We appreciate and would like to thank the Howard Hughes Medical Institute for the HHMI Professor award to U.B. which made this project possible. We thank the college at the University of California, Los Angeles for providing facilities and teaching infrastructure support for this work. The work was also supported with funding from Midwestern University and a generous donation from the Charity Fidelity Gift Fund. We thank John VandenBrooks for comments on the manuscript and Krista Pearman for her technical assistance.

Cette procédure est une procédure de deux jours. Jour 1 (2 h + temps de dissection) Dissection 1. pupes yeux disque <ol><li> Sélectionnez une nymphe pour la dissection. NOTE: L&#39;âge de la nymphe à être disséqué sera déterminé par les besoins expérimentaux. Toutefois, si l&#39;examen de la morphologie des cellules, cela est souvent effectué à 42 h après la formation de pupe (CSA) à 25 ° C, ce qui est l&#39;âge de l&#39;pupes montré dans la vidéo. Recueillir pupes blanc (considéré comme de l&#39;APF 0 h) avec un pinceau mouillé et les disposer dans l&#39;ordre chronologique (en fonction du temps de collecte) dans un nouveau flacon de nourriture à la mouche maintenue à 25 ° C. Disséquer les nymphes ans dans ce même ordre chronologique afin de maintenir les différences de temps entre les nymphes aussi près que possible. </li><li> Transférer la nymphe dans une goutte de solution saline tamponnée au phosphate (PBS, pH 7,4, voir matériaux pour la recette) sur une plaque de silicone de dissection. </li><li> Tenez le haut de la coque de nymphose avec une pince et percer et ouvrir la lpartie de tournesol de la pupe avec une paire de pinces coupantes. Retirer la nymphe de l&#39;affaire dans le trou nouvellement formé. </li><li> Faire une coupe en diagonale avec des ciseaux, mi-thorax, à la nymphe. Retirer les débris ou transférer la tête de nymphe à une nouvelle baisse de PBS. Nettoyer avec le tube pipette de ventilateur (voir Figure 1 et matériaux pour plus de détails) et ajouter d&#39;autres PBS selon les besoins. Assurez-vous d&#39;éviter la déshydratation. </li><li> La poignée fermement le haut de la tête cuticule nymphale avec une paire de pinces. Avec la pointe du tube de ventilateur, poussez doucement sur la coque de nymphose à extruder le cerveau et d&#39;autres tissus, y compris les disques de l&#39;oeil, sur la coque de nymphose. Soyez sûr d&#39;éviter l&#39;accaparement tout tissu dans la cuticule. </li><li> Utilisation du tube de soufflage remplie de PBS, soufflez doucement le tissu de corps gras et d&#39;autres débris sur le cas de la tête de séparer clairement et identifier les disques du cerveau et des yeux. REMARQUE: Garder le cerveau fixé au boîtier de la tête de pupe permettra au chercheur d&#39;utiliser l&#39;ilannonce cas comme une &quot;poignée&quot; de manipulation avenir du tissu sans endommager les disques des yeux ou le cerveau. Cependant, il est courant de déloger le cerveau et les yeux de disques le cas de la tête et ce ne nuit pas à l&#39;ensemble de la procédure et les résultats. Il faut souvent plusieurs tentatives de souffler et pousser à extraire avec succès les disques œil et le cerveau de l&#39;intérieur de la tête pupe et afin qu&#39;ils soient clairement séparés du tissu environnant. Le nettoyeur et un plus séparé peuvent obtenir les disques d&#39;oeil du tissu environnant aidera à prévenir les tissus étrangers de fixation sur les disques de l&#39;oeil. </li><li> Après dissection, transférer immédiatement le tissu dans un plat en verre de 3 bien sur la glace avec 0,5 ml de solution Fix (voir matériaux). Flotter le tissu sur la surface de la solution Fix pendant quelques minutes (généralement le temps qu&#39;il faut pour disséquer le prochain jeu de disques de l&#39;œil). REMARQUE: Si ce ne sont pas le premier disque d&#39;oeil à être disséqué dans cette ronde de dissection, Completely immerger le disque de l&#39;œil précédente dans la solution Fix. Si l&#39;exécution des procédures de coloration spéciales (par exemple, les espèces réactives de l&#39;oxygène, les lysosomes, etc.) cela doit être fait avant la fixation. Pour ce faire, stocker les disques oculaires disséqués dans du PBS glacé avant de continuer avec le protocole de coloration appropriée. </li><li> Après dissection et l&#39;immersion de tous les disques de l&#39;oeil étant disséqué, déplacer le plat en verre de 3 à bien un rocker et le rock à la température ambiante pendant 30 min. </li></ol> 2. Immunocoloration <ol><li> Retirez délicatement la solution Fix et ajouter 0,5 ml de PBS avec 0,3% de Triton X-100 (0,3% PBT, voir matériaux) et laver le tissu sur une bascule pour 10 min. Répétez cette étape trois fois pour un total de 4 lavages (10 minutes chacun). REMARQUE: élimination soigneuse de la solution dans le puits peut être réalisée sous un microscope à dissection pour prévenir les lésions tissulaires ou de la perte. La solution peut être aspiré soigneusement avec une pipette; nous préférons une amende-tipped, jetable pipette de transfert qui a eu l&#39;ouverture aplatie pour réduire le risque de tissus aspiration. </li><li> A la fin de la 4ème lavage, retirer le lavage et ajouter 0,5 ml de solution de bloc (voir matériaux) pour le bien et incuber pendant 30 min. NOTE: Ce délai peut être prolongé en fonction de vos besoins. </li><li> A la fin du temps de blocage de 30 min, la pipette 10 ul de la solution d&#39;anticorps primaire (dilué dans une solution Block) dans le nombre approprié de puits dans une plaque de microtitration. REMARQUE: Un puits d&#39;une microplaque pourra accueillir environ 4 paires de disques oculaires attachés à la coque de nymphose ou 8 paires isolées de disques oculaires avec des cerveaux connectés. Les anticorps primaires utilisés dans ce protocole pour les données représentatives sont antiphosphotyrosine (dilution 1: 500) et anti-Cut (dilution 1: 200). </li><li> Transférer le tissu disséqué dans les puits de plaques de microtitration remplies de la solution d&#39;anticorps primaire de la boîte de verre de 3 puits. </li><li> Lieula microplaque sur une serviette en papier mouillée (pour prévenir la déshydratation) dans une boîte vide de la pipette de pointe (ou d&#39;autres contenants similaires) et conserver à 4 ° C pendant la nuit. </li></ol> Jour 2 (4 h + temps de montage) <ol start="6"><li> Transférer le tissu de la microplaque à un nouveau plat en verre de 3 et de 0,3% de PBT (≈0.5 ml) et laver le tissu pendant 10 min sur la bascule. Répétez cette étape trois fois pour un total de 4 lavages (10 min chacun) </li><li> Retirez le dernier lavage et ajouter 0,5 ml de solution de blocs pour le bien. Bloquer pendant 30 min. NOTE: Ce délai peut être prolongé en fonction de vos besoins. </li><li> Retirer la solution de bloc et ajouter 0,5 ml de la solution d&#39;anticorps secondaire (fait à partir d&#39;anticorps marqués par fluorescence correspondant à votre anticorps primaire et longueur d&#39;onde fluorescente, dilué dans une solution Block) pour le bien et protéger le plat en verre de 3 et de la lumière avec une couverture ou aluminium déjouer et incuber pendant au moins 2 h à la salletempérature. NOTE: Toutes les étapes suivantes devraient également être protégés contre l&#39;exposition de la lumière. Cette incubation peut être prolongée pour une nuit à 4 ° C, dans une configuration similaire à l&#39;incubation d&#39;anticorps primaire, mais il peut également être effectuée dans un récipient en verre 3 de puits recouvert d&#39;un film de laboratoire, ou une plaque de microtitration, si la conservation de l&#39;anticorps est souhaitée. </li><li> À la fin de l&#39;incubation d&#39;anticorps, enlever la solution d&#39;anticorps secondaire et ajouter 0,5 ml de 0,3% PBT pour le bien et laver sur la bascule pour 10 min. Répétez cette étape pour un total de 4 lavages (10 min chacun). NOTE: Si la coloration DAPI est souhaité, avant de commencer le premier lavage, ajouter 0,5 pi de solution DAPI des stocks (voir matériaux) pour les 500 pi de PBT qui seront utilisés pour le premier lavage (1: 1000 dilution). </li></ol> 3. Montage <ol><li> A la fin de la dernière lavage, monter les disques yeux des antennes / sur une diapositive en utilisant 70% de glycérol ou PBS comme un moyen de supprimer le largeain et d&#39;autres tissus non désirés dans une piscine d&#39;un côté de la lame. </li><li> Déplacez doucement le tissu en utilisant la pression hydrostatique entre les forceps, ou très doucement avec une pointe forceps, et aligner les disques d&#39;oeil dans une colonne vers le centre de la diapositive. </li><li> Retirer tous glycérol étrangère ou PBS à travers le tissu (en utilisant effet de mèche d&#39;un laboratoire lingette ou les forceps travailler pour cela). </li><li> Placer une petite quantité (≈10 pi) de milieu de montage sur un côté de la colonne de disque de l&#39;oeil et placer une lamelle couvre-objet sur le tissu de façon à répartir le milieu de montage sur les disques de l&#39;oeil. </li><li> Laissez diapositive sit pendant 1 à 2 minutes pour permettre le / milieu de montage de tissu pour étaler complètement. </li><li> Essuyez tout support de montage de congé supplémentaire autour des bords de la lamelle et le sceller avec du vernis à ongles transparent. </li><li> Capturer des images fluorescentes des disques de l&#39;oeil à l&#39;aide de la microscopie confocale. </li></ol>

<ol>
	<li>Ready, D. F., Hanson, T. E., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+Drosophila+retina,+a+neurocrystalline+lattice.">Development of the Drosophila retina, a neurocrystalline lattice.</a> Dev Biol. 53 (2), 217-240 (1976).</li><li>Wolff, T., Ready, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+beginning+of+pattern+formation+in+the+Drosophila+compound+eye:+the+morphogenetic+furrow+and+the+second+mitotic+wave.">The beginning of pattern formation in the Drosophila compound eye: the morphogenetic furrow and the second mitotic wave.</a> Development. 113 (3), 841-850 (1991).</li><li>Carthew, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pattern+formation+in+the+Drosophila+eye.">Pattern formation in the Drosophila eye.</a> Curr Opin Genet Dev. 17 (4), 309-313 (2007).</li><li>Bao, S., Cagan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preferential+adhesion+mediated+by+Hibris+and+Roughest+regulates+morphogenesis+and+patterning+in+the+Drosophila+eye.">Preferential adhesion mediated by Hibris and Roughest regulates morphogenesis and patterning in the Drosophila eye.</a> Dev cell. 8 (6), 925-935 (2005).</li><li>Grzeschik, N. A., Knust, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=IrreC/rst-mediated+cell+sorting+during+Drosophila+pupal+eye+development+depends+on+proper+localisation+of+DE-cadherin.">IrreC/rst-mediated cell sorting during Drosophila pupal eye development depends on proper localisation of DE-cadherin.</a> Development. 132 (9), 2035-2045 (2005).</li><li>Sprecher, S. G., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Switch+of+rhodopsin+expression+in+terminally+differentiated+Drosophila+sensory+neurons.">Switch of rhodopsin expression in terminally differentiated Drosophila sensory neurons.</a> Nature. 454 (7203), 533-537 (2008).</li><li>Zuker, C. S., Cowman, A. F., Rubin, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+structure+of+a+rhodopsin+gene+from+D.+melanogaster.">Isolation and structure of a rhodopsin gene from D. melanogaster.</a> Cell. 40 (4), 851-858 (1985).</li><li>Voas, M. G., Rebay, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signal+integration+during+development:+insights+from+the+Drosophila+eye.">Signal integration during development: insights from the Drosophila eye.</a> Dev Dyn. 229 (1), 162-175 (2004).</li><li>Hsiao, H. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+and+immunohistochemistry+of+larval,+pupal+and+adult+Drosophila+retinas.">Dissection and immunohistochemistry of larval, pupal and adult Drosophila retinas.</a> J Vis Exp. , 4347 (2012).</li><li>Call, G. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genomewide+clonal+analysis+of+lethal+mutations+in+the+Drosophila+melanogaster+eye:+comparison+of+the+X+chromosome+and+autosomes.">Genomewide clonal analysis of lethal mutations in the Drosophila melanogaster eye: comparison of the X chromosome and autosomes.</a> Genetics. 177 (2), 689-697 (2007).</li><li>Chen, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discovery-based+science+education:+functional+genomic+dissection+in+Drosophila+by+undergraduate+researchers.">Discovery-based science education: functional genomic dissection in Drosophila by undergraduate researchers.</a> PLoS Biol. 3 (2), 59 (2005).</li><li>Wolff, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+techniques+for+pupal+and+larval+Drosophila+eyes.">Dissection techniques for pupal and larval Drosophila eyes.</a> CSH Protoc. 2007, (2007).</li><li>Williamson, W. R., Hiesinger, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+developing+and+adult+Drosophila+brains+and+retinae+for+live+imaging.">Preparation of developing and adult Drosophila brains and retinae for live imaging.</a> J Vis Exp. 37, 1936 (2010).</li><li>Xu, T., Rubin, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+genetic+mosaics+in+developing+and+adult+Drosophila+tissues.">Analysis of genetic mosaics in developing and adult Drosophila tissues.</a> Development. 117 (4), 1223-1237 (1993).</li><li>Duffy, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GAL4+system+in+Drosophila:+a+fly+geneticist's+Swiss+army+knife.">GAL4 system in Drosophila: a fly geneticist's Swiss army knife.</a> Genesis. 34 (1-2), 1-15 (2002).</li></ol>

The authors have nothing to disclose.

While it appears that the process is simple and easy to perform, in reality, this technique requires a great deal of practice to master. Routinely, we start students off by learning to dissect and mount third instar eye discs12, which are much easier to work with. This practice helps to develop an appropriate dissection position of the arms, hands and fingers13 so that manipulation of the forceps under the dissecting microscope is stable, easy and experienced. In essence, the practice period shou...

Les domaines de la biologie du développement et de la cellule ont été fortement touchés par l&#39;organisme modèle: Drosophila melanogaster. Dans ce modèle, les études du disque de l&#39;œil ont contribué beaucoup de connaissances sur la signalisation, biologie cellulaire et d&#39;autres domaines. Le troisième disque stade de l&#39;œil larvaire fin a été largement étudié et est un modèle puissant à utiliser, car il donne un aperçu d&#39;une série de périodes de développement, chacun avec ses propres molécules et des processus de signalisation unique, comme le sillon morphogénétique progresse à travers le disque de l&#39;oeil 8. Cependant, il est nécessaire d&#39;étendre notre compréhension des processus de développement dans la phase de chrysalide de développement. Bien qu&#39;il y ait eu des études sur le disque d&#39;œil pupe 3-7, notre connaissance ne se rapproche pas de l&#39;ampleur du travail qui a été effectué sur le troisième disque stade de l&#39;œil. Cela est dû, en partie, à la plus grande difficulté à disséquer le disque d&#39;oeil chrysalide. En conséquence, une présentation de labonne méthode de dissection pourrait grandement augmenter la recherche dans ce domaine. Bien qu&#39;il existe des étapes dans le développement du disque de l&#39;œil des pupes qui sont facilement disséqués, en particulier autour de la période mi-pupe, d&#39;autres périodes sont beaucoup plus difficiles à disséquer. Ce protocole représente une méthode pour disséquer disques oculaires de nymphose qui peuvent être universellement utilisés pour tous les délais de développement des pupes. Ce protocole peut être utilisé comme une alternative à un autre protocole que 9 montre un procédé plus facile et plus rapide à disséquer les disques de l&#39;oeil à partir des points de temps midpupal. Ce protocole a été filmé et mis au point pour la formation des étudiants avancés de premier cycle dans le cycle UCLA Consortium de recherche en génomique fonctionnelle (URCFG) 10,11 dans la technique de dissection de l&#39;oeil chrysalide. Beaucoup d&#39;étudiants de premier cycle ont pu utiliser cette vidéo et méthode pour apprendre cette technique difficile.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Phosphate-buffered saline (PBS, pH 7.4)</td>
			<td></td>
			<td></td>
			<td>80g NaCl, 2g&#160; KCl, 14.4g Na2HPO4, 2.4g KH2PO4, Bring volume to 1 l, adjust the pH to 7.4, autoclave or filter sterilize, dilute to 1X PBS with autoclaved ddH2O before using.</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Promega</td>
			<td>H5142</td>
			<td>Caution: Irritant! Wear gloves.</td>
		</tr>
		<tr>
			<td>0.3% PBT</td>
			<td></td>
			<td></td>
			<td>1.5 ml of Triton X-100, 500 ml 1X PBS.</td>
		</tr>
		<tr>
			<td>37% formaldehyde solution</td>
			<td>Fisher Scientific</td>
			<td>F75P1GAL</td>
			<td>Caution: Toxic, probable human carcinogen! Wear gloves.&#160;</td>
		</tr>
		<tr>
			<td>Fix Solution</td>
			<td></td>
			<td></td>
			<td>(&#8776;4% Formaldehyde in PBS) 50 &#956;l of 37% Formaldehyde, 450 &#956;l 1XPBS, make fresh before use</td>
		</tr>
		<tr>
			<td>Normal goat serum</td>
			<td>Rockland antibodies &#38; assays</td>
			<td>B304</td>
			<td>Aliquot in 1 ml volumes and store at -80C</td>
		</tr>
		<tr>
			<td>Block Solution</td>
			<td></td>
			<td></td>
			<td>10% NGS in PBT.&#160; This can be made and stored at 4 &#176;C for a few days prior to use.</td>
		</tr>
		<tr>
			<td>DAPI stock solution</td>
			<td>Life Technologies</td>
			<td>D3571</td>
			<td>For coutnerstaining nuclei. Prepare a 1 mg/ml solution with ddH2O.</td>
		</tr>
		<tr>
			<td>VectaShield Mounting Medium</td>
			<td>Vector Labs</td>
			<td>H-1000</td>
			<td>Mounting medium</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sigma</td>
			<td>G5516</td>
			<td>For mounting. Prepare 70% dilution with ddH2O.</td>
		</tr>
		<tr>
			<td colspan="4">Equipment</td>
		</tr>
		<tr>
			<td>Nutating mixer</td>
			<td>VWR</td>
			<td>82007-202</td>
			<td>Used to rock tissue in 3 well glass dish</td>
		</tr>
		<tr>
			<td>SylGard 182 Silicone Elastomer Kit</td>
			<td>Krayden</td>
			<td>NC9897184</td>
			<td>Used to make silicone dissection dish</td>
		</tr>
		<tr>
			<td>Silicone dissecting dish</td>
			<td></td>
			<td></td>
			<td>Mix Sylgard elastomer kit (above) according to directions gently (to avoid bubbles). Pour mixture into Petri dish (any size). Allow SylGard to cure overnight in 37 &#176;C incubator.</td>
		</tr>
		<tr>
			<td>3 well glass dish</td>
			<td>Corning</td>
			<td>7220-85</td>
			<td>The 3 well variety of these are no longer available, this is the 9 well product.</td>
		</tr>
		<tr>
			<td>72 well microwell minitray</td>
			<td>Nunc</td>
			<td>438733</td>
			<td></td>
		</tr>
		<tr>
			<td>Sharp forceps (Dumont #55)</td>
			<td>Fine Science Tools</td>
			<td>11255-20</td>
			<td></td>
		</tr>
		<tr>
			<td><a name="RANGE!A19">Vannas-type Micro Scissors, Straight, 5mm blade</a></td>
			<td>Ted Pella</td>
			<td>1346</td>
			<td></td>
		</tr>
		<tr>
			<td>100 mm Borosilicate glass capillaries</td>
			<td>World Precision Instruments</td>
			<td>1B100-4</td>
			<td>Pull with needle puller to make fine point tip that allows a small stream of PBS to flow.</td>
		</tr>
		<tr>
			<td>Disposable Transfer Pipets, Fine Tip</td>
			<td>Samco Scientific</td>
			<td>231</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">Tubing dimensions given are inner diameter (ID) x outer diameter (OD) x wall thickness in inches</td>
		</tr>
		<tr>
			<td>PVC tubing (1/8 x 3/16 x 1/32)</td>
			<td>Nalgene</td>
			<td>8000-0010</td>
			<td rowspan="3">Use these with pulled needle to assemble the blower tube as shown in Figure 2.</td>
		</tr>
		<tr>
			<td>Tygon Silicone tubing (3/32 x 5/32 x 1/32)</td>
			<td>Saint Gobain Performance Plastics</td>
			<td>ABW00004</td>
		</tr>
		<tr>
			<td>Tygon Silicone tubing (1/32 x 3/32 x 1/32)</td>
			<td>Saint Gobain Performance Plastics</td>
			<td>ABW00001</td>
		</tr>
	</tbody>
</table>

dissection and mounting of drosophila pupal eye discs

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes. It contains approximately 800 separate eye units, termed ommatidia1. Each ommatidium contains eight neuronal photoreceptors that develop from undifferentiated cells following the passage of the morphogenetic furrow in the third larval instar2. Following the sequential differentiation of the photoreceptors, non-neuronal cells develop, including cone and pigment cells, along with mechanosensory bristle cells3. Final differentiation processes, including the structured arrangement of all the ommatidial cell types, programmed cell death of undifferentiated cell types and rhodopsin expression, occurs through the pupal phase4-7. This technique focuses on manipulating the pupal eye disc, providing insight and instruction on how to dissect the eye disc during the pupal phase, which is inherently more difficult to perform than the commonly dissected third instar eye disc. This technique also provides details on immunostaining to allow the visualization of various proteins and other cell components.

A titre d&#39;exemple de l&#39;utilisation de ce protocole, des résultats illustrant midpupal (42 heures APF à 25 ° C) des disques de l&#39;oeil immunocolorées avec des anticorps différents sont présentés dans la figure 2. En utilisant un anticorps dirigé contre les résidus de phosphotyrosine, la membrane des cellules peut être observée (figure 2A). Ceci peut être utilisé pour identifier l&#39;agencement régulier de cellules dans l&#39;œil ommatidial nymphal suivant les procédés de mise...

Watch this Scientific Journal Video about Dissection and Mounting of Drosophila Pupal Eye Discs at JoVE.com

Dissection and Mounting of Drosophila Pupal Eye Discs

The goal of this technique is to enable researchers to perform dissection, immunostaining and mounting of pupal eye discs from Drosophila melanogaster of any age.

Dissection et de montage de<em&gt; Drosophila</em&gt; pupes disques yeux

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes.  It contains approximately 800 ...

dissection-and-mounting-of-drosophila-pupal-eye-discs

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Biology

Dissection and Mounting of Drosophila Pupal Eye Discs

12.1K vues.  University of California at Los Angeles (UCLA). The goal of this technique is to enable researchers to perform dissection, immunostaining and mounting of pupal eye discs from Drosophila melanogaster of any age.

Vidéo: Dissection and Mounting of Drosophila Pupal Eye Discs

Dissection of Drosophila Ovaries

Dissection des ovaires chez la drosophile

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Biologie

Dissection of Imaginal Discs from 3rd Instar Drosophila Larvae

This protocol demonstrates the dissection technique used for isolating imaginal discs from drosophila larvae at the 3rd instar stage. Methods for fixing the tissue after saying and removing the wing, leg, and eye discs are demonstrated directly on a microscope slide for subsequent visualization.

Dissection de disques imaginaux de la 3ème Instar larves de drosophile

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

Drosophile larvaire JNM Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Dissection et la coloration des Drosophile Ovaires larvaires

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

L&#39;immunohistochimie drosophile Abdomen Nymphe

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

Imagerie à travers le cas de la nymphe<em&gt; Drosophila melanogaster</em

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called imaginal discs. These discs give rise to a high percentage of adult structures that are found within the adult fly. Here we describe a protocol that has been optimized to recover these discs and prepare them for analysis with antibodies, transcriptional reporters and protein traps. This procedure is best suited for thin tissues like imaginal discs, but can be easily modified for use with thicker tissues such as the larval brain and adult ovary. The written protocol and accompanying video will guide the reader/viewer through the dissection of third instar larvae, fixation of tissue, and treatment of imaginal discs with antibodies. The protocol can be used to dissect imaginal discs from younger first and second instar larvae as well. The advantage of this protocol is that it is relatively short and it has been optimized for the high quality preservation of the dissected tissue. Another advantage is that the fixation procedure that is employed works well with the overwhelming number of antibodies that recognize Drosophila proteins. In our experience, there is a very small number of sensitive antibodies that do not work well with this procedure. In these situations, the remedy appears to be to use an alternate fixation cocktail while continuing to follow the guidelines that we have set forth for the dissection steps and antibody incubations.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

Dissection et Immunocoloration Imaginal disques de<em&gt; Drosophila melanogaster</em

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called ...

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes differentiation from a stem cell population, mitotic amplification, and meiosis. In addition, post-meiotic germ cells undergo a dramatic morphological reshaping process as well as a global epigenetic reconfiguration of the germ line chromatin&#8212;the histone-to-protamine switch.
Studying the role of a protein in post-meiotic spermatogenesis using mutagenesis or other genetic tools is often impeded by essential embryonic, pre-meiotic, or meiotic functions of the protein under investigation. The post-meiotic phenotype of a mutant of such a protein could be obscured through an earlier developmental block, or the interpretation of the phenotype could be complicated. The model organism Drosophila melanogaster offers a bypass to this problem: intact testes and even cysts of germ cells dissected from early pupae are able to develop ex vivo in culture medium. Making use of such cultures allows microscopic imaging of living germ cells in testes and of germ-line cysts. Importantly, the cultivated testes and germ cells also become accessible to pharmacological inhibitors, thereby permitting manipulation of enzymatic functions during spermatogenesis, including post-meiotic stages.
The protocol presented describes how to dissect and cultivate pupal testes and germ-line cysts. Information on the development of pupal testes and culture conditions are provided alongside microscope imaging data of live testes and germ-line cysts in culture. We also describe a pharmacological assay to study post-meiotic spermatogenesis, exemplified by an assay targeting the histone-to-protamine switch using the histone acetyltransferase inhibitor anacardic acid. In principle, this cultivation method could be adapted to address many other research questions in pre- and post-meiotic spermatogenesis.

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

This protocol describes the dissection and cultivation of intact testes and germ-line cysts from Drosophila melanogaster pupae. This method allows microscopic observation of spermatogenesis ex vivo. Furthermore, we describe a pharmacological assay of the effect of inhibitors on specific stages of germ-cell development in pupal testes.

Ex vivo Culture de la drosophile pupes testicule et seul mâle kystes lignée germinale: Dissection, l&#39;imagerie et le traitement pharmacologique

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes ...

Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton

The posterior pharyngeal segments of the vertebrate head give rise to the branchial skeleton, the primary site of food processing in fish. The morphology of the fish branchial skeleton is matched to a species' diet. Threespine stickleback fish (Gasterosteus aculeatus) have emerged as a model system to study the genetic and developmental basis of evolved differences in a variety of traits. Marine populations of sticklebacks have repeatedly colonized countless new freshwater lakes and creeks. Adaptation to the new diet in these freshwater environments likely underlies a series of craniofacial changes that have evolved repeatedly in independently derived freshwater populations. These include three major patterning changes to the branchial skeleton: reductions in the number and length of gill raker bones, increases in pharyngeal tooth number, and increased branchial bone lengths. Here we describe a detailed protocol to dissect and flat-mount the internal branchial skeleton in threespine stickleback fish. Dissection of the entire three-dimensional branchial skeleton and mounting it flat into a largely two-dimensional prep allows for the easy visualization and quantification of branchial skeleton morphology. This dissection method is inexpensive, fast, relatively easy, and applicable to a wide variety of fish species. In sticklebacks, this efficient method allows the quantification of skeletal morphology in genetic crosses to map genomic regions controlling craniofacial patterning.

The branchial skeleton, including gill rakers, pharyngeal teeth, and branchial bones, serves as the primary site of food processing in most fish. Here we describe a protocol to dissect and flat-mount this internal skeleton in threespine sticklebacks. This method is also applicable to a variety of other fish species.

Dissection et Flat-montage de l&#39;épinoche à trois épines branchiale Squelette

The posterior pharyngeal segments of the vertebrate head give rise to the branchial skeleton, the primary site of food processing in fish. The morphology ...

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Diversified species of Drosophila (fruit fly) provide opportunities to study mechanisms of development and genetic changes responsible for evolutionary changes. In particular, the adult stage is a rich source of morphological traits for interspecific comparison, including wing pigmentation comparison. To study developmental differences among species, detailed observation and appropriate staging are required for precise comparison. Here we describe protocols for staging of pupal periods and quantification of wing pigmentation in a polka-dotted fruit fly, Drosophila guttifera. First, we describe the method for detailed morphological observation and definition of pupal stages based on morphologies. This method includes a technique for removing the puparium, which is the outer chitinous case of the pupa, to enable detailed observation of pupal morphologies. Second, we describe the method for measuring the duration of defined pupal stages. Finally, we describe the method for quantification of wing pigmentation based on image analysis using digital images and ImageJ software. With these methods, we can establish a solid basis for comparing developmental processes of adult traits during pupal stages.

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Protocols for staging pupal periods and measurement of wing pigmentation of Drosophila guttifera are described. Staging and quantification of pigmentation provide a solid basis for studying developmental mechanisms of adult traits and enable interspecific comparison of trait development.

Méthodes pour la mise en scène des périodes nymphale et mesure de Pigmentation de l’aile de Drosophila guttifera

Diversified species of Drosophila (fruit fly) provide opportunities to study mechanisms of development and genetic changes responsible for evolutionary ...