KB es apoyado por el NIH subvención GM54544.

Parte 1: Equipos y Suministros <ol><li> Un microscopio de disección se requiere una buena disección de embriones, se sugiere la ampliación de 40X, 25X, con los ojos las piezas para aumentar al máximo aumento. </li><li> Pinzas finas (número 5) se requieren para la selección manual de los embriones y devitellinization de embriones. </li><li> Equipo para realizar y modificar las finas agujas de vidrio es necesario. Tira de vidrio se requieren agujas para la disección. Preferimos vidrio sólido para la disección (duran más), pero la norma de paredes gruesas tubo de vidrio (diámetro externo 1-1,5 mm) funciona tan bien. Algunas personas utilizan agujas muy finas de tungsteno, electrólisis de la nitidez deseada en un baño de NaOH 1 M con una batería de 10 + autotransformador Amp. Afiladas agujas de disección de tungsteno tienen la ventaja de que son relativamente resistentes y de larga duración. Agujas de vidrio hueco (similar a la forma de parches electrodos pinza) están conectados a un tubo de plástico y se utiliza durante la disección de succión y expulsión de solución salina, así como la entrega controlada de la cola. Una variedad de extractores de vidrio están disponibles para la fabricación de agujas de cristal. Programada por ordenador extractores puede ser programado para tirar de una variedad de formas y tamaños. El Brown-Flaming modelos (Sutter Instrument Co.) son ampliamente favorecidos. Un microscopio compuesto (20-40X objetivo) debe estar disponible para inspeccionar y modificar los electrodos antes de su uso. </li><li> Dos tipos de soluciones se utilizan comúnmente para baño de los tejidos embrionarios durante la disección: 1) &quot;estándar&quot; (Jan y Jan, 1976a) o &quot;norma modificada&quot; (Broadie, 2000) salinas, en base a soluciones de uso común para la grabación en otros sistemas de invertebrados , y 2) &quot;hemolinfa-like&quot; (HL) Salines (Stewart et al 1994), un compromiso entre la solución salina normal y las concentraciones iónicas medidas en la hemolinfa de Drosophila. Cabe señalar que ninguna de estas salinas de reproducir fielmente la composición química de la hemolinfa en vivo (Broadie, 2000). Salino estándar contiene (en mM): 135 NaCl, 5 KCl, 4 MgCl 2, 1,8 CaCl 2, 72 sacarosa, 5 TES, pH 7,2. Los experimentadores también puede considerar salina preparada comercialmente de insectos (por ejemplo, los medios de comunicación de insectos de Schneider), que no es apta para registro electrofisiológico pero lo más probable para proteger la salud de los tejidos expuestos. </li></ol> Parte 2: La estadificación de embriones y la disección <ol><li> Para la recolección de huevos óptima, mantener jóvenes (&lt;7 días), varones sanos y las moscas hembras (20-40) en fresco por la que se ollas durante 2-3 días. Por la que se ollas comunes constan de 100 vasos de plástico ml (Tri-Pour) perforada para proporcionar la circulación de aire adecuada, cubriendo una placa de agar de 60 mm. Placas de agar contienen jugo de manzana o de uva endurecido con agar. Existen varias recetas, pero un ejemplo es el siguiente: 700 ml de agua, 25 a 30 gramos de agar, 300 ml de jugo concentrado (uva o manzana), 0,5 g de metilparabeno (p-hydroxymethylbenzoate), y el azúcar 30 g. Autoclave el agua y el agar, y por separado hervir el p-hydroxymethylbenzoate con el azúcar y el jugo. Mezcle el jugo de agar y soluciones de forma rápida y verter en placas de 60 mm, la eliminación de las burbujas. </li><li> Antes de la recolección de huevos, las moscas se alimentan de pasta de levadura (7 gramos de levadura de panadería en 9 ml de agua se almacenan a 4 ° C) en las placas nuevas, cambiar por lo menos dos veces al día durante al menos dos días. El día 3, una buena olla debe producir 100-200 huevos por hora. Mejor puesta de huevos se observa en una olla mantenido a su lado con el agar en la placa de rayado. Muchos embriones se establecerán en o junto a los arañazos. </li><li> Para recoger un gran número de embriones, la transferencia de los embriones con cuidado de la placa de agar en una cesta con un pincel. Una cesta se puede hacer con un tubo de centrífuga de 15 ó 50 ml. Cortar la parte inferior dejando la superficie lo suficiente para atrapar a una pantalla (70 micras de malla) entre la tapa y la parte superior del tubo. Enjuague los embriones con dH 2 0 y poner en una placa de Petri de agua dulce 50% al 100% de lejía para eliminar el corion externo. Por otra parte, los huevos pueden ser recogidos por separado utilizando unas pinzas finas, y dechorionated mediante la colocación de forma manual en una gota de lejía. De-chorionation puede tardar desde 30 segundos a 2 minutos (lejía fresca es mucho más rápido), y deben ser controlados bajo el microscopio de disección. La eliminación del corion expone la membrana brillante, transparente vitelino. Huevos dechorionated debe enjuagarse brevemente, o se colocan en una solución salina, como el cloro destruye rápidamente los tejidos devitellinized. </li><li> Es fundamental para la etapa cuidadosamente los embriones a la edad deseada de desarrollo. La Drosophila definido las fases embrionarias (1-17; Campos-Ortega y Hartenstein, 1985) no son útiles, como todo el desarrollo neuromuscular funcional ocurre en 16 etapas finales o de la etapa 17. Por lo tanto, los embriones son protagonizadas por hora de desarrollo a 25 ° C en una incubadora humidificada, en horas después de la fecundación o, más comúnmente, después de la puesta de huevos (AEL). En estas condiciones, la embriogénesis dura 21 + / - 1 hora a 25 ° C. </li><li> Los huevos procedentes de un tiempo de huevo laicos (1-2 hrs) que estén debidamente años están al lado de vista de los criterios morfológicos. Después de un prolongado lavado en dH 2 O, los huevos dechorionated se colocan en una placa de cultivo de plástico (en dH 2 O) para su visualización. Puesta en escena correcta se puede confirmar con criterios morfológicos utilizando la luz reflejada con un microscopio de disección (Campos-Ortega y Hartenstein, 1985). Madura la última etapa de embriones (22-24h AEL) al borde de la eclosión son generalmente reconocidos por la segmentación de la cutícula y la tráquea infladas. Los embriones también pueden ser genotipo utilizando equilibradores de las buenas prácticas agrarias y un microscopio de disección de fluorescencia con los filtros adecuados. </li><li> Para la disección, embriones y dechorionated etapas de desarrollo están sujetos (lado dorsal hacia arriba) en un cubreobjetos en una gota de solución salina (véase el paso 1.5). Los embriones se extraen de la membrana vitelina por punción de la membrana cerca del extremo anterior o posterior, con una micropipeta de vidrio o una aguja de tungsteno, y luego suavemente liberar al embrión de la membrana. En preparación para la disección, el embrión debe ser colocado en el portaobjetos con la superficie hasta dorsal (parte dorsal hasta la disección se exponga el sistema nervioso ventral y neuromusculatura para los experimentos. Sin embargo, el embrión, obviamente, se puede colocar en otras maneras de facilitar el acceso a otros tejidos.) </li><li> Embriones prematuros (&lt;16 h AEL) se conectan directamente a limpiar el cristal o de vidrio recubierto con polilisina (poli-L-lisina bromhidrato; Broadie y Bate, 1993a, b). Mayores embriones (&gt; 16 hrs AEL), tras la formación de la cutícula, deben ser pegados, lo ideal es que Sylgard recubiertos (Dow Corning) cubreobjetos. Un agua de polimerización pegamento de cianoacrilato quirúrgico o veterinario se utiliza (Broadie y Bate, 1993c, d). El pegamento se entrega a través de una pipeta de vidrio pequeño electrodo (10-20 punta micrómetro de diámetro interior) conectada a un tubo de goma con el flujo de cola controlado por la presión de la boca. Se debe tener cuidado durante el parto de cola, como el pegamento se polimeriza rápidamente en contacto con la solución salina. Tenga en cuenta también que pegar no se puede realizar en una solución salina sin o bajas concentraciones de iones divalentes, como el pegamento requiere iones divalentes para polimerizar. Pequeñas cantidades de pegamento se utilizó por primera vez para sujetar firmemente la cabeza y la cola. </li><li> Una vez que los extremos del embrión se pegan a los cubreobjetos, se hace una incisión a lo largo de la línea media dorsal con un electrodo de vidrio o una aguja de tungsteno afilada. Esto es más fácil con cuidado perforando la línea de incisión antes de hacer el corte final. Los lados de la incisión se adjuntarán a los cubreobjetos con más cola, para extender suavemente el piso embrión. Los órganos internos, incluyendo los intestinos, los órganos de la grasa y, opcionalmente, la tráquea se retirar mediante aspiración con una pipeta de vidrio conectado a un tubo de goma (Broadie y Bate 1993a-c). El embrión debe adjuntar planos de la cubreobjetos, la epidermis hacia abajo con el ventral del sistema nervioso central, nervios periféricos y músculos somáticos expuestos para la experimentación. </li></ol>

Clasificación precisa de los embriones es crítica debido a la rápida maduración de las propiedades funcionales en el transcurso de tiempo de tan sólo varias horas. Varios problemas complican la puesta en escena. En primer lugar, la mayoría de los investigadores utilizan huevo tiempo pone a los embriones de etapa, pero el tiempo de puesta de huevos puede variar enormemente de un animal a otro en diferentes condiciones (Broadie et al. 1992). En particular, las mujeres con una dieta limitada tienden a retener los hue...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
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<tr>
<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
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harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

Drosophila es un modelo genético de primera para el estudio de ambos el desarrollo embrionario y funcional neurociencia. Tradicionalmente, estos campos son muy aislados unos de otros, con historias en gran medida independiente y la comunidad científica. Sin embargo, la relación entre estos campos por lo general diferentes programas de desarrollo es la base de adquisición de las propiedades funcionales de señalización eléctrica y la diferenciación de las sinapsis químicas funcional durante las fases finales de la formación de circuitos neuronales. Esta interfaz es una zona muy importante para la investigación. En Drosophila, estas fases de desarrollo funcional se producen durante un período de &lt;8 horas (a 25 ° C) durante el último tercio de la embriogénesis. Este período de desarrollo tardío fue considerado durante mucho tiempo difíciles de investigación debido a la deposición de una cutícula dura, epidérmico impermeable. Un avance avance fue la aplicación de agua de polimerización pegamento quirúrgico que puede aplicarse localmente a la cutícula para permitir la disección controlada de la última etapa de embriones. Con una incisión longitudinal dorsal, el embrión puede ser en plano, dejando al descubierto el cordón nervioso ventral y la musculatura de la pared del cuerpo para la investigación experimental. Este sistema ha sido muy utilizada para aislar y caracterizar mutantes genéticos que impiden la formación de sinapsis embrionario, por lo que revelan los mecanismos moleculares que regulan la especificación y diferenciación de las sinapsis y conexiones sinápticas propiedades funcionales de señalización.

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Esta técnica expone al neuromusculatura Drosophila embrionarias para inmunohistoquímica o de registro electrofisiológico. Es útil para el estudio de los primeros eventos en el desarrollo neuromuscular o la realización de electrofisiología en los mutantes que no pueden eclosionar.

La recolección y preparación de Drosophila Los embriones de registro electrofisiológico y otros procedimientos

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

harvesting-preparing-drosophila-embryos-for-electrophysiological

Research

JoVE Journal

Biology

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K vistas.  University of Illinois. Esta técnica expone al neuromusculatura Drosophila embrionarias para inmunohistoquímica o de registro electrofisiológico. Es útil para el estudio de los primeros eventos en el desarrollo neuromuscular o la realización de electrofisiología en los mutantes que no pueden eclosionar.

Video: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

Preparación de cultivos neuronales a partir de embriones de Drosophila Midgastrula Etapa

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Investigación

Biología

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

Los métodos electrofisiológicos para la grabación de los potenciales sinápticos de la UNM de larvas de Drosophila

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

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

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 ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Imagen de la posición vertical Drosophila Los embriones

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

Estrategia de transfección optimizados para la expresión y el registro electrofisiológico de los canales iónicos recombinantes del voltaje en las células HEK-293T

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Aislamiento y purificación de Kinesin desde Drosophila Los embriones

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Preparación Individual Drosophila Huevos Cámaras de Imágenes en directo

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

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