Wir bedanken uns bei Susan Gerbi, Heidi Smith und David Lambert für die Versorgung Material in-vivo-Zentrifugation auf Sciara coprophila und Ilyanassa obsoleta, verglichen. Wir danken Mitglieder der Welte-Labor für die Kommentare auf die Technik und Handschrift. Entwicklung und Verfeinerung der in-vivo-Assay wurde durch Zentrifugation NIGMS gewähren GM64687 zu MAW unterstützt. Die Bilder in Abb.. 1 und Abb.. 5E wurden F bisher 2 veröffentlicht und sind hier mit freundlicher Genehmigung abgedruckt.

<ol>
	<li>Wieschaus, E., N&#252;sslein-Volhard, C., Roberts, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Looking+at+embryos.">Looking at embryos.</a> Drosophila: A Practical Approach. , 179-214 (1998).</li><li>Cermelli, S., Guo, Y., Gross, S. P., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lipid+droplet+proteome+reveals+that+droplets+are+a+protein+storage+depot.">The lipid droplet proteome reveals that droplets are a protein storage depot.</a> Curr Biol. 16, 1783-1795 (2006).</li><li>Kiehart, D. P., Crawford, J. M., Montague, R. A., Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+microinjection+of+Drosophila+embryos.">Quantitative microinjection of Drosophila embryos.</a> Drosophila Protocols. , 345-359 (2000).</li><li>LaJeunesse, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three+new+Drosophila+markers+of+intracellular+membranes.">Three new Drosophila markers of intracellular membranes.</a> Biotechniques. 36 (784-788), 790-790 (2004).</li><li>Schuh, M., Lehner, C. F., Heidmann, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incorporation+of+Drosophila+CID/CENP-A+and+CENP-C+into+centromeres+during+early+embryonic+anaphase.">Incorporation of Drosophila CID/CENP-A and CENP-C into centromeres during early embryonic anaphase.</a> Curr Biol. 17, 237-243 (2007).</li><li>Clarkson, M., Saint, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+His2AvDGFP+fusion+gene+complements+a+lethal+His2AvD+mutant+allele+and+provides+an+in+vivo+marker+for+Drosophila+chromosome+behavior.">A His2AvDGFP fusion gene complements a lethal His2AvD mutant allele and provides an in vivo marker for Drosophila chromosome behavior.</a> DNA Cell Biol. 18, 457-462 (1999).</li><li>Welte, M. A., Gross, S. P., Postner, M., Block, S. M., Wieschaus, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+regulation+of+vesicle+transport+in+Drosophila+embryos:+forces+and+kinetics.">Developmental regulation of vesicle transport in Drosophila embryos: forces and kinetics.</a> Cell. 92, 547-557 (1998).</li><li>Tran, S. L., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=unpublished+observations.">unpublished observations.</a> , .</li><li>Rothwell, W. F., Sullivan, W., Sullivan, W., Ashburner, M., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+analysis+of+Drosophila+embryos.">Fluorescent analysis of Drosophila embryos.</a> Drosophila Protocols. , 141-157 (2000).</li><li>Guo, Y., Jangi, S., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organelle-specific+Control+of+Intracellular+Transport:+Distinctly+Targeted+Isoforms+of+the+Regulator+Klar.">Organelle-specific Control of Intracellular Transport: Distinctly Targeted Isoforms of the Regulator Klar.</a> Mol Biol Cell. 16, 1406-1416 (2005).</li><li>Meyer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Overlapping+Functions+of+Argonaute+Proteins+in+Patterning+and+Morphogenesis+of+Drosophila+Embryos.">Overlapping Functions of Argonaute Proteins in Patterning and Morphogenesis of Drosophila Embryos.</a> PLoS Genet. 2, 1224-1239 (2006).</li><li>Shubeita, G. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Consequences+of+motor+copy+number+on+the+intracellular+transport+of+kinesin-1-driven+lipid+droplets.">Consequences of motor copy number on the intracellular transport of kinesin-1-driven lipid droplets.</a> Cell. 135, 1098-1107 (2008).</li><li>Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+lipid-droplet+transport+by+the+Perilipin+homologue+LSD2.">Regulation of lipid-droplet transport by the Perilipin homologue LSD2.</a> Curr. Biol. 15, 1266-1275 (2005).</li><li>Phadnis, N., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Phadnis, N., Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Morgan, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+XXII:+The+redistribution+of+the+visible+materials+of+the+egg+by+centrifuging.">Chapter XXII: The redistribution of the visible materials of the egg by centrifuging.</a> Experimental Embryology. , (1927).</li><li>Welte, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unpublished+Observations.">Unpublished Observations.</a> , .</li><li>Bownes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abnormal+oogenesis+and+embryogenesis+resulting+from+centrifuging+Drosophila+melanogaster+females.">Abnormal oogenesis and embryogenesis resulting from centrifuging Drosophila melanogaster females.</a> J Embryol Exp Morphol. 40, 65-81 (1977).</li></ol>

Kritische Schritte Stellen Sie sicher, dass der Agar-Konzentration hoch genug ist. Wenn es zu niedrig ist, wird der Agar während der Zentrifugation zerfallen, und die Embryonen platzen oder zufällig orientiert. Wenn der Agar-Konzentration ist nur etwas zu niedrig ist oder wenn der Agar ist nass (unzureichend getrocknet oder unzureichende Entfernung von überschüssigem TSS in Schritt 4,3), werden die Embryonen float aus ihren Löchern während des Laufs und werden falsch ausgerichtet. Wenn der Agar zu dünn gegossen ist, wird es auch zusammenbrechen oder Crack während der Zentrifugation. Stellen Sie sicher, dass Embryonen nicht über Zellularisierung Stadien fortgeschritten. Anderenfalls wird die Trennung von Organellen nicht funktioniert (Abb. 5C) seit Organellen wird sequestriert in Tausenden von einzelnen Zellen. In jüngeren Embryonen auf der einen Zell-Stadium sind Organellen frei, um große Strecken wandern und reichern sich entsprechend ihrer charakteristischen Dichte. Um zu vermeiden, zu analysieren Embryonen, die zu alt sind, stellen Sie sicher, (i) eine Pre-Collection, damit Frauen, alte Eier (Schritt 2,4), (ii) zu sammeln Embryonen für 3 Stunden oder weniger (Schritt 2,5), (iii) lagen visuell auswählen Embryonen vor Zellularisierung Stadien für die Einbettung in Agar (Schritt 4,2), (iv) halten die Einbettung der Zeit kurz ist, um die Entwicklung bis zum Ende des Zellularisierung (Schritt 4,6) zu verhindern, und (v) zu verwerfen Embryonen, die nicht korrekt Schicht (Schritt 5,6) . Sie sparen nicht auf die Zentrifugationszeit. Obwohl einige der Embryonen schöne Schichtung auch nach nur 10 Minuten Zentrifugation zeigen, ist die Trennung inkonsistent vom Embryo zum Embryo. 30 min dreht geben sehr konsistente Ergebnisse. Auch, desto länger die Zentrifugation Zeit, je länger die Schichten stabil sind nach der Zentrifugation wurde beendet. Dies ist für diejenigen Anwendungen, bei denen es braucht Zeit, um die Embryonen weiter verarbeiten vor Gebrauch wichtig (z. B., wenn sie für die Transplantation Versuche verarbeitet werden oder angebracht zum konfokalen Analyse benötigen). Mögliche Änderungen Eiersammlung und Chorion Entfernung Viele verschiedene Protokolle gibt es für die Schritte 2 und 3 1, 3, 9, und wird ebenso wie Arbeiten der hier beschriebenen, solange die Embryonen verwendet jung sind, ist Ei Ausbeute hoch, und der Anteil der mis- inszeniert Embryonen wird minimiert. Anterior-posteriore Ausrichtung Das Protokoll orientiert alle Embryonen, so dass das hintere Ende in der Agar ist begraben und am vorderen Ende ist (Abb. 3). Diese Orientierung ist gut für die Konsistenz, so dass es möglich ist, die Mikropyle als Landmarke, die am Ende während der Zentrifugation (Abb. 1, 5) wies verwenden. Doch mit der Praxis ist es einfach, wählen Sie die Ausrichtung des zentrifugiert Embryonen durch die unterschiedliche Aussehen der Lipid-Tröpfchen und Eigelb Schichten von Hellfeld-Mikroskopie. Dann wird es möglich, Embryonen mit einem Ende auffahren; diese Flexibilität beschleunigt die Montage-Prozess. Unorientierten Zentrifugation Die-zeitaufwendig und technisch anspruchsvollen Aspekt des Protokolls ist die Montage des Embryos in Agar (Schritt 4). Es erfordert berühren Embryo zweimal (einmal, um sie in das Loch und einmal, um es nach der Zentrifugation zu erholen). Als Alternative kann man Embryonen im Schritt 4.1 Transfer in Reaktionsgefäßen mit TSS gefüllt. Wenn diese Rohre in einer Mikrozentrifuge (13.000 rpm, 10-30 min) werden zentrifugiert, Embryonen wird in der Regel auch Schicht gut 10-13. Diese Variante ermöglicht es, Hunderte von Embryonen zur gleichen Zeit sehr schnell zu verarbeiten. Allerdings ist Schichtung nicht so konsequent, nur einen Bruchteil der Embryonen nicht entwickeln unterschiedliche Schichten trotz wesentlich höheren Zentrifugalkräften (~ 16.000 g) als in dem Protokoll über aufgetragen. Anspruchsvoller ist die Tatsache, dass die Ausrichtung der Embryonen Variable ist, und man kann die Trennung in verschiedenen Winkeln zu den großen Embryo Achse zu beobachten. Diese Variabilität erfordert die Versuchsleiter zu klären, nach der Zentrifugation, wie ein bestimmter Embryo im Zentrifugenglas wurde angeordnet. Mit dem Einsatz des Dotters Autofluoreszenz als Marker für die-dichte Teil des Embryos, ist dies oft möglich, obwohl es viel Übung und mehr Urteil, um es sicher zu tun erfordert. Wenn es genügend Embryonen in der Probe vorhanden sind, kann man in der Lage sein zu klären, die Fälle, in denen Embryonen geschehen, wie in Abb. arrangiert werden. 1. Zentrifugation von Embryonen in ihren Eierschalen Es ist möglich, Embryonen in Agar für Schritt 4 einzubetten, ohne den Chorion ersten 2. In diesem Ansatz werden Embryonen für die Sammlung Platte mit Halocarbonöl anstelle von 50% Bleichmittel in Schritt 3,2 (Halogenkohlenwasserstoff wird das Chorion transluzent) abgedeckt. Embryonen von geeigneten Stufen gewählt werden und auf eine neue Agarplatte mit einer Pinzette, durch Greifen nur das Ei Filamente mit der Pinzette, Schäden ander Embryo richtige ist vermieden. Die Embryonen werden wie in Schritt 4.4 durch 4,8 eingebettet, mit der vorderen Ei Filamente ragte aus dem Loch in der Agar. Die Eierschale ist um 50% Bleichmittel Behandlung entfernt werden, nachdem die Embryonen zentrifugiert wurde und aus dem Agar als in Schritt 5.4 gegraben. Zentrifugation in der Gegenwart der Eischale kann zur Verbesserung der Rate von devitellinization nach der Fixierung (Schritt 6.2), aber die Auswahl der richtigen Stadien für die Zentrifugation (Schritt 4.2) ist eine größere Herausforderung. Anwendungen Embryo Zentrifugieren ist besonders nützlich für Kolokalisation Experimente, dh zu testen, ob ein bestimmtes Protein lokalisiert zu einem gewissen wichtigen Organellen. Zum Beispiel der Klar-Proteins an der frühen Embryonalentwicklung Lipidtröpfchen wird, ist noch dieser Lokalisation eine Herausforderung, in intakten Embryonen zu demonstrieren. Zum Teil ist dies, weil sowohl Klar puncta und Fetttröpfchen in den Embryo Peripherie 10 sind reichlich vorhanden, so dass störende Kolokalisation schwer auszuschließen. Allerdings konzentriert sich Zentrifugation Fetttröpfchen in eine deutliche Schicht, und somit wird Kolokalisation mit Klar-Signal auf der Hand 10. Ähnliche Analyse hat eindeutig die Lokalisierung von anderen Proteinen zu Lipidtröpfchen 8, 13-15, darunter überraschende Beispiele wie bestimmte Histone 2 und in Fällen, in denen ein Protein vorhanden ist ubiquitär nachgewiesen, sondern bereichert auf Lipidtröpfchen 8. Als Embryonen können optisch durch Entwicklungsstadium (Schritt 4,2) gewählt werden, ist es sogar möglich, entwicklungsregulierten Rekrutierung von Proteinen an Lipidtröpfchen 8, 15 zu erkennen. Testing für Kolokalisation mit Fetttröpfchen ist besonders einfach, da die Fetttröpfchen in eine optisch deutliche &quot;cap&quot; akkumulieren, so dass keine weiteren Marker oder Flecken ist erforderlich, um diese Schicht zu identifizieren. In der Tat, wegen seiner Leichtigkeit und potenziell auffälligen optischen Ergebnisse (Abb. 1, Abb.. 5 D, E), beschäftigt unser Labor diese Technik nun routinemäßig als ein erster Test, ob ein Kandidat Protein Lipidtröpfchen lokalisiert ist. Im Prinzip sollte ähnlich Kolokalisation Studien für die anderen Organellen in Abb. arbeiten. 1, und wahrscheinlich für andere (einschließlich intrazelluläre Parasiten), deren Verteilung in zentrifugiert Embryonen ist noch nicht dokumentiert werden. Da Zentrifugation konzentriert Organellen in engen Bändern, erleichtert es Isolierung einer Fraktion in dieser Organellen angereichert. Dieser Ansatz wurde bereits verwendet, um die Fetttröpfchen zur Transplantation in Spender Embryonen 2. Datenwiederherstellung. Es kann auch möglich sein, biochemisch analysieren die angereicherte Fraktion (zB durch Abschneiden Schichten aus Embryonen nach der Fixierung 15). Die hier beschriebene Methode hat Anwendungen jenseits Drosophila-Embryonen, wie Zentrifugation kann verwendet werden, um die Eier der vielen verschiedenen Arten (einschließlich Frösche, Nematoden, Manteltiere, Ringelwürmer, Seeigel, Muscheln und Nesseltiere) 16 schichten werden. In unserem eigenen Labor haben wir das Protokoll oben für Embryonen von der Stubenfliege Musca domestica 2 verwendet und eingesetzt unorientierten Zentrifugation in der Analyse von Embryonen aus der Trauermücke Sciara coprophila 17 und die Schnecke Ilyanassa obsoleta 17. Schließlich ist diese Methode nicht für Embryonen beschränkt: Die Zentrifugation kann auch sinnvoll sein, andere große Zellen, wie Oozyten (zB in Drosophila 2, 18 und die milkweed bug Oncopeltus fasciatus 17) schichten. Für verschiedene Muster, müssen die genauen Zentrifugation Bedingungen optimiert werden: Zum Beispiel ist 10 Minuten bei 3000 g ausreichend ist, um effizient zu schichten Embryonen von Musca domestica 2, und wenn zentrifugiert viel länger, neigen die Embryonen auseinander zu brechen während der Fixierung und Immunfärbung.

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<tr>
<td>60x15 mm Petri dishes</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>#35-1007</td>
<td>From Falcon</td>
</tr>
<tr>
<td>Wire mesh</td>
<td>&nbsp;</td>
<td>Small Parts</td>
<td>CX-0150</td>
<td>stainless steel type 304; mesh size #150 x150, wire diameter: 0.0026 inches</td>
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<td>Tungsten needles</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>10130-10</td>
<td>0.25 mm</td>
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<td>Moria Nickel Plated Pin/Needle Holder</td>
<td>&nbsp;</td>
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<td>Fly Cages</td>
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<td>Genesee Scientific</td>
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in-vivo centrifugation of drosophila embryos

Eine wichtige Strategie zur Reinigung und Isolierung verschiedener intrazellulären Organellen ist, sie voneinander zu trennen basierend auf Unterschieden in Schwebedichte. Allerdings, wenn die Zellen vor der Zentrifugation, Proteinen und Organellen in dieser nicht-native Umgebung gestört werden oft fälschlicherweise aneinander kleben. Hier beschreiben wir eine Methode zur Trennung von Organellen Dichte in intakten, lebenden Drosophila-Embryos. Frühe Embryonen vor Zellularisierung sind aus Käfigen Bevölkerung geerntet, und ihre äußeren Eierschalen werden durch Behandlung mit 50% Bleichmittel entfernt. Die Embryonen werden dann zu einem kleinen Agarplatte und eingefügt, hinteren Ende zuerst, in kleinen vertikalen Löcher in den Agar übertragen. Die Platten mit eingebetteten Embryonen werden für 30 min bei 3000g zentrifugiert. Der Agar unterstützt die Embryonen und hält sie in einer definierten Orientierung. Anschließend werden die Embryonen aus dem Agar mit einer stumpfen Nadel gegraben. Zentrifugation trennt wichtigsten Organellen in verschiedene Schichten, eine Schichtung gut sichtbar von Hellfeld-Mikroskopie. Eine Reihe von fluoreszierenden Markern zur Verfügung stehen erfolgreiche Schichtung in lebenden Embryonen zu bestätigen. Proteine ​​mit bestimmten Organellen assoziiert wird in einer bestimmten Schicht angereichert werden, zeigt Kolokalisation. Einzelne Schichten können zur biochemischen Analyse oder Transplantation in gespendeten Eizellen gewonnen werden. Diese Technik ist für Organelle Trennung in anderen großen Zellen, einschließlich der Eier und Eizellen von verschiedenen Arten.

Watch this Scientific Journal Video about In-vivo Centrifugation of Drosophila Embryos at JoVE.com

In-vivo Centrifugation of Drosophila Embryos

Wir beschreiben eine Methode zur Trennung von Organellen Dichte in lebenden<em&gt; Drosophila</em&gt; Embryonen. Die Embryonen werden in Agar eingebettet und zentrifugiert. Diese Technik liefert reproduzierbare Trennung von wichtigen Organellen entlang der anterior-posterior Embryo-Achse. Diese Methode erleichtert die Kolokalisation Experimente und liefert Organelle Fraktionen für die biochemische Analytik und Transplantation Experimente.

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 ...

in-vivo-centrifugation-of-drosophila-embryos

Research

JoVE Journal

Biology

In-vivo Centrifugation of Drosophila Embryos

18.3K Aufrufe.  University of Rochester. Wir beschreiben eine Methode zur Trennung von Organellen Dichte in lebenden Drosophila Embryonen. Die Embryonen werden in Agar eingebettet und zentrifugiert. Diese Technik liefert reproduzierbare Trennung von wichtigen Organellen entlang der anterior-posterior Embryo-Achse. Diese Methode erleichtert die Kolokalisation Experimente und liefert Organelle Fraktionen für die biochemische Analytik und Transplantation Experimente.

Serie: In-vivo Centrifugation of Drosophila Embryos

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

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Die Herstellung Drosophila Embryonen für Live-Imaging mit dem Hanging-Drop-Protokoll

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

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

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

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

Mesoscopic Fluoreszenz-Tomographie für In-vivo Imaging der Entwicklungsländer Drosophila</em

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

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 ...

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 ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

Primären Zellkulturen aus Drosophila Gastrula Embryonen

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

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

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

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

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

In-vivo-Imaging von Intact Drosophila-Larven auf subzellulärer Auflösung

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

RNAi Interference by dsRNA Injection into Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements and tools for genetic manipulation have become available in Drosophila 2. Soon after the initial discovery by Frie and Mello 3 that double stranded RNA can be used to knockdown the activity of individual genes in Caenorhabditis elegans, RNA interference (RNAi) was shown to provide a powerful reverse genetic approach to analyze gene functions in Drosophila organ development 4, 5.
Many organs, including lung, kidney, liver, and vascular system, are composed of branched tubular networks that transport vital fluids or gases 6, 7. The analysis of Drosophila tracheal formation provides an excellent model system to study the morphogenesis of other tubular organs 8. The Berkeley Drosophila genome project has revealed hundreds of genes that are expressed in the tracheal system. To study the molecular and cellular mechanism of tube formation, the challenge is to understand the roles of these genes in tracheal development. Here, we described a detailed method of dsRNA injection into Drosophila embryo to knockdown individual gene expression. We successfully knocked down endogenous dysfusion(dys) gene expression by dsRNA injection. Dys is a bHLH-PAS protein expressed in tracheal fusion cells, and it is required for tracheal branch fusion 9, 10. dys-RNAi completely eliminated dys expression and resulted in tracheal fusion defect. This relatively simple method provides a tool to identify genes requried for tissure and organ development in Drosophila.

RNAi Interference by dsRNA Injection into Drosophila Embryos

RNA interference has been proven very effective to analyze gene function in Drosophila tracheal development. A detailed protocol used by Jiang lab to inject dsRNA into fly embryos to knockdown gene expression is illustrated. This technique has the potential for screening genes required for tissue and organ development in Drosophila.

RNAi-Interferenz durch dsRNA Injektion in Drosophila Embryos

Genetic screening is one of the most powerful methods available for gaining insights into complex biological process 1. Over the years many improvements ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Imaging Zellgröße Veränderung der Lebens- Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

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 ...