Nous tenons à remercier Susan Gerbi, Heidi Smith, et ​​David Lambert pour la fourniture du matériel à tester in vivo centrifugation sur Sciara coprophila et Ilyanassa obsoleta, respectivement. Nous remercions les membres du laboratoire de Welte pour les commentaires sur la technique et du manuscrit. Développement et le raffinement de l&#39;essai de centrifugation in vivo a été soutenue par NIGMS octroi GM64687 au MAE. Les images montrées dans la Fig. 1 et Fig. 5E, F 2 ont déjà été publiés et sont reproduits ici avec sa permission.

<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>

Les étapes critiques Assurez-vous que la concentration en gélose est suffisamment élevée. Si elle est trop faible, l&#39;agar-agar va se désintégrer pendant la centrifugation, et les embryons vont éclater ou être orientés de manière aléatoire. Si la concentration en gélose est légèrement trop faible ou si l&#39;agar-agar est humide (retrait insuffisamment séchés ou insuffisante du TSS en excès dans l&#39;étape 4.3), les embryons va flotter hors de leurs trous pendant la centrifugation et de devenir mal orientées. Si l&#39;agar-agar est versé trop mince, il sera également l&#39;effondrement ou du crack pendant la centrifugation. Assurez-vous que les embryons n&#39;ont pas avancé au-delà stades cellularisation. Sinon, la séparation des organites ne fonctionnera pas (Fig. 5C) depuis organites deviendra séquestrées dans des milliers de cellules individuelles. Chez les jeunes embryons au stade une cellule, les organites sont libres de migrer de grandes distances et s&#39;accumulent en fonction de leur densité caractéristique. Pour éviter d&#39;analyser les embryons qui sont trop vieux, assurez-vous de (i) comprennent une collection pré-pour permettre aux femelles de pondre des œufs anciens (étape 2.4), (ii) percevoir des embryons pour 3 heures ou moins (étape 2.5), (iii) sélectionner visuellement les embryons avant les étapes cellularisation pour s&#39;intégrer dans la gélose (étape 4.2), (iv) tenir intégrant peu de temps pour empêcher le développement jusqu&#39;à la fin de cellularisation (étape 4.6), et (v) de jeter les embryons qui n&#39;ont pas la couche correctement (étape 5.6) . Ne lésinez pas sur le temps de centrifugation. Bien que certains des embryons montrera superposition belle même après seulement 10 minutes de centrifugation, la séparation est incompatible de l&#39;embryon à l&#39;embryon. 30 tours min donnent des résultats très cohérents. En outre, plus le temps de centrifugation, le plus les couches sont stables après la centrifugation est terminée. Ceci est important pour les applications où il faut du temps pour traiter les autres embryons avant de les utiliser (par exemple, s&#39;ils ont besoin d&#39;être traitées pour des expériences de transplantation ou montés pour l&#39;analyse confocale). D&#39;éventuelles modifications Collecte des oeufs et de chorion retrait de nombreux protocoles différents existent pour les étapes 2 et 3 1, 3, 9, et fonctionne aussi bien que celles décrites ici aussi longtemps que les embryons utilisés sont des jeunes, le rendement d&#39;oeuf est élevé, et la fraction de mal embryons en scène est minimisé. Antéro-postérieure d&#39;alignement Le protocole oriente tous les embryons de telle sorte que l&#39;extrémité postérieure est enterré dans la gélose et l&#39;extrémité antérieure est en haut (fig. 3). Cette orientation est bonne pour la cohérence, de sorte qu&#39;il est possible d&#39;utiliser le micropyle comme un repère qui bout pointu pendant la centrifugation (Fig. 1, 5). Cependant, avec la pratique, il est facile de repérer l&#39;orientation des embryons centrifugé par l&#39;apparition distincte du lipide-goutte et les couches de jaune lumineux par microscopie à champ. Alors il devient possible de monter des embryons avec soit finissent; cette flexibilité accélère le processus de montage. Centrifugation non orientés L&#39;aspect le plus-de temps et techniquement difficiles de ce protocole est de fixation des embryons dans l&#39;agar (étape 4). Elle exige de toucher chaque embryon (une fois pour l&#39;insérer dans le trou et une fois de récupérer l&#39;après centrifugation). Comme alternative, on peut transférer des embryons à l&#39;étape 4.1 dans des microtubes rempli de TSS. Lorsque ces tubes sont centrifugés dans une microcentrifugeuse (13.000 rpm, 10-30 min), les embryons seront aussi généralement couche bien 10-13. Cette variation permet de traiter des centaines d&#39;embryons dans le même temps très rapidement. Cependant, la superposition n&#39;est pas aussi cohérente, une fraction des embryons ne se développent pas de couches distinctes, malgré les forces de centrifugation beaucoup plus élevé (~ 16 000 g) que appliquée dans le protocole précédent. Plus difficile est le fait que l&#39;orientation de l&#39;embryon est variable, et on peut observer une séparation à différents angles à l&#39;axe de l&#39;embryon majeur. Cette variabilité impose à l&#39;expérimentateur de trier après la centrifugation comment un embryon particulier a été organisé dans le tube à centrifuger. Avec l&#39;utilisation de l&#39;autofluorescence jaune comme un marqueur de la partie la plus dense de-l&#39;embryon, ce qui est souvent possible, mais il exige beaucoup de pratique et plus de jugement pour le faire de manière fiable. S&#39;il ya des embryons suffisamment dans l&#39;échantillon, on peut être capable de trier les cas où les embryons se trouvait être disposés comme dans la Fig. 1. La centrifugation des embryons dans leur coquille d&#39;œuf, il est possible d&#39;intégrer des embryons dans l&#39;agar pour l&#39;étape 4 sans avoir à retirer le chorion 2 premiers. Dans cette approche, les embryons sur la plaque de collecte sont couverts avec de l&#39;huile d&#39;halocarbures au lieu d&#39;eau de Javel à 50% à l&#39;étape 3.2 (halocarbures tourne le chorion translucide). Les embryons de stades appropriés sont sélectionnés et transférés à une plaque de gélose nouvelle aide de pinces; en saisissant uniquement les filaments d&#39;oeuf avec la pince à épiler, des dommages auxle bon embryon est évitée. Les embryons sont intégrées comme dans l&#39;étape 4.4 à 4.8, avec les filaments d&#39;oeuf antérieure collant vers le haut du trou dans la gélose. La coquille d&#39;œuf est éliminé par traitement javel 50% après les embryons ont été centrifugés et creusé de l&#39;agar comme à l&#39;étape 5.4. La centrifugation en présence de la coquille d&#39;oeuf peut améliorer le taux de devitellinization après fixation (étape 6.2), mais la sélection des étapes correctes pour la centrifugation (étape 4.2) est plus difficile. Applications Centrifugation embryon est particulièrement utile pour des expériences de colocalisation, soit pour tester si une protéine particulière se localise dans une certaine organite majeur. Par exemple, la protéine est présente sur Klar début gouttelettes lipidiques embryonnaire, pourtant, cette localisation est difficile à démontrer dans les embryons intacts. En partie, cela est parce que les deux points lacrymaux et Klar gouttelettes lipidiques sont abondants dans la périphérie 10 embryons, ce qui rend difficile de colocalisation fallacieuses exclure. Toutefois, la centrifugation concentre gouttelettes lipidiques dans une couche distincte, et donc, colocalisation avec signal Klar devient évident 10. Une analyse similaire a démontré sans équivoque la localisation d&#39;autres protéines à 8 gouttelettes lipidiques, 13-15, y compris des exemples surprenants, comme les histones certains 2 et dans les cas où une protéine est présente de façon ubiquitaire, mais enrichi sur 8 gouttelettes lipidiques. Comme les embryons peuvent être sélectionnées visuellement par le stade de développement (étape 4.2), il est même possible de détecter le recrutement régulés par le développement de protéines à 8 gouttelettes lipidiques, 15. Test de colocalisation avec des gouttelettes lipidiques est particulièrement facile puisque gouttelettes lipidiques s&#39;accumulent dans un visuellement distincte &quot;bouchon&quot;, donc pas de marqueur supplémentaires ou de la teinture est nécessaire pour identifier cette couche. En fait, en raison de sa facilité et potentiellement frappante résultats visuels (Fig. 1, Fig. 5 D, E), notre laboratoire utilise cette technique maintenant couramment comme un premier test si une protéine candidate est localisée à gouttelettes lipidiques. En principe, les études de colocalisation similaires devraient travailler pour le autres organites Fig. 1, et probablement pour d&#39;autres (y compris les parasites intracellulaires) dont la distribution dans les embryons centrifugé n&#39;a pas encore été documentée. Depuis centrifugation concentre organites en bandes serrées, il facilite l&#39;isolement d&#39;une fraction enrichie dans ces organites. Cette approche a déjà été utilisé pour récupérer des gouttelettes lipidiques pour la transplantation en 2 embryons donateurs. Il peut également être possible d&#39;analyser biochimiquement la fraction enrichie (par exemple, en coupant les couches hors embryons après fixation 15). La méthode décrite ici a des applications au-delà des embryons de drosophile, comme la centrifugation peuvent être utilisées pour stratifier les œufs de différentes espèces (y compris les grenouilles, les nématodes, les tuniciers, les annélides, les oursins, les mollusques, et les cnidaires) 16. Dans notre propre laboratoire, nous avons utilisé le protocole ci-dessus pour les embryons de la mouche domestique Musca domestica 2 et ont eu recours à la centrifugation non orientés dans l&#39;analyse des embryons de la coprophila sciarides Sciara 17 et l&#39;escargot Ilyanassa obsoleta 17. Enfin, cette méthode n&#39;est pas limitée aux embryons: Centrifugation peut également être utile pour stratifier les autres grandes cellules, telles que des ovocytes (par exemple, chez la drosophile 2, 18 et l&#39;asclépiade bug Oncopeltus fasciatus 17). Pour obtenir un spécimen différent, les conditions de centrifugation exactes doivent être optimisés: Par exemple, 10 minutes à 3000 g est suffisant pour stratifier efficacement embryons de Musca domestica 2, et si centrifugé beaucoup plus long, les embryons ont tendance à se briser lors de la fixation et immunocoloration.

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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>
<td>Fine Science Tools</td>
<td>26016-12</td>
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<td>Fly Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific</td>
<td>59-100 or 59-101</td>
<td>For 65 mm or 100 mm diameter Petri dishes, respectively</td>
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<td>Halocarbon Oil 27</td>
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in-vivo centrifugation of drosophila embryos

Une stratégie importante pour purifier et d&#39;isoler les différents types d&#39;organites intracellulaires est de les séparer les uns des autres en fonction des différences dans la densité de flottaison. Toutefois, lorsque les cellules sont perturbés avant la centrifugation, les protéines et les organites dans cet environnement non indigènes souvent de manière inappropriée coller les unes aux autres. Nous décrivons ici une méthode pour séparer par des organites de densité dans intactes, des embryons de drosophile vivant. Embryons précoces, avant cellularisation sont récoltés dans des cages de la population, et leurs coquilles d&#39;œuf extérieures sont éliminés par traitement à l&#39;eau de Javel à 50%. Les embryons sont ensuite transférés à une plaque de gélose de petites et insérées, extrémité postérieure d&#39;abord, dans de petits trous verticaux dans la gélose. Les plaques contenant des embryons incorporés sont centrifugés pendant 30 min à 3000g. L&#39;agar soutient les embryons et les maintient dans une orientation définie. Ensuite, les embryons sont creusés dans la gélose avec une aiguille émoussée. La centrifugation sépare organites majeurs en couches distinctes, une stratification facilement visible par microscopie à champ lumineux. Un certain nombre de marqueurs fluorescents sont disponibles pour confirmer la stratification succès dans les embryons vivants. Protéines associées à certains organites sera enrichie d&#39;une couche particulière, démontrant colocalisation. Couches individuelles peuvent être récupérées pour analyse biochimique ou transplantation dans des oeufs de donneur. Cette technique est applicable pour la séparation des organites dans d&#39;autres grandes cellules, y compris les œufs et les ovocytes d&#39;espèces diverses.

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

In-vivo Centrifugation of Drosophila Embryos

Nous décrivons une méthode d&#39;organites séparés par la densité dans la vie<em&gt; Drosophile</em&gt; Embryons. Les embryons sont intégrés dans l&#39;agar et centrifugé. Cette technique donne une séparation reproductible des organites majeurs long de l&#39;axe antéro-postérieure d&#39;embryon. Cette méthode facilite les expériences de colocalisation et les rendements des fractions organite pour l&#39;analyse biochimique et des expériences de transplantation.

In vivo d&#39;embryons de drosophile Centrifugation

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

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JoVE Journal

Biology

In-vivo Centrifugation of Drosophila Embryos

18.3K vues.  University of Rochester. Nous décrivons une méthode d'organites séparés par la densité dans la vie Drosophile Embryons. Les embryons sont intégrés dans l'agar et centrifugé. Cette technique donne une séparation reproductible des organites majeurs long de l'axe antéro-postérieure d'embryon. Cette méthode facilite les expériences de colocalisation et les rendements des fractions organite pour l'analyse biochimique et des expériences de transplantatio...

Vidéo: 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.

Préparation des cultures de neurones d&#39;embryons de drosophile Midgastrula scène

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

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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.

La préparation des Drosophile Les embryons destinés à Live-imagerie utilisant le protocole goutte suspendue

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.

Mésoscopique fluorescence tomographie pour In vivo L&#39;imagerie du développement Drosophile</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.

Préparation d&#39;embryons pour la microscopie électronique des Drosophile Tube cardiaque embryonnaire

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.

Imagerie verticale de<em&gt; Drosophila</em&gt; Embryons

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.

Cultures de cellules primaires à partir Drosophile Embryons Gastrula

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.

L&#39;imagerie in vivo chez la drosophile larves Intact au sub-cellulaire Résolution

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.

Interférences ARNi par injection dans les ARNdb Drosophile Embryons

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.

Changement de forme d&#39;imagerie cellulaire en vie Drosophile Embryons

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.

Isolement et purification de la kinésine de Drosophila Embryons

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