Este protocolo se utiliza actualmente en nuestro laboratorio y ha sido perfeccionado a lo largo de los años por muchas personas como los doctores David Sharp, Kwon Mijung, Sommi Patrizia, Cheerambathur Dhanya. Se agradece al Dr. Bill Sullivan (UCSC), que nos ha proporcionado excelentes consejos sobre la manipulación y la microinyección de los primeros embriones de Drosophila, cuando nuestro trabajo en este sistema se está iniciando (ref. 13). Agradecemos a todos los miembros del laboratorio Scholey. Nuestro trabajo en la mitosis en Drosophila es apoyado por el NIH subvención GM55507.

Recetas: Placas de jugo de uvas: <ul><li> 5,5 g de agar Bacto </li><li> 14,5 g de dextrosa o glucosa </li><li> 7,15 g de sacarosa </li><li> 45 ml de jugo concentrado de uva (jugo 100%). </li><li> 204.5 ml H 2 0 </li><li> 625 l de NaOH 10N </li></ul> Mezcle todos los ingredientes y horno de microondas hasta que hierva. Añadir la mezcla 2,8 ml de ácido (mezcla de ácidos: 20,9 ml de ácido propiónico, 2,1 ml de ácido fosfórico, 27 ml H 2 0) Mezclar y verter en placas de Petri de 35 mm. Deje que solidifique a temperatura ambiente durante uno o dos días. Si las placas no se va a utilizar muy pronto, sellar con parafilm y se mantenga a 4 ° C (hasta que se estabilice a RT antes de usar). Levadura pasta: Disuelva una cucharadita de levadura (Sigma YSC2, la levadura Saccharomyces cerevisiae de tipo II) en agua para formar una pasta espesa. Coloque una pequeña cantidad de esta en cada placa de jugo de uva antes de su uso. G-PEM buffer: <ul><li> 80 mM Na-TUBOS, pH 6,9 </li><li> MgCl2 1 mM </li><li> 1 mM EGTA </li><li> 1 mM GTP </li></ul> Inyección de tampón: <ul><li> 150 mM K-aspartato </li><li> 10 mM K-fosfato </li><li> 20 mM imidazol, pH 7,2 </li></ul> Heptano cola: Desenrollar cinta adhesiva de doble pegajosa y colocar en un frasco de 100 ml, agregar aproximadamente 50 ml de heptano, el sello de la botella, y el rock durante varios días. Cuando se utiliza, agrega heptano si la cola es demasiado grueso. Cámara de deshidratación: Tome una cápsula de Petri de 100 mm, poner una parte de un plato de 35 mm en el interior para hacer una &quot;mesa&quot; y añadir Drierite (sulfato de calcio anhidro) a su alrededor para que la altura de Drierite no es mayor que la &quot;mesa&quot; y la tapa. El cubreobjetos con embriones se colocan en esta &quot;mesa&quot; para la deshidratación antes de la inyección. Es una buena idea usar al menos algunas Drierite indicando el cambio y que cuando se ha cambiado de color. Protocolo: Este protocolo se puede utilizar para la inyección de casi cualquier reactivo soluble en el embrión de Drosophila sincitial: Por ejemplo, ya sea una proteína fluorescente para la observación o un inhibidor de la proteína diana o ambas cosas. Preparando todo: 1 .- 2 a 3 días antes del experimento, hacer las placas de zumo de uva y establecer laicos jaula: Tome una botella de plástico (6 oz, de fondo redondo de Drosophila Científica Aplicada), corte dos agujeros pequeños (de 1 cm 2) en los lados opuestos de la botella y rellenar con un trozo de algodón (esto permitirá que el aire circule), toque las moscas en la botella y la tapa con una placa de jugo de uva. Mantener a 25 ° C (o temperatura ambiente), las moscas deben establecer los embriones de unos 10 días. 2 .- Por lo menos un día antes del experimento de sacar las agujas para la inyección, se utiliza una aguja Kopf / pipeta modelo de extractor de 720 y delgados filamentos de vidrio de paredes (World Precision Instruments tw100f). Para la inyección de la tubulina fluorescente, mantenga agujas a 4 ° C para evitar que la temperatura inducida por polimerización en la aguja. Tubulina preparación: Nota: Todos los trabajos con la tubulina se debe hacer a 4 ° C para prevenir la polimerización. <ol><li> Tome tubulina marcado fuera del congelador y ponerla en hielo durante 5 a 15 minutos. </li><li> Diluir con el G-PEM buffer (4 a 10 veces dependiendo de la intensidad requerida). </li><li> Se centrifuga a 13k rpm a 4 ° C durante 10-15 minutos. </li><li> Carga 1 a 2 l en la aguja. </li></ol> Anticuerpos o inhibidores químicos: Prepare agujas como para la tubulina, utilizando una concentración adecuada (esto puede ser necesario para ser probado). El anticuerpo puede ser en el buffer de la inyección, el G-PEM buffer o PBS. Si se necesita otro buffer, el buffer tendrá que ser inyectado como un control para asegurarse de que el buffer no causa ningún daño por sí mismo. Nota: Inhibidor de la tubulina y fluorescentes se pueden mezclar e inyectar juntos si las concentraciones lo permiten. De lo contrario, inyectar la tubulina fluorescente y esperar de 5 a 10 minutos antes de la inyección del inhibidor. Cuando se realiza una doble inyección, empieza con más embriones, ya que menos embriones se recuperará con normalidad. De recogida de embriones: <ol><li> Ponga un plato de jugo de uva nuevo en la jaula estaba. </li><li> Eliminar esta placa después de una hora, esta es la primera colección del día y muchas veces no es muy buena. Puede ser descartada. </li><li> Cambiar la placa de cada hora para mantener la recogida de embriones durante una hora cada uno. </li></ol> Cubreobjetos de preparación: <ol><li> Ponga un cubreobjetos de 50 x 22 mm en un lado de un portaobjetos de microscopio. Cinta de las cuatro esquinas de la diapositiva para que no se mueve. </li><li> El uso de un aplicador con punta de algodón, poner una capa de pegamento heptano en una sola línea sobre el cubreobjetos (la cola no debe ser viscoso y se seca en pocos segundos, de lo contrario añadir más heptano). </li><li> Ponga un pedazo dedoble cinta adhesiva en la diapositiva hacia el lado del cubreobjetos. </li></ol> Embrión de la preparación: Nota: Los embriones deben ser reflejados de aproximadamente 2 horas después del inicio de la recolección, a fin de comenzar los siguientes pasos con tiempo suficiente para acabar con ellos dentro de este plazo. <ol><li> Con una brocha humedecida cuidadosamente recoger los embriones de la placa de jugo de uva y el lugar en una cinta adhesiva doble en la diapositiva. </li><li> Utilizando la parte exterior de las pinzas, rollo de los embriones en la cinta adhesiva de doble hasta el corion (la membrana externa) se rompe. </li><li> Levante el embrión suavemente rodando sobre el corion por lo que se pega a la pinza y colocarlo en la cola de heptano en el cubreobjetos con el lado largo del paralelo de embriones para el lado largo del cubreobjetos. Conjunto entre un 10 y 20 embriones en una fila. </li><li> Quitar el cubreobjetos y colocarlo en la cámara de deshidratación de 3 a 8 minutos (esto depende de la humedad de la habitación y la cantidad a inyectar). </li><li> Colocar en una cámara de metal con grasa de vacío. </li><li> Cubrir los embriones con 700 halocarbonos aceite para evitar una mayor deshidratación. Los embriones están listos para la inyección. </li></ol> Embrión de inyección: <ol><li> Buscar los embriones en un objetivo de 16x. </li><li> Mueva los embriones de distancia, y encontrar la aguja sin moverse del plano focal. </li><li> Centro de la aguja y moverla hacia arriba sin moverlo en dirección xo y.. </li><li> Si la aguja no está abierto, puesto al borde del cubreobjetos en el campo de visión, pero no cuando la aguja se va, y bajar la aguja con el mismo plano focal. Con mucho cuidado mover el cubreobjetos hasta que llega la aguja y con cuidado se rompe abierto. Si la aguja está abierto, vaya al paso 5. (Las agujas se pueden abrir con el ácido fluorhídrico antes del llenado). </li><li> Ponga los embriones a la vista (pero no en donde la aguja se), bajar la aguja en el aceite y asegúrese de obtener gotas de buen líquido de la aguja. </li><li> Con cuidado pero sin pausa mover el embrión en la aguja e inyectar una gota en el embrión y mover el embrión de distancia. (O siga las instrucciones de su aparato de inyección). </li><li> Después de todos los embriones se han inyectado, están listos para su observación en un microscopio confocal. </li></ol> Imagen: Imagen de los embriones en un microscopio confocal con un objetivo de 60 o 100 veces: Notas: <ol><li> El intervalo de tiempo en una serie lapso de tiempo debe ser como máximo de unos segundos, ya que la mitosis en el embrión es muy rápido, según el proceso exacto para ser estudiados. </li><li> Una serie en 3D o un solo plano, se pueden adquirir en función del proceso de interés. </li><li> De acción rápida inhibidores de la transferencia del microscopio de inyección de microscopio de imagen tiene que ser rápido. </li></ol><img src="/files/ftp_upload/1382/1382fig1.jpg" alt="la figura 1" /> Figura 1: Estudio de la mitosis en el embrión de Drosophila sincitial. Núcleos en el embrión temprano sufren divisiones rápida sin citocinesis, durante los ciclos de 10 a 13 núcleos forman una monocapa en la corteza. A. Esquemática del embrión temprano, con núcleos en la corteza. Embrión puede expresar las buenas prácticas agrarias y / o Solicitud de Propuestas proteínas marcadas y se pueden microinyección con otras proteínas marcadas y / o inhibidores. B. Tiempo de imágenes a intervalos de embriones que expresan GFP-tubulina y el PP-histonas. Parcela C de la separación de Polo-Polo de ciclos de 11 , 12 y 13 en un embrión de tipo silvestre. Longitud de husillo presenta períodos de duración isométrica y los períodos de elongación, que son muy consistentes y reproducibles. D. Dinámica de los microtúbulos del huso. La inyección de baja concentración de tubulina rodamina lleva a manchas formadas por una subunidades de tubulina pocos, que puede ser usado para estudiar la dinámica de los microtúbulos.

<ol>
	<li>Brust-Mascher, I., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitotic+spindle+dynamics+in+Drosophila.">Mitotic spindle dynamics in Drosophila.</a> Int Rev Cytol. 259, 139-172 (2007).</li><li>Duffy, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GAL4+system+in+Drosophila:+a+fly+geneticist's+Swiss+army+knife.">GAL4 system in Drosophila: a fly geneticist's Swiss army knife.</a> Genesis. 34, 1-15 (2002).</li><li>Morris, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+methods+for+analyzing+the+functions+of+kinesins+in+early+embryos.">Microinjection methods for analyzing the functions of kinesins in early embryos.</a> Methods Mol Biol. 164, 163-172 (2001).</li><li>Brust-Mascher, I., Sommi, P., Cheerambathur, D. K., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin-5-dependent+poleward+flux+and+spindle+length+control+in+Drosophila+embryo+mitosis.">Kinesin-5-dependent poleward flux and spindle length control in Drosophila embryo mitosis.</a> Mol Biol Cell. 20, 1749-1762 (2009).</li><li>Brust-Mascher, I., Civelekoglu-Scholey, G., Kwon, M., Mogilner, A., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+for+anaphase+B:+role+of+three+mitotic+motors+in+a+switch+from+poleward+flux+to+spindle+elongation.">Model for anaphase B: role of three mitotic motors in a switch from poleward flux to spindle elongation.</a> Proc Natl Acad Sci U S A. 101, 15938-15943 (2004).</li><li>Brust-Mascher, I., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubule+flux+and+sliding+in+mitotic+spindles+of+Drosophila+embryos.">Microtubule flux and sliding in mitotic spindles of Drosophila embryos.</a> Mol Biol Cell. 13, 3967-3975 (2002).</li><li>Cheerambathur, D. K., Brust-Mascher, I., Civelekoglu-Scholey, G., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+partitioning+of+mitotic+kinesin-5+crosslinkers+between+microtubule-bound+and+freely+diffusing+states.">Dynamic partitioning of mitotic kinesin-5 crosslinkers between microtubule-bound and freely diffusing states.</a> J Cell Biol. 182, 421-428 (2008).</li><li>Cheerambathur, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+analysis+of+an+anaphase+B+switch:+predicted+role+for+a+microtubule+catastrophe+gradient.">Quantitative analysis of an anaphase B switch: predicted role for a microtubule catastrophe gradient.</a> J Cell Biol. 177, 995-1004 (2007).</li><li>Kwon, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chromokinesin,+KLP3A,+dives+mitotic+spindle+pole+separation+during+prometaphase+and+anaphase+and+facilitates+chromatid+motility.">The chromokinesin, KLP3A, dives mitotic spindle pole separation during prometaphase and anaphase and facilitates chromatid motility.</a> Mol Biol Cell. 15, 219-233 (2004).</li><li>Sharp, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+coordination+of+three+mitotic+motors+in+Drosophila+embryos.">Functional coordination of three mitotic motors in Drosophila embryos.</a> Mol Biol Cell. 11, 241-253 (2000).</li><li>Sharp, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+bipolar+kinesin,+KLP61F,+cross-links+microtubules+within+interpolar+microtubule+bundles+of+Drosophila+embryonic+mitotic+spindles.">The bipolar kinesin, KLP61F, cross-links microtubules within interpolar microtubule bundles of Drosophila embryonic mitotic spindles.</a> J Cell Biol. 144, 125-138 (1999).</li><li>Sharp, D. J., Rogers, G. C., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoplasmic+dynein+is+required+for+poleward+chromosome+movement+during+mitosis+in+Drosophila+embryos.">Cytoplasmic dynein is required for poleward chromosome movement during mitosis in Drosophila embryos.</a> Nat Cell Biol. 2, 922-930 (2000).</li><li>Sharp, D. J., Yu, K. R., Sisson, J. C., Sullivan, W., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antagonistic+microtubule-sliding+motors+position+mitotic+centrosomes+in+Drosophila+early+embryos.">Antagonistic microtubule-sliding motors position mitotic centrosomes in Drosophila early embryos.</a> Nat Cell Biol. 1, 51-54 (1999).</li><li>Waterman-Storer, C. M., Desai, A., Bulinski, J. C., Salmon, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+speckle+microscopy,+a+method+to+visualize+the+dynamics+of+protein+assemblies+in+living+cells.">Fluorescent speckle microscopy, a method to visualize the dynamics of protein assemblies in living cells.</a> Curr Biol. 8, 1227-1230 (1998).</li></ol>

Este protocolo es relativamente sencillo, sin embargo, cada paso requiere de práctica para asegurarse de que los embriones no están dañados. Cuidadosos experimentos de control siempre se necesita hacer para asegurar que los resultados se están obteniendo fiables. Una buena manera de comenzar es mediante la microinyección de tubulina buffer o rodamina en embriones de control que expresan GFP-tubulina para asegurarse de que la mitosis procede normalmente a través de todos los ciclos en la superficie del embrión (ci...

microinjection techniques for studying mitosis in the drosophila melanogaster syncytial embryo

Este protocolo describe el uso del embrión de Drosophila melanogaster sincitial para el estudio de la mitosis 1. Drosophila tiene una genética útil con un genoma secuenciado, y puede ser fácil de mantener y manipular. Muchos mutantes de mitosis existen, y las moscas transgénicas que expresan funcionales fluorescente (GFP, por ejemplo) - etiquetado proteínas mitótico han estado y están generando. Expresión de genes diana es posible utilizar el sistema GAL4/UAS 2. El embrión de Drosophila temprana lleva a cabo múltiples mitosis muy rápidamente (la duración del ciclo celular, ≈ 10 min). Es muy adecuado para obtener imágenes de la mitosis, ya que durante los ciclos de 10-13, los núcleos se dividen rápidamente y de forma sincrónica sin intervenir citocinesis en la superficie del embrión en una sola monocapa justo debajo de la corteza cerebral. Estos núcleos se dividen rápidamente, probablemente el uso de la maquinaria mitótica igual que otras células, pero que están optimizados para la velocidad, el puesto de control general, se cree que no se estrictas, lo que permite el estudio de las proteínas mitótico cuya ausencia podría causar detención del ciclo celular en células con un control fuerte . Los embriones que expresan proteínas GFP etiquetados o microinyección con la etiqueta fluorescente proteínas pueden ser fácilmente imágenes para seguir la dinámica de vida (Fig. 1). Además, los embriones pueden ser microinyección con el bloqueo de la función-anticuerpos o inhibidores de proteínas específicas para estudiar el efecto de la pérdida o la perturbación de su función 3. Estos reactivos se difunden en todo el embrión, llegando a muchos ejes para producir un gradiente de concentración del inhibidor, lo que a su vez se traduce en un gradiente de defectos comparable a una serie de alelos de mutantes. Idealmente, si la proteína diana es marcado con fluorescencia, el gradiente de inhibición se puede visualizar directamente 4. Se supone que el más fuerte fenotipo es comparable con el fenotipo nulo, a pesar de que es difícil excluir formalmente la posibilidad de que los anticuerpos pueden tener efectos dominantes en raras ocasiones, los controles tan rigurosos y una prudente interpretación debe ser aplicada. Más lejos del sitio de la inyección, la función de proteínas es sólo parcialmente perdido permitir otras funciones de la proteína diana a hacerse evidente.

Watch this Scientific Journal Video about Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo at JoVE.com

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Este protocolo describe el uso de la microinyección y la imagen de alta resolución en el<em&gt; Drosophila melanogaster</em&gt; Embrión sincitial para estudiar mitosis.

Técnicas de microinyección para el estudio de la mitosis en el Drosophila melanogaster Embrión sincitial

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

microinjection-techniques-for-studying-mitosis-drosophila

Research

JoVE Journal

Biology

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

14.0K vistas.  University of California, Davis. Este protocolo describe el uso de la microinyección y la imagen de alta resolución en el Drosophila melanogaster Embrión sincitial para estudiar mitosis.

Video: Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

La disección del sistema nervioso central de larvas de Drosophila melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Investigación

Biología

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Vídeo de alta resolución de Seguimiento de la locomoción en adultos de Drosophila melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Registro electrofisiológico en el Drosophila Embrión

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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  ...

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 ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

El estudio de puesto de control mitótico, ilustrando comportamiento dinámico de proteínas cinetocoro y el movimiento de cromosomas en vivo<em&gt; Drosophila</em&gt; Los embriones Sincitial

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Medición de la esperanza de vida en<em&gt; Drosophila melanogaster</em

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

Imaging A través de la envoltura pupal de<em&gt; Drosophila melanogaster</em

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular biology has been greatly fueled by the discovery of noncoding RNAs as major players in post-transcriptional regulation. Such a revolutionary perspective has been accompanied and triggered by the development of powerful technologies for profiling short RNAs expression, both at the high-throughput level (genome-wide identification) or as single-candidate analysis (steady state accumulation of specific species). Although several state-of-art strategies are currently available for dosing or visualizing such fleeing molecules, Northern Blot assay remains the eligible approach in molecular biology for immediate and accurate evaluation of RNA expression. It represents a first step toward the application of more sophisticated, costly technologies and, in many cases, remains a preferential method to easily gain insights into RNA biology. Here we overview an efficient protocol (Enhanced Northern Blot) for detecting weakly expressed microRNAs (or other small regulatory RNA species) from Drosophila melanogaster whole embryos, manually dissected larval/adult tissues or in vitro cultured cells. A very limited amount of RNA is required and the use of material from flow cytometry-isolated cells can be also envisaged.

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The aim of this publication is to visualize and discuss the operative steps of an Enhanced Northern Blot protocol on RNA extracted from Drosophila melanogaster embryos, cells, and tissues. This protocol is particularly useful for the efficient detection of small RNA species.

Mejorada del Norte Blot Detección de los Pequeños ARN Especies en<em&gt; Drosophila melanogaster</em

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular ...

Generation of Transgenic Hydra by Embryo Microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Hydra is used as a model for the study of regeneration, pattern formation, and stem cells. However, research efforts have been hampered by lack of a reliable method for gene perturbations to study molecular function. The development of transgenic methods has revitalized the study of Hydra biology1. Transgenic Hydra allow for the tracking of live cells, sorting to yield pure cell populations for biochemical analysis, manipulation of gene function by knockdown and over-expression, and analysis of promoter function. Plasmid DNA injected into early stage embryos randomly integrates into the genome early in development. This results in hatchlings that express transgenes in patches of tissue in one or more of the three lineages (ectodermal epithelial, endodermal epithelial, or interstitial). The success rate of obtaining a hatchling with transgenic tissue is between 10% and 20%. Asexual propagation of the transgenic hatchling is used to establish a uniformly transgenic line in a particular lineage. Generating transgenic Hydra is surprisingly simple and robust, and here we describe a protocol that can be easily implemented at low cost.

Generation of Transgenic Hydra by Embryo Microinjection

Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi.