KB wird durch NIH GM54544 unterstützt.

Teil 1: Ausrüstung und Zubehör <ol><li> Eine gute Dissektionsmikroskop ist für den Embryo Dissektionen erforderlich; 40-facher Vergrößerung wird vorgeschlagen, mit 25X Okulare bis maximal die Vergrößerung zu erhöhen. </li><li> Feine Pinzette (Nummer 5) sind für die manuelle Selektion von Embryonen und devitellinization von Embryonen erforderlich. </li><li> Ausrüstung zu machen und zu ändern gutes Glas Nadeln erforderlich ist. Gezogen-Glas-Nadeln sind für die Präparation erforderlich. Wir bevorzugen massivem Glas für die Präparation (länger), aber Standard dickwandigen Glasröhren (Außendurchmesser 1-1,5 mm) arbeitet als gut. Einige Leute benutzen feinen Wolfram-Nadeln, elektrolysiert, um die gewünschte Schärfe in ein Bad von 1M NaOH mit einem 10 + Amp Spartransformator Batterie. Geschärft Wolfram Dissektion Nadeln haben den Vorteil, dass sie relativ robust und langlebig sind. Hohlglas Nadeln (gebildet ähnlich Klemmelektroden Patch) sind Kunststoffschlauch befestigt und verwendet bei der Präparation für Saug-und Ausweisung von Kochsalzlösung, sowie die kontrollierte Abgabe von Leim. Eine Vielzahl von Glas-Abzieher sind für die Herstellung von Glas Nadeln. Computer-Programmierung Abzieher kann voreingestellt werden, um eine Vielzahl von Formen und Größen zu ziehen. Die Brown-Flaming Modelle (Sutter Instrument Co.) sind weit verbreitet begünstigt. Ein zusammengesetztes Mikroskop (20-40fach Objektiv) zur Verfügung stehen sollte untersuchen und verändern Elektroden vor dem Gebrauch. </li><li> Zwei Arten von Lösungen werden häufig verwendet, um die Bad embryonalen Gewebe während der Präparation: 1) &quot;Standard&quot; (Jan und Jan, 1976a) oder &quot;modifizierten Standardtarif&quot; (Broadie, 2000) Salinen, basierend auf Lösungen üblicherweise für die Aufnahme in andere wirbellose eingesetzt und 2) &quot;Hämolymphe-like&quot; (HL) Salinen (Stewart et al 1994), einen Kompromiss zwischen den Standard-Kochsalzlösung und die Ionenkonzentrationen in der Drosophila Hämolymphe gemessen. Es wird darauf hingewiesen, dass keines dieser Salinen genau imitieren die chemische Zusammensetzung der Hämolymphe in vivo (Broadie, 2000). Standard-Kochsalzlösung enthält (in mM): 135 NaCl, 5 KCl, 4 MgCl 2, 1,8 CaCl 2, 72 Saccharose, 5 TES, pH 7,2. Experimentatoren möchten vielleicht auch kommerziell hergestellte Insekten Kochsalzlösung (z. B. Schneider Insect Media), die ungeeignet für die elektrophysiologische Ableitung aber wahrscheinlich für die Gesundheit der exponierten Gewebe zu schützen, ist zu prüfen. </li></ol> Teil 2: Embryo Staging und Dissection <ol><li> Für eine optimale Eiersammlung, halten jung (&lt;7 Tage), gesunde männliche und weibliche Fliegen (20-40) auf frischen Verlegung Töpfe für 2-3 Tage. Legen Töpfen häufig bestehen aus 100 ml Kunststoffbecher (Tri-Pour) perforiert, um eine ausreichende Luftzirkulation, für eine 60 mm-Agar-Platte geben. Agar-Platten enthalten Apfel-oder Traubensaft mit Agar gehärtet. Einige Rezepte gibt, aber ein Beispiel ist wie folgt: 700 ml Wasser, 25-30 g Agar, 300 ml Saft-Konzentrat (Trauben-oder Apfel), 0,5 g Methylparaben (p-hydroxymethylbenzoate) und 30g Zucker. Autoclave das Wasser und Agar-Agar, und separat kochen p-hydroxymethylbenzoate mit dem Zucker und Saft. Mix der Agar-und Saft-Lösungen und schnell in 60 mm-Platten zu gießen, Entfernung von Blasen. </li><li> Vor Entnahme der Eizellen, sind Fliegen mit Hefepaste (7 Gramm Bäckerhefe in 9 ml Wasser bei 4 ° C gelagert) auf frische Platten, mindestens zweimal täglich für mindestens zwei Tage gewechselt zugeführt. Am Tag 3, sollte ein guter Topf produzieren 100-200 Eier pro Stunde. Bessere Eiablage befindet sich auf einem Topf auf die Seite mit dem Agar in eine Metallplatte geritzt gepflegt beobachtet. Viele Embryonen werden in gelegt werden oder neben dem Kratzer. </li><li> Zum Sammeln einer großen Anzahl von Embryonen, sanft Transfer der Embryonen von der Agarplatte in einen Korb mit einem Pinsel. Ein Korb kann mit einem 15 oder 50 ml Zentrifugenröhrchen erfolgen. Schneiden Sie den unteren, wobei genügend Fläche, um einen Bildschirm (70 um Maschenweite) zwischen dem Deckel und dem oberen Ende des Röhrchens Falle. Spülen Sie die Embryonen mit dH 2 0 und in einer Petrischale mit frischem 50% bis 100% Bleichmittel auf die äußere Chorion zu entfernen. Alternativ können Eier einzeln gesammelt werden mit feinen Pinzetten und dechorionated, indem man sie manuell in einen Tropfen Bleichmittel. De-chorionation kann von 30 Sekunden bis 2 Minuten (frische Bleiche ist viel schneller) zu nehmen, und sollte unter der Dissektionsmikroskop überwacht werden. Die Entfernung des Chorion macht die glänzenden, transparent Dotterhaut. Dechorionated Eier sollten kurz abgespült werden, oder die in Kochsalzlösung, als Bleichmittel schnell zerstört devitellinized Gewebe. </li><li> Es ist wichtig, sorgfältig zu Embryonen, um die gewünschte Entwicklungsalter. Die definierten Drosophila Embryonalstadien (1-17; Campos-Ortega und Hartenstein, 1985) sind nicht sinnvoll, da alle funktionellen neuromuskulären Entwicklung tritt im späten Stadium 16 oder Stufe 17. So Embryonen werden von Entwicklungs-Stunden inszeniert bei 25 ° C in einem befeuchteten Inkubator, in Stunden nach der Befruchtung oder, häufiger, nach der Eiablage (AEL). Unter diesen Bedingungen dauert der Embryogenese 21 + / - 1 Stunde bei 25 ° C. </li><li> Eier von einer zeitlich Ei-lay (1-2 Std.s), die entsprechend gealtert sind als nächstes für morphologischen Kriterien angesehen. Nach gründlichem Waschen in dH 2 O, sind die dechorionated Eier in einer Kunststoff-Petrischale (unter dH 2 O) für die Anzeige platziert. Richtig Inszenierung kann mit morphologischen Kriterien mit reflektierte Licht mit einem Dissektionsmikroskop (Campos-Ortega und Hartenstein, 1985) bestätigt werden. Ältere im Spätstadium Embryonen (22-24h AEL) am Rande der Schlupf in der Regel durch die Segmentierung der Kutikula und aufgeblasen Luftröhre anerkannt. Embryonen können auch genotypisiert mit GFP-Balancer und ein Fluoreszenz-Dissektionsmikroskop mit geeigneten Filtern werden. </li><li> Für Dissektion sind dechorionated und entwicklungspolitisch inszeniert Embryonen beigefügt (Rückenseite nach oben) auf einem Deckglas unter einem Tropfen Kochsalzlösung (siehe Schritt 1.5). Die Embryonen werden von der Dotterhaut durch Punktion der Membran in der Nähe der vorderen oder hinteren Ende mit einem Glas-Mikropipette oder Wolfram Nadel, und dann vorsichtig die Befreiung der Embryo aus der Membran entfernt. In Vorbereitung für die Präparation, sollte der Embryo auf dem Deckglas mit der dorsalen Fläche nach oben (dorsal bis Dissektion wird das ventrale Nervensystem und neuromusculature für Experimente aussetzen positioniert werden. Doch der Embryo kann natürlich auch auf andere Weise positioniert werden, um Zugang zu anderen zu erleichtern Gewebe.) </li><li> Embryonen im Frühstadium (&lt;16 Stunden AEL) direkt an Glas oder Glas mit Polylysin (Poly-L-Lysin-Hydrobromid; Broadie und Bate, 1993a, b) beschichtet reinigen. Ältere Embryonen (&gt; 16 Stunden AEL), folgende Nagelhaut Bildung muss geklebt werden, ideal für Sylgard-beschichtet (Dow Corning) Deckgläser. Ein Wasser-Polymerisation von Cyanacrylat chirurgische oder tierärztliche Klebstoff verwendet wird (Broadie und Bate, 1993c, d). Der Leim wird durch einen kleinen Glaselektrode Pipette (10-20 Mikrometer Innendurchmesser Spitze) an einen Gummischlauch mit dem Leimfluss durch den Mund Druck gesteuert ausgeliefert. Es muss während der Leim Lieferung genommen werden, da der Kleber polymerisiert schnell bei Kontakt mit der Salzlösung. Beachten Sie auch, dass Kleben nicht in Kochsalzlösung mit keinen oder geringen zweiwertigen Ionen-Konzentrationen durchgeführt werden, wie der Klebstoff erfordert zweiwertige Ionen zu polymerisieren. Kleine Mengen von Klebstoff werden zuerst verwendet werden, um fest an den Kopf und Schwanz. </li><li> Nachdem die Enden der Embryo in die Deckglas geklebt wird ein Schnitt entlang der dorsalen Mittellinie mit einer Glaselektrode oder geschärft Wolframnadel gemacht. Dies wird dadurch erleichtert, indem Sie vorsichtig Perforieren der Schnittführung vor dem endgültigen Schnitt. Die Seiten des Einschnitts werden dann auf das Deckglas mit mehr geklebt, sanft verteilen Embryo flach. Die inneren Organe einschließlich des Darms, Fettkörper und, optional, Luftröhre werden dann durch Absaugen mittels einer Glaspipette an Gummischlauch (Broadie und Bate 1993a-c) entfernt. Der Embryo ist jetzt angebracht flach auf dem Deckglas werden, Epidermis sich mit der Bauchseite des zentralen Nervensystems, der peripheren Nerven und somatische Muskulatur für Experimente ausgesetzt. </li></ol>

Präzise Inszenierung von Embryonen ist kritisch aufgrund der schnellen Reifung der funktionellen Eigenschaften über den zeitlichen Verlauf von nur einigen Stunden. Mehrere Probleme erschweren diese Inszenierung. Erstens verwenden die meisten Forscher timed Eier legt, um Embryonen, sondern die Eiablage Zeit enorm variieren von Tier zu Tier unter verschiedenen Bedingungen (Broadie et al. 1992). Insbesondere neigen Frauen auf eine begrenzte Diät, um befruchtete Eier über einen längeren Zeitraum vor der Verlegung zu be...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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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 ist ein führender genetisches Modell für die Untersuchung der beiden embryonalen Entwicklung und funktionelle Neurowissenschaften. Traditionell sind diese Felder ganz von einander getrennt, mit weitgehend unabhängigen Geschichten und Wissenschaft. Allerdings ist die Schnittstelle zwischen diesen in der Regel unterschiedliche Felder der Entwicklungsprogramme zugrunde liegenden Erwerb von funktionellen elektrischen Signal-Eigenschaften und die Differenzierung von funktionellen chemischen Synapsen während der letzten Phasen der neuronalen Schaltkreis Bildung. Diese Schnittstelle ist ein äußerst wichtiges Gebiet für die Untersuchung. In Drosophila treten diese Phasen der funktionalen Entwicklung über einen Zeitraum von &lt;8 Stunden (bei ​​25 ° C) während des letzten Drittels der Embryogenese. Diese späte Entwicklungsphase galt lange Zeit als unlösbar zu Ermittlungen wegen der Ablagerung von einem harten, undurchlässigen epidermale Kutikula. Ein Durchbruch Fortschritt war die Anwendung von Wasser-Polymerisation chirurgische Kleber, die lokal an der Nagelhaut kann angewendet werden, um kontrollierte Präparation des Late-Stage-Embryonen zu ermöglichen. Mit einem dorsalen Längsschnitt, kann der Embryo flach gelegt werden, Freilegung der Bauchmark und Körper Wand Muskulatur experimentelle Untersuchung. Dieses System wurde stark genutzt zu isolieren und zu charakterisieren genetische Mutanten, die embryonale Synapsenbildung beeinträchtigen und damit zeigen, die molekularen Mechanismen, über die Spezifikation und Differenzierung von Synapsen-Verbindungen und funktionelle synaptische Signalisierung Eigenschaften.

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

Diese Technik macht die Drosophila embryonale neuromusculature für die Immunhistochemie oder elektrophysiologische Ableitung. Es ist für das Studium frühen Ereignisse in neuromuskulären Entwicklung oder Durchführung Elektrophysiologie in Mutanten, die nicht brüten können nützlich.

Ernte und Vorbereitung Drosophila Embryonen für die elektrophysiologische Ableitung und andere Verfahren

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 Aufrufe.  University of Illinois. Diese Technik macht die Drosophila embryonale neuromusculature für die Immunhistochemie oder elektrophysiologische Ableitung. Es ist für das Studium frühen Ereignisse in neuromuskulären Entwicklung oder Durchführung Elektrophysiologie in Mutanten, die nicht brüten können nützlich.

Serie: 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.

Vorbereitung der neuronalen Kulturen aus Midgastrula Bühne Drosophila Embryonen

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

Forschung

Biologie

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.

Elektrophysiologischen Methoden für die Aufzeichnung Synaptic Potentials aus dem NMJ von Drosophila-Larven

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.

Elektrophysiologische Aufzeichnung in der Drosophila Embryo

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.

Vorbereitung der Embryonen für Elektronenmikroskopie der Drosophila Embryonale Herz Rohr

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.

In-vivo-Zentrifugation von Drosophila Embryonen

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.

Upright Imaging von Drosophila Embryos

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).

Optimierte Transfektion Strategie für Expression und elektrophysiologische Ableitung von rekombinantem Voltage-Gated Ion Channels in HEK-293T-Zellen

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.

Isolierung und Reinigung von Kinesin aus Drosophila Embryonen

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.

Der Ausarbeitung der einzelnen Drosophila Ei Chambers für 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, ...

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.

Studieren mitotischen Checkpoint durch Veranschaulichung Dynamische Kinetochor Protein Verhalten und Chromosome Motion in Lebens-<em&gt; Drosophila</em&gt; Syncytial Embryonen