Nous remercions Mme V. Lephart pour l&#39;entretien des stocks de volée Ce travail a été soutenu par les Instituts nationaux de la santé de subvention (GM62916) et la National Science Foundation subvention (MCB0817107).

Le protocole suivant pour la préparation des chromosomes polytènes de squash est adaptée de la procédure décrite dans Johansen et al. (2009). 1. Culture de larves de drosophile troisième stade Afin d&#39;obtenir optimale chromosomes polytènes de haute qualité des préparations de squash, des conditions de culture sont essentiels fréquentés (c&#39;est à dire, placer environ 20 ponte des mouches femelles dans une norme de 4 bouteille volée &quot;et changer pour une nouvelle bouteille chaque jour). Sélectionnez la plus grosse des individus de la première récolte de l&#39;escalade troisième stade larvaire pendant qu&#39;ils sont encore nomades, mais juste avant la nymphose. Nous avons régulièrement la culture à 21 ° C mais 18 ° C donnera plus gros chromosomes qui peuvent être plus adaptés à certaines fins, comme par exemple lorsque la bande / interbandes régions ont besoin d&#39;être visualisés à haute résolution. 2. Matériaux de squash polytènes <ul><li> Pinces à dissection Drummond (2) </li><li> Boîtes de Pétri (60 x 15 mm) </li><li> Lames de microscope givré (n ° 12-544-3 Fisher) (poly-lysine) </li><li> 22 x 22 mm glisse n ° couvrent 15 (Fisher n ° 12-520B) (enduits de Sigmacote; Sigma # SL2) </li><li> 22 x 40 mm glisse n ° couvrent 15 (Fisher n ° 12-530B) </li><li> Kim-lingettes </li><li> Microscope à contraste de phase avec l&#39;objectif 20X </li><li> Petite Dewar (par exemple, fiole à vide ou un thermos bouteille) </li><li> Pince longue </li><li> Lames de rasoir </li><li> Coplin Jarre </li><li> Caoutchouc-Maid ou le plateau de Tupperware (ou équivalent Bac plombable) </li><li> Parafilm (coupé en carrés 22 mm) </li><li> 175 g de poids </li></ul> 3. Fixatifs et solutions <ol><li> Stocks de formaldéhyde 5X. Préparer une nouvelle solution de 0,74 g. paraformaldéhyde dans 4,0 ml de dH 2 O et avec 28 l de KOH 1N. Chaude à 65 ° C pour dissoudre le paraformaldéhyde et stocker ensuite sur la glace. </li><li> Fixateur 1. Préparer une nouvelle solution de 0,5 ml PBS 10X, 50 pl Triton X-100, 3,45 ml dH 2 O et 1,0 ml d&#39;un stock de formaldéhyde 5X. Chaud pour disperser le Triton X-100 et l&#39;utilisation en 1 heure. </li><li> Fixateur 2. Préparer une nouvelle solution de 1,5 ml dH 2 O, 2,5 ml d&#39;acide acétique glacial et 1,0 ml d&#39;un stock de formaldéhyde 5X et d&#39;utiliser moins de 1 heure. </li><li> Solution d&#39;acide Lactoacetic. Préparer une solution de 1 ml d&#39;acide lactique, 2 ml dH 2 O, et 3 ml d&#39;acide acétique. </li></ol> 4. Préparation polytènes de squash chromosome: <ol><li> Rincer à l&#39;eau et les larves de transfert à PBS dans une boîte de culture tissulaire pour la dissection. </li><li> Saisir l&#39;extrémité de la bouche des crochets avec une paire de pinces, tenir le corps environ 2 / 3 du chemin vers le bas avec l&#39;autre paire, et tirez sur la bouche des crochets de sorte les glandes salivaires sont exposés. Séparer les glandes salivaires du cerveau et des yeux antennaire disques, et de disséquer le cadavre de graisse et les tissus associés par les glandes. </li><li> Ajouter 200 microlitres de fixateur 1 dans le premier puits d&#39;une lame de deux dépression bien et 200-300 microlitres de 2 fixatif pour bien deux. Transfert une paire de glandes salivaires à la fois pour une même fixateur dans un et incuber pendant le temps requis pour l&#39;épitope cible, habituellement environ 1-2 minutes. (Remarque: certains épitopes, comme pour la plupart des modifications des histones, peuvent nécessiter 5 minutes ou plus de fixation.) </li><li> Pince en utilisant le transfert des glandes salivaires à 2 fixateur dans le puits 2 et incuber pendant deux minutes. </li><li> Transfert des glandes à 10-30 ul de solution d&#39;acide Lactoacetic sur une lamelle propre Sigmacoted. Abaissez délicatement une lame de microscope polylysine enduit sur ​​la lamelle et, sans appliquer une pression verticale, ramasser la lamelle afin glandes sont entre la lame et la lamelle. Immédiatement faciliter la lyse cellulaire et chromosomique propagation en prenant soin saisir la lamelle avec une pince sur un bord et doucement se déplaçant légèrement en arrière, en essayant de minimiser toute pression verticale sur le tissu. Sinon, utiliser le côté gomme d&#39;un crayon ou même un doigt pour déplacer doucement la lamelle en arrière. Notez que tout retard dans le déplacement de la lamelle en arrière est susceptible de diminuer la propagation des armes chromosomiques comme ils auront tendance à devenir plus rigides à l&#39;exposition à la solution d&#39;acide Lactoacetic. Opacité de la solution est souvent une bonne indication de la séparation des cellules. Tapotant doucement la lamelle oblique (pour éviter une pression verticale) avec le côté gomme d&#39;un crayon à quelques reprises peut aussi aider à propager chromosomiques. </li><li> Examine immédiatement le tissu sous un microscope à contraste de phase avec un objectif 20 ou 40x pour déterminer si les chromosomes sont bien réparties. </li><li> Lorsque la propagation chromosomique est suffisant, mettre la diapositive avec le côté lamelle sur une pile de nettoyage Kim-lingettes. Placez une seconde Kim-tampon sur le dessus et aplatir les chromosomes en plaçant votre pouce sur l&#39;endroit où la lamelle est positionné et en appuyant fermement, en évitant tout mouvement horizontal de la lamelle qui cisaillementles chromosomes. </li><li> Examinez la lame à nouveau sous le microscope afin de déterminer si la préparation est adaptée pour l&#39;usage prévu. Si les chromosomes ont déménagé de façon significative sur les écraser, utiliser moins de volume de solution d&#39;acide Lactoacetic dans vos préparations ultérieures. Répétez les étapes 4.1 à 4.8 jusqu&#39;à ce qu&#39;un nombre suffisant de diapositives appropriées ont été obtenues. Placez 175 g de poids sur la lamelle de la glisse. </li><li> Remplir un petit Dewar (par exemple, une bouteille thermos) à l&#39;azote liquide. En utilisant une pince à long, tremper glisser dans l&#39;azote liquide 2 jusqu&#39;à l&#39;arrêt ébullition, retirez diapositives et utiliser immédiatement le bord d&#39;une lame de rasoir propre à un coin flip off à la lamelle. Si diapositive sera utilisée immédiatement, placer dans un bocal de Coplin avec du PBS à 4 ° C. Sinon recueillir les glisser dans une jarre de Coplin remplie d&#39;éthanol à 95%. Examiner la lamelle une fois le gel est sublimé pour confirmer que le tissu adhère à la diapositive et ne pas rester avec la lamelle. Répéter avec le reste de la glisse jusqu&#39;à ce que toutes les diapositives sont stockées soit dans du PBS (pour une utilisation au sein de plusieurs heures) ou de l&#39;éthanol (pour le stockage à long terme). </li></ol> Les résultats représentatifs La première étape critique pour l&#39;obtention de haute qualité de la préparation de squash polytènes est de faire croître les larves grasses avec des noyaux des glandes salivaires grands. La seconde est une bonne technique à la procédure épandage, qui peut prendre un peu de pratique. Une astuce pour réussir la propagation améliorée est de trouver le volume minimal de solution d&#39;acide lactoacetic pendant l&#39;étape de l&#39;écrasement. Cela favorisera la propagation suffisante de chromosomes sans générer des forces excessives streaming qui peuvent se laver les bras du chromosome loin. Il est également intéressant de noter que tout retard dans le déplacement de la lamelle en arrière permettra de réduire la propagation des armes chromosomiques, ils deviennent plus rigides lorsqu&#39;ils sont exposés à la solution d&#39;acide Lactoacetic. Si tout va bien, il devrait y avoir de nombreux bien répartis chromosomes polytènes. La figure 1 montre un exemple d&#39;une telle préparation, double marqué avec un marqueur pour les régions interbandes en vert et avec un colorant qui colore les régions bagué en bleu. Si l&#39;insuffisance d&#39;épandage est obtenue, les chromosomes se ressemblent petites boules, comme le montre la figure 2. D&#39;autre part, si une trop grande propagation a eu lieu, le chomosomes sera trop mince et étendu ou, dans certains cas, fragmentée en petits morceaux comme le montre la figure 3. <img src="/files/ftp_upload/1748/1748fig1.jpg" alt="Figure 1" /> Figure 1. Préparation de squash polytènes doubles étiquetés avec l&#39;anticorps de la JIL-1 histone kinase H3S10 (en rouge) et Hoechst (bleu). <img src="/files/ftp_upload/1748/1748fig2.jpg" alt="Figure 2" /> Figure 2. Courges polytènes avec la propagation insuffisante. <img src="/files/ftp_upload/1748/1748fig3.jpg" alt="Figure 3" /> Figure 3. Courges polytènes avec trop de propagation.

<ol>
	<li>Cai, W., Bao, X., Deng, H., Jin, Y., Girton, J., Johansen, J., Johansen, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+polymerase+II-mediated+transcription+at+active+loci+does+not+require+histone+H3S10+phosphorylation+in+Drosophila.">RNA polymerase II-mediated transcription at active loci does not require histone H3S10 phosphorylation in Drosophila.</a> Development. 135, 2917-2925 (2008).</li><li>DiMario, P., Rosby, R., Cui, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+visualization+of+GFP-fusion+proteins+on+polytene+chromosomes.">Direct visualization of GFP-fusion proteins on polytene chromosomes.</a> Dros. Inf. Serv. 89, 115-118 (2006).</li><li>Johansen, K. M., Cai, W., Deng, H., Bao, X., Zhang, W., Girton, J., Johansen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+studying+transcription+and+epigenetic+chromatin+modification+in+Drosophila+polytene+chromosome+squash+preparations+using+antibodies.">Methods for studying transcription and epigenetic chromatin modification in Drosophila polytene chromosome squash preparations using antibodies.</a> Methods. 48, 387-397 (2009).</li></ol>

L&#39;inclusion d&#39;acides acétique et lactique dans les protocoles conventionnels de fixation facilite la résolution de squash et deux interbandes bras chromosomique propagation mais malheureusement, certains épitopes ne survivent pas à ce traitement. Un exemple d&#39;un tel épitope est H3S10ph (Cai et al., 2008). Depuis traitement à l&#39;acide a aussi l&#39;inconvénient qu&#39;il désaltère la fluorescence intrinsèque des protéines étiquetées GFP, DiMario et al. (2006) a récemment dé...

preparation of drosophila polytene chromosome squashes for antibody labeling

Drosophile a longtemps été un modèle de système de prédilection pour étudier la relation entre la structure de la chromatine et la régulation des gènes en raison des avantages cytologique fournie par le géant de chromosomes polytènes salivaires des glandes du stade larvaire tiers. Dans ce tissu les chromosomes subissent de nombreux cycles de réplication, en l&#39;absence de la division cellulaire donnant naissance à environ 1000 exemplaires. L&#39;ADN reste alignée après chaque cycle réplicatif résultant dans des chromosomes fortement agrandie qui fournissent une occasion unique de mettre en corrélation la morphologie chromatine avec la localisation des protéines spécifiques. En conséquence, il a été un haut niveau d&#39;intérêt dans la définition des modifications épigénétiques présents à des gènes différents et à différentes étapes du processus de transcription. Un outil important pour de telles études est l&#39;étiquetage des chromosomes polytènes avec des anticorps à l&#39;enzyme, facteur de transcription, ou la modification des histones d&#39;intérêt. Ce protocole vidéo illustre la technique de squash utilisés dans le laboratoire de Johansen pour préparer la drosophile chromosomes polytènes pour l&#39;étiquetage des anticorps.

Watch this Scientific Journal Video about Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling at JoVE.com

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Ce protocole vidéo illustre la technique de squash utilisés dans le laboratoire de Johansen pour préparer<em&gt; Drosophile</em&gt; Chromosomes polytènes pour l&#39;étiquetage des anticorps.

Préparation des Courges drosophile chromosomes polytènes pour marquage de l&#39;anticorps

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological ...

preparation-drosophila-polytene-chromosome-squashes-for-antibody

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31.0K vues.  Iowa State University. Ce protocole vidéo illustre la technique de squash utilisés dans le laboratoire de Johansen pour préparer Drosophile Chromosomes polytènes pour l'étiquetage des anticorps.

Vidéo: Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

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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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 ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

Marquage fluorescent Drosophile</emOuvrages d&#39;art> Coeur

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

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 ...

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

Dissection en direct de Drosophile Embryons: méthodes simplifiées pour le dépistage de collections de mutants par immunofluorescence

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

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 ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

Préparation des Drosophile Les cellules S2 pour la microscopie optique

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Préparation des adultes Drosophile Yeux pour la coupe mince et analyse microscopique

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

Etudier Checkpoint mitotique en illustrant le comportement dynamique des protéines Kinetochore et Motion chromosomique chez Vivre<em&gt; Drosophila</em&gt; Les embryons syncytial

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Chromosome Preparation From Cultured Cells

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as fluorescence in situ hybridization (FISH), this assay has the powerful ability to analyze individual cells for aberrations that involve gains or losses of portions of the genome&#160;and rearrangements involving one or more chromosomes. In humans, chromosome abnormalities occur in approximately 1 per 160 live births1,2, 60-80% of all miscarriages3,4, 10% of stillbirths2,5, 13% of individuals with congenital heart disease6, 3-6% of infertility cases2, and in many patients with developmental delay&#160;and birth defects7. Cytogenetic analysis of malignancy is routinely used by researchers and clinicians, as observations of clonal chromosomal abnormalities have been shown to have both diagnostic and prognostic significance8,9.&#160; Chromosome isolation is invaluable for gene therapy and stem cell research of organisms including nonhuman primates and rodents10-13.
Chromosomes can be isolated from cells of live tissues, including blood lymphocytes, skin fibroblasts, amniocytes, placenta, bone marrow, and tumor specimens. Chromosomes are analyzed at the metaphase stage of mitosis, when they are most condensed and therefore more clearly visible. The first step of the chromosome isolation technique involves the disruption of the spindle fibers by incubation with Colcemid, to prevent the cells from proceeding to the subsequent anaphase stage. The cells are then treated with a hypotonic solution and preserved in their swollen state with Carnoy&#39;s fixative. The cells are then dropped on to slides and can then be utilized for a variety of procedures. G-banding involves trypsin treatment followed by staining with Giemsa to create characteristic light and dark bands. The same procedure to isolate chromosomes can be used for the preparation of cells for procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY)14,15.

Chromosomes can be isolated from live cells such as lymphocytes or skin fibroblasts, and from organisms including humans or mice. These chromosome preparations can be further utilized for routine G-banding and molecular cytogenetic procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY).

Chromosome Préparation partir des cellules cultivées

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as ...