (Figura 1A). El primero de ellos es un polipéptido de la caspasa-escindible 146 amino ácido el modelo de la caspasa biosensor Apoliner (Figura 1b). 28 Este polipéptido se deriva de DIAP1 (inhibidor de Drosophila de apoptosis) que contiene un único sitio de la caspasa natural que se escinde durante la ap...

Watch this Scientific Journal Video about In Vivo Biosensor Tracks Non-apoptotic Caspase Activity in Drosophila at JoVE.com

In Vivo Biosensor Tracks Non-apoptotic Caspase Activity in Drosophila

Para detectar las células sanas en animales enteros que contienen niveles bajos de actividad caspasa, el biosensor altamente sensible designado CaspaseTracker se generó para Drosophila. se detecta actividad biosensor dependiente de caspasa en las células sanas de larga vida a través de los órganos internos de los animales adultos criados en condiciones optimizadas en ausencia de estímulos de muerte.

En Vivo biosensor Pistas de apoptosis no caspasa actividad en Drosophila

Caspases are the key mediators of apoptotic cell death via their proteolytic activity. When caspases are activated in cells to levels detectable by ...

in-vivo-biosensor-tracks-non-apoptotic-caspase-activity-in-drosophila

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JoVE Journal

Biology

In Vivo Biosensor Tracks Non-apoptotic Caspase Activity in Drosophila

9.0K vistas.  Johns Hopkins University Bloomberg School of Public Health.  Para detectar las células sanas en animales enteros que contienen niveles bajos de actividad caspasa, el biosensor altamente sensible designado CaspaseTracker se generó para Drosophila. se detecta actividad biosensor dependiente de caspasa en las células sanas de larga vida a través de los órganos internos de los animales adultos criados en condiciones optimizadas en ausencia de estímulos de muerte. 

Video: In Vivo Biosensor Tracks Non-apoptotic Caspase Activity in Drosophila

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Mesoscópica fluorescencia Tomografía por En vivo Imágenes de desarrollo Drosophila</em

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Investigación

Biología

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 ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

En vivo-La centrifugación de embriones de Drosophila

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

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

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

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

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

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

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

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

Patch clamp y técnicas de perfusión para el estudio de los canales iónicos expresados ​​en Xenopus Ovocitos

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

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 ...

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  ...

An in vivo Crosslinking Approach to Isolate Protein Complexes From Drosophila Embryos

Many cellular processes are controlled by multisubunit protein complexes. Frequently these complexes form transiently and require native environment to assemble. Therefore, to identify these functional protein complexes, it is important to stabilize them in vivo before cell lysis and subsequent purification. Here we describe a method used to isolate large bona fide protein complexes from Drosophila embryos. This method is based on embryo permeabilization and stabilization of the complexes inside the embryos by in vivo crosslinking using a low concentration of formaldehyde, which can easily cross the cell membrane. Subsequently, the protein complex of interest is immunopurified followed by gel purification and analyzed by mass spectrometry. We illustrate this method using purification of a Tudor protein complex, which is essential for germline development. Tudor is a large protein, which contains multiple Tudor domains - small modules that interact with methylated arginines or lysines of target proteins. This method can be adapted for isolation of native protein complexes from different organisms and tissues.

An in vivo Crosslinking Approach to Isolate Protein Complexes From Drosophila Embryos

Multi-component protein complexes play crucial roles during cellular function and development. Here we describe a method used to isolate native protein complexes from Drosophila embryos after in vivo crosslinking followed by purification of the crosslinked complexes for subsequent structure-function analysis.

Un In vivo Enfoque de reticulación para aislar Complejos de Proteína Desde Drosophila Los embriones

Many cellular processes are controlled by multisubunit protein complexes. Frequently these complexes form transiently and require native environment to ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

Reconstitución funcional y la actividad del canal Las mediciones de Purificada Wildtype y mutante CFTR Proteína

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

La medición de las larvas La actividad en el<em&gt; Drosophila</em&gt; Monitor de Actividad

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...