Ce travail a été soutenu par des subventions GM096076 du programme AREA NIH.

1. Préparation des mouches pour la dissection <ol><li> Anesthésier les mouches saines du génotype désiré (par exemple, l&#39;expression d&#39;une protéine GFP-étiqueté) qui sont inférieures à une semaine. Sélectionnez 10-15 femelles de grande taille avec des arrondis de couleur crème abdomen. </li><li> Transférez les mouches sélectionnées et quelques (3-5) mâles dans un flacon contenant de nouveaux aliments légèrement yeasted et permettre aux mouches pour engraisser pendant deux jours à 25 ° C. Reportez-vous à Weil et al. 6 pour les détails de préparation drosophile. Engraissement les mouches veille à ce que des stades différents, en particulier les étapes 8-12, sera disponible pour l&#39;imagerie. Mal engraissés ou unfattened vole habituellement ne contiennent que des stades très précoces (1-6) et matures (stade 14) ovocytes. </li></ol> 2. Préparation des ovocytes d&#39;imagerie utilisant un microscope composé vertical <ol><li> Préparer une lame de verre standard par l&#39;apposition de deux lamelles de la lame, à environ 1 cm de distance, à l&#39;aide de silicone GREASe. Utiliser uniquement une graisse suffisante pour assurer une bonne étanchéité entre la lamelle et un toboggan. En appuyant fermement sur chaque lamelle la volonté répartir la graisse couche mince et régulière. Ajouter un peu d&#39;huile halocarbone 27 (Sigma) à la jonction entre les lamelles et le faire glisser. Ils serviront de cales pour empêcher blessant ovocytes isolés dans les étapes ultérieures. N ° 1.5 lamelles fonctionnent bien, avec une épaisseur moyenne (0.16-0.18 mm) qui correspond à celui des ovocytes à un stade avancé. </li><li> Lieu 2-3 gouttes d&#39;huile 27 halocarbure sur la surface d&#39;une lamelle 22 mm 2. Pour obtenir les ovocytes sains possibles, une femelle est retirée du flacon à l&#39;aide alimentaire pipetage à la bouche et lentement déposée dans l&#39;huile sans anesthésiant premier halocarbure. </li><li> En utilisant une pince à dissection (tels que Dumont # 5), la broche à la volée contre la lamelle et retirer le bout de l&#39;abdomen. Enlever les ovaires en les faisant glisser le long de la surface de la lamelle couvre-objet, en vertu de l&#39;huile. Cela permettra de promouvoir adhesion des ovocytes à la lamelle. </li><li> Doucement démêler les ovaires, à la recherche d&#39;ovocytes qui sont 10B étape au moins de l&#39;ovogenèse. A ce stade, les ovocytes peuvent être identifiés par recherche de chambres d&#39;oeuf dans lequel l&#39;ovocyte occupe au moins 50-60% du volume total de la chambre d&#39;oeuf. À ce stade de l&#39;ovogenèse, les cellules folliculaires qui recouvrent l&#39;ovocyte sont devenus colonne, et entourent l&#39;ovocyte de tous les côtés, sauf pour une petite région où les connexions à des cellules nourricières restent. Utilisez la pince pour enlever ovocytes individuels de la gaine ovarioles. </li><li> Procéder, en utilisant des femelles 1-3, 5-8 ovocytes intacts jusqu&#39;à ce que ont été isolées sur la lamelle. Retirez tous les tissus inutilisables. </li><li> Retourner avec précaution la lamelle contenant les ovocytes sur la lame de pré-préparée de telle sorte que les ovocytes sont placés entre les deux entretoises. Ne pas appuyer sur la lamelle inversée car cela endommager les ovocytes. Si nécessaire, ajouter une petite quantité d&#39;huile halocarbone à til bords du &quot;sandwich&quot; pour assurer l&#39;espace est rempli. (Facultatif:. Laisser reposer diapositive pendant plusieurs minutes pour permettre le tassement de la lamelle) La diapositive est maintenant prêt pour l&#39;imagerie. </li></ol> 3. Préparation des ovocytes d&#39;imagerie utilisant un microscope composé inversé <ol><li> Les ovocytes sont disséqués essentiellement comme décrit dans la section 2, sauf que les petites boîtes de Pétri avec un insert à fond de verre (MatTek, 35 mm) sont utilisés au lieu des lamelles. Il n&#39;est pas nécessaire de recouvrir l &#39;échantillon, et après la dissection des tissus peuvent être visualisés directement dans les plats, mais des précautions doivent être prises pour s&#39;assurer que les ovocytes restent couverts d&#39;huile et ne sont pas autorisés à se dessécher. </li></ol> 4. Vivez l&#39;imagerie <ol><li> Apportez un ovocyte dans le foyer en utilisant DIC ou optique à contraste de phase et d&#39;examiner l&#39;ovocyte pour tout signe de dommage, comme une fuite du cytoplasme, ou gonflement des cellules folliculaires. Ovocytes seulement les images qui semblent en bonne santé et en bon état. </li><li> Streaming peut être contrôlée directement sans qu&#39;il soit nécessaire pour l&#39;étiquetage GFP, depuis vésicules vitellines dans le cytoplasme sont autofluorescente 7. Ce signal peut être capté dans la gamme FITC de la portée, mais il faudra probablement des réglages de gain plus élevés que ceux qui sont utilisés pour des protéines GFP-marqués. </li><li> Les meilleures images de diffusion en continu sont obtenues à partir de coupes optiques qui sont juste au-dessous de la surface de l&#39;ovocyte qui est en appui contre la vitre couvre-objet / de Petri en verre. Ici, l&#39;ovocyte est légèrement aplati, de permettre une meilleure plan de mise au point. Les images peuvent être capturées à 200X - 1.000 X. En utilisant le dispositif décrit ici, les objectifs de l&#39;air doit être utilisé, puisque la lamelle forme une barrière entre l&#39;ovocyte et objective. Ceci est optimale car elle minimise grandement la dérive et le mouvement de l&#39;échantillon. </li><li> Une fois une région d&#39;intérêt est mis au point, éteignez l&#39;optique brighfield et passer à l&#39;imagerie confocale. Utilisation du balayage rapide / preview fonction du logiciel, ajuster et d&#39;acquérirle cas échéant. </li><li> Les paramètres de capture varie d&#39;un microscope à microscope, mais les paramètres généraux sont les suivants: Utilisation d&#39;une longueur d&#39;onde d&#39;excitation de 488 nm et ~ une longueur d&#39;onde d&#39;émission de ~ 515 nm. Mise en place de l&#39;axe Z de telle sorte que la capture de l&#39;axe Z reste constante, avec un délai de 10 secondes entre les images. Capturer une séquence test de trames de 5-10 pour assurer l&#39;ovocyte est fixe et le plan de focalisation est approprié. </li><li> Une séquence typique de temps écoulé entre la capture sera de 20 à 50 cadres, et peut être examinée immédiatement après l&#39;achèvement d&#39;évaluer la qualité vidéo. Le processus peut être répété pour d&#39;autres cellules dans la diapositive, ou de femelles supplémentaires peuvent être disséqués selon les besoins. </li></ol> 5. Dépannage et commune <ol><li> Drift - Lors de l&#39;utilisation d&#39;un microscope droit, les cellules peuvent parfois dérives dues à la diffusion de l&#39;huile. Cela peut être minimisé en utilisant juste assez d&#39;huile pour couvrir la zone sous la lamelle. Si le problème persiste, utilisez g de moins en siliconeRease au intercalaires apposer à la diapositive. </li></ol> Remarque: Certains logiciels permettent le suivi confocale de cellules, mais en général, il est préférable de renoncer à l&#39;imagerie d&#39;ovocytes à la dérive et au lieu d&#39;utiliser un autre spécimen. <ol start="2"><li> Aucun signe de streaming - Si le streaming n&#39;est pas évident, il est probablement dû à l&#39;une des deux causes: soit le plan focal pour la capture d&#39;image n&#39;est pas correcte (généralement trop profondément à l&#39;intérieur de l&#39;ovocyte) ou l&#39;ovocyte est endommagé. Temps et la pratique peut réduire à la fois les erreurs. </li></ol>

<ol>
	<li>Becalska, A. N., Gavis, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lighting+up+mRNA+localization+in+Drosophila+oogenesis.">Lighting up mRNA localization in Drosophila oogenesis.</a> Dev. 136, 2493-2503 (2009).</li><li>He, L., Wang, X., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shining+light+on+Drosophila+oogenesis:+Live+imaging+of+egg+development.">Shining light on Drosophila oogenesis: Live imaging of egg development.</a> Curr. Opin. Gen. Dev. 21, 612-619 (2011).</li><li>King, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Ovarian development Drosophila melanogaster. , (1970).</li><li>Spradling, A. C., Bate, M., Martinez-Arias, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+genetics+of+oogenesis.">Developmental genetics of oogenesis.</a> The Development of Drosophila melanogaster. 1, 1-70 (1993).</li><li>Serbus, L. R., Cha, B. -. J., Theurkauf, W. E., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynein+and+the+actin+cytoskeleton+control+kinesin-driven+cytoplasmic+streaming+in+Drosophila+oocytes.">Dynein and the actin cytoskeleton control kinesin-driven cytoplasmic streaming in Drosophila oocytes.</a> Dev. 132, 3743-3752 (2005).</li><li>Weil, T. T., Parton, R. M., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+Individual+Drosophila+Egg+Chambers+for+Live+Imaging.">Preparing Individual Drosophila Egg Chambers for Live Imaging.</a> J. Vis. Exp. (60), e3679 (2012).</li><li>Theurkauf, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Premature+microtubule-dependent+cytoplasmic+streaming+in+cappuccino+and+spire+mutant+oocytes.">Premature microtubule-dependent cytoplasmic streaming in cappuccino and spire mutant oocytes.</a> Science. 265, 2093-2096 (1998).</li><li>Morin, X., Daneman, R., Zavortink, M., Chia, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protein+trap+strategy+to+detect+GFP-tagged+proteins+expressed+from+their+endogenous+loci+in+Drosophila.">A protein trap strategy to detect GFP-tagged proteins expressed from their endogenous loci in Drosophila.</a> PNAS. 98, 15050-15055 (2001).</li><li>R&#246;per, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rtnl1+is+enriched+in+a+specialized+germline+ER+that+associates+with+ribonucleoprotein+granule+components.">Rtnl1 is enriched in a specialized germline ER that associates with ribonucleoprotein granule components.</a> J. Cell Sci. 120, 1081-1092 (2007).</li><li>Pokrywka, N. J., Payne-Tobin, A., Raley-Susman, K. M., Swartzman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubules,+the+ER+and+Exu:+New+associations+revealed+by+analysis+of+mini+spindles+mutations.">Microtubules, the ER and Exu: New associations revealed by analysis of mini spindles mutations.</a> Mech. Dev. 126, 289-300 (2009).</li><li>Petri, W. H., Mindrinos, M. N., Lombard, M. F., Margaritis, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+development+of+the+Drosophila+chorion.ih+a+chemically+defined+organ+culture+medium.">In vitro development of the Drosophila chorion.ih a chemically defined organ culture medium.</a> Wilhelm Roux's Arch., Devl. Biol. 186, 351-362 (1979).</li></ol>

Je n&#39;ai rien à déclarer.

Ovocytes chez la drosophile constituent un système modèle polyvalent pour traiter une variété de questions liées à la fonction et l&#39;organisation des cellules et des composants subcellulaires. Une compréhension complète de la fonction des protéines nécessite le suivi de ces deux aspects spatiaux et temporels du comportement des protéines. Les progrès dans les deux systèmes d&#39;imagerie et les outils génétiques disponibles pour Drosophilists rendre possible, même pour les petits laboratoires pour effectuer des expériences d&#39;imagerie de haute qualité dans des ovocytes de vie. Ici, je ébaucher un protocole d&#39;analyser le comportement des protéines GFP-étiqueté ou autofluorescente et les structures du milieu à la fin de l&#39;ovogenèse qui peut être utilisé en conjonction avec une variété de contextes génétiques pour sonder la fonction des protéines. Dans ce protocole I décrire le processus par lequel les ovocytes sont préparés pour l&#39;imagerie et les réglages de base confocaux qui permettront d&#39;acquisition vidéo laps de temps de protéines fluorescentes et des structures. Supplémentaires details sur la préparation des ovocytes sont décrits par Weil et al. 6 Une grande variété de souches de mouches exprimant des protéines qui ont été marqués de façon endogène par l&#39;intermédiaire de l&#39;exon de piégeage sont disponibles 8 et variants de la protéine infiniment plus peuvent être conçus et réintroduite pour les mouches. En travaillant avec les tissus vivants, la santé des spécimens est d&#39;une importance primordiale. Chambres d&#39;œufs de stade 10B pouvez compléter ovogenèse d&#39;une manière essentiellement autonome 11, ce qui les rend idéales pour des études à court terme d&#39;imagerie. Des précautions doivent être prises lors de plusieurs étapes clés pour obtenir des données de haute qualité. Lors de la dissection, il est important de manipuler les ovaires et d&#39;ovocytes aussi peu que possible, et de minimiser la quantité de temps entre la dissection et l&#39;achèvement de l&#39;imagerie. Idéalement, une station de dissection petite doit être situé dans la même pièce où se trouve le champ confocal, et seulement quelques chambres d&#39;œufs en plusieurs fois devrait être imagé. Le nombre recommandé ici (5-8) Garantit que le temps de dissection est réduit au minimum tout en maximisant la probabilité d&#39;obtenir des images de bonne qualité. Je recommande la capture d&#39;une courte série d&#39;images pour évaluer la qualité d&#39;image et de confirmer que les paramètres de capture sont optimisés, avant de capturer une séquence plus longue laps de temps. La plupart des logiciels permettant d&#39;images individuelles pour être sauvé, et ces images peuvent ensuite être analysées dans une variété de façons en utilisant ImageJ 12 ou d&#39;autres logiciels.

<table border="1"><tbody><tr><td> Nom </td><td> Entreprise </td><td> Numéro de catalogue </td><td> Commentaires </td></tr><tr><td> Huile halocarbone 27 </td><td> Sigma-Aldrich </td><td> H8773-100ML </td><td></td></tr><tr><td> # 5SF forceps </td><td> Outils Fine Science </td><td> 11252-00 </td><td></td></tr><tr><td> Graisse au silicone </td><td> Sigma Aldrich </td><td> Z273554-1EA </td><td></td></tr><tr><td> À fond de verre des boîtes de Pétri </td><td> MatTek </td><td> P35G 0-20-C- </td><td></td></tr></tbody></table>

live imaging of gfp-labeled proteins in drosophila oocytes

L&#39;ovocyte chez la drosophile a été établi comme un système polyvalent d&#39;enquêter sur des questions fondamentales telles que la fonction du cytosquelette, l&#39;organisation des cellules, et la structure et la fonction des organites. La disponibilité de diverses protéines GFP-taggés signifie que de nombreux processus cellulaires peuvent être surveillés dans les cellules vivantes au cours des minutes ou des heures, et l&#39;utilisation de cette technique, des procédés tels que la RNP transport, la morphogenèse épithéliale, et le remodelage tissulaire ont été décrits en détail ovocytes chez la drosophile 1,2. La possibilité d&#39;effectuer une imagerie vidéo combiné avec un riche répertoire de mutants permet une grande variété de gènes et des processus à examiner en détail incroyable. Un exemple en est le processus de diffusion ovoplasmique, qui initie à la mi-ovogenèse 3,4. Ce mouvement vigoureux de vésicules cytoplasmiques est microtubules et la kinésine-dépendante 5 et fournit un sys utilestème d&#39;enquêter sur la fonction du cytosquelette lors de ces étapes. Ici, je présente un protocole pour l&#39;imagerie laps de temps de vie des ovocytes aide de pratiquement n&#39;importe quelle configuration microscopie confocale.

Imagerie de protéines GFP-marqués varie en fonction de la force avec la protéine marquée est exprimée. Un ovocyte à mi-étape exprimer reticulon-like 1 (Rtnl-1), l&#39;exon étiquetés avec la GFP 8,9 produit un signal fort. Rtnl-1 semble co-localisés avec les microtubules présents pendant de longues continu ovoplasmique 10 (figure 1A). Rtnl-1 est un bon marqueur pour observer en continu, et est également un indicateur de l&#39;organisation des microtubules dans les ovocytes à un stade avancé. Le streaming ovoplasmique dans une chambre d&#39;œufs de type sauvage est évident que le mouvement notable de vésicules vitellines autofluorescentes lorsque vous utilisez un taux de capture d&#39;une image à chaque (figure 1B) 10 sec. Une projection maximale de cinq images peuvent être générés pour marquer le chemin de vésicules mobiles. <img alt="Figure 1" src="/files/ftp_upload/50044/50044fig1.jpg" /> Figure 1. Vivez l&#39;imagerie à la fois autoflstructures uorescentes et GFP-étiqueté. A) Un seul time-lapse image d&#39;une étape 11 ovocytes exprimant Rtnl1-GFP à 400X. Rtnl1-GFP fibres se déplacent avec un mouvement ondulatoire dans les ovocytes en streaming. B) Une projection maximale de cinq châssis d&#39;une chambre oeufs de stade 10B dans laquelle les vésicules vitellines autofluorescentes ont été imagées toutes les 10 secondes pendant 50 sec au total, à un grossissement de 400X. Barre d&#39;échelle = 30 um

Watch this Scientific Journal Video about Live Imaging of GFP-labeled Proteins in Drosophila Oocytes at JoVE.com

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Un protocole d&#39;imagerie en temps réel des protéines GFP-marqués ou des structures autofluorescentes en individuel<em&gt; Drosophila</em&gt; Ovocytes est décrite.

Vivez l&#39;imagerie des protéines GFP-étiquetés<em&gt; Drosophila</em&gt; Les ovocytes

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

live-imaging-of-gfp-labeled-proteins-in-drosophila-oocytes

Research

JoVE Journal

Biology

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

11.3K vues.  Vassar College. Un protocole d'imagerie en temps réel des protéines GFP-marqués ou des structures autofluorescentes en individuel Drosophila Ovocytes est décrite.

Vidéo: Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Imagerie de la migration en direct des cellules gliales dans le disque Eye drosophile Imaginal

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

Recherche

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 ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

En direct de l&#39;imagerie Drosophila melanogaster Migrations hémocytaires embryonnaires

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

Marquage fluorescent du COS-7 exprimant des protéines de fusion SNAP-tag pour imagerie de cellules vivantes

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

Préparation de développement et de l&#39;adulte Drosophile Cerveaux et la rétine d&#39;imagerie en temps réel

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

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 ...

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 ...

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.

Préparation individuelle Drosophila Œuf Chambres pour l&#39;imagerie en direct

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

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Deux et trois dimensions imagerie de cellules vivantes de protéines endommagent l&#39;ADN de réponse

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.