Ringraziamo la signora V. Lephart di mantenimento delle scorte volare Questo lavoro è stato sostenuto dal National Institutes for Health di sovvenzione (GM62916) e la National Science Foundation di sovvenzione (MCB0817107).

Il protocollo seguito per la preparazione di politene da squash cromosoma è adattato dalla procedura descritta in Johansen et al. (2009). 1. Cultura del terzo larve instar Drosophila Al fine di ottenere ottimali cromosomi politenici per le preparazioni di alta qualità da squash, condizioni di coltura poco affollate sono essenziali (cioè, posto a circa 20 la deposizione delle uova le mosche femmina in uno standard 4 &quot;bottiglia di volare e passare a una nuova bottiglia ogni giorno). Selezionare la più grassa individui dal primo raccolto di arrampicata 3 larve instar mentre sono ancora vaganti, ma appena prima di impupamento. rotazione abbiamo cultura a 21 ° C, ma 18 ° C produrrà più grassi cromosomi che possono essere più adatti per alcuni scopi come ad esempio quando la banda / regioni interbanda devono essere visualizzate ad alta risoluzione. 2. Materiali da squash polytene <ul><li> Pinza dissezione Drummond (2) </li><li> Petri (60 x 15 mm) </li><li> Vetrini da microscopio Frosted (Fisher n ° 12-544-3) (poli-lisina rivestita) </li><li> 22 x 22 mm n. 15 coprioggetto (Fisher n ° 12-520B) (rivestito con Sigmacote; Sigma # SL2) </li><li> 22 x 40 mm n. 15 coprioggetto (Fisher n ° 12-530B) </li><li> Kim-salviette </li><li> Microscopio a contrasto di fase con obiettivo 20X </li><li> Piccolo Dewar (ad esempio, borraccia o thermos vuoto bottiglia) </li><li> Pinze lunghe </li><li> Lamette da barba </li><li> Coplin Jar </li><li> Gomma-Colf o vassoio Tupperware (o equivalente cassetto richiudibile) </li><li> Parafilm (tagliato in 22 piazze mm) </li><li> 175 g di peso </li></ul> 3. Fissativi e soluzioni <ol><li> Magazzino formaldeide 5X. Preparare una soluzione fresca di 0,74 g. paraformaldeide in 4,0 ml di dH 2 O e con 28 ml di KOH 1N. Calda a 65 ° C per sciogliere la paraformaldeide e quindi memorizzare su ghiaccio. </li><li> Fissativo 1. Preparare una soluzione fresca di 0,5 ml di PBS 10X, 50μl Triton X-100, 3,45 ml di dH 2 O e 1,0 ml di uno stock di formaldeide 5X. Caldo per disperdere i Triton X-100 e l&#39;utilizzo entro 1 ora. </li><li> Fissativo 2. Preparare una soluzione fresca di 1,5 ml di dH 2 O, 2,5 ml di acido acetico glaciale, e 1,0 ml da uno stock di formaldeide 5X e utilizzare entro 1 ora. </li><li> Soluzione di acido Lactoacetic. Preparare una soluzione di 1 ml di acido lattico, 2 ml di dH 2 O, e 3 ml di acido acetico. </li></ol> 4. Polytene cromosoma preparazione di squash: <ol><li> Risciacquare con acqua larve e trasferimento in PBS in un piatto di coltura per la dissezione. </li><li> Afferrare la punta della bocca ganci con un paio di pinze, tenere il corpo circa 2 / 3 del modo verso il basso con l&#39;altra coppia, e tirare sulla bocca ganci così le ghiandole salivari sono esposti. Separare le ghiandole salivari dal cervello e occhi antennali dischi, e sezionare via il corpo di grasso e qualsiasi altri tessuti associati dalle ghiandole. </li><li> Aggiungere 200 microlitri di Fissativo 1 al primo pozzo di una diapositiva due depressione bene e 200-300 microlitri di Fissativo da 2 a ben due. Trasferimento di una coppia di ghiandole salivari in un momento di Fissativo 1 nel bene uno e incubare per la quantità di tempo necessario per l&#39;epitopo bersaglio, di solito circa 1-2 minuti. (Nota: alcuni epitopi, come per la maggior parte delle modificazioni degli istoni, può richiedere 5 minuti o più a lungo fissazione.) </li><li> Pinze utilizzando il trasferimento delle ghiandole salivari di Fissativo 2 in ben 2 e incubare per due minuti. </li><li> Trasferire le ghiandole a 10-30 ml di soluzione acida Lactoacetic su un vetrino pulito Sigmacoted. Abbassare delicatamente uno polilisina rivestita vetrino da microscopio sul vetrino e, senza applicare pressione verticale, sollevare il vetrino così le ghiandole sono tra il vetrino ed il coprioggetto. Immediatamente facilitare la lisi cellulare e cromosomiche diffondere con attenzione afferrando il vetrino con una pinza su un lato delicatamente e spostando leggermente avanti e indietro, cercando di minimizzare qualsiasi pressione verticale sul tessuto. In alternativa, utilizzare il lato gomma di una matita o anche un dito per spostare delicatamente il coprioggetto avanti e indietro. Si noti che qualsiasi ritardo nello spostamento del coprioggetto avanti e indietro è probabile che a diminuire la diffusione delle braccia cromosomiche come tenderanno a diventare più rigida su esposizione alla soluzione di acido Lactoacetic. Intorbidimento della soluzione è spesso una buona indicazione di separazione delle cellule. Picchiettando leggermente sulla coprioggetto obliquamente (per evitare una pressione verticale) con il lato gomma di una matita un paio di volte può anche aiutare nella diffusione dei cromosomi. </li><li> Immediatamente esaminare il tessuto al microscopio a contrasto di fase con un obiettivo 20 o 40X per determinare se i cromosomi sono ben diffuse. </li><li> Quando il cromosomico diffusione è sufficiente impostare la diapositiva con il lato coprioggetti in giù su una pila di pulizia Kim-salviette. Posto un secondo Kim-wipe sulla parte superiore e appiattire cromosomi, ponendo il pollice su dove è posizionato il coprioggetto e premendo con decisione, evitando qualsiasi movimento orizzontale del vetrino che al taglioi cromosomi. </li><li> Esaminare il vetrino di nuovo sotto il microscopio per determinare se il preparato è adatto per lo scopo previsto. Se i cromosomi sono mossi in maniera significativa su schiacciamento, l&#39;uso meno volume di soluzione acida Lactoacetic nella vostra preparazione successive. Ripetere i passaggi 4,1-4,8 fino a quando un numero sufficiente di diapositive adatte sono stati ottenuti. Mettere 175 g di peso sul vetrino delle diapositive. </li><li> Riempire un piccolo Dewar (ad esempio, thermos) con azoto liquido. Utilizzando una pinza lunga, tuffo scivolare in N 2 liquido fino a quando la smette di ebollizione, togliere diapositiva e utilizzare immediatamente il bordo di una lama di rasoio pulito in un angolo a buttar via il coprioggetto. Se diapositive sarà utilizzato immediatamente, mettere in una vaschetta Coplin con PBS a 4 ° C. Mettere insieme il vetrino in una vaschetta Coplin pieno di etanolo al 95%. Esaminare il vetrino una volta che il gelo ha sublimato via per confermare che il tessuto aderito alla diapositiva e non rimanere con il coprioggetto. Ripetete con il resto delle diapositive fino a quando tutte le diapositive vengono memorizzati in PBS (per l&#39;uso entro alcune ore) o etanolo (per la conservazione a lungo termine). </li></ol> Rappresentante Risultati Il primo passo fondamentale per l&#39;ottenimento di alta qualità polytene preparazione zucca è di far crescere le larve di grasso con grandi nuclei delle ghiandole salivari. Il secondo è una buona tecnica con la procedura di diffusione, che può richiedere una certa pratica. Un consiglio per una migliore diffusione successo è quello di trovare il volume minimo di soluzione di acido lactoacetic durante la fase di schiacciamento. Ciò promuoverà sufficiente diffusione dei cromosomi senza generare eccessive forze di streaming in grado di lavare via le armi dei cromosomi. E &#39;anche opportuno ricordare che i ritardi nello spostamento del coprioggetto avanti e indietro riduce la diffusione delle braccia cromosomiche man mano che diventano più rigidi quando sono esposti alla soluzione acida Lactoacetic. Se tutto va bene, ci dovrebbe essere numerosi cromosomi ben distribuiti politenici. La Figura 1 mostra un esempio di una tale preparazione, doppie etichettato con un marcatore per le regioni interbanda in verde e con un colorante che macchia le regioni a bande di colore blu. Se insufficiente diffusione è ottenuto, i cromosomi sarà simile a palline, come mostrato nella Figura 2. D&#39;altra parte, se troppo diffusione si è verificata, il chomosomes sarà troppo sottile e prolungato o in alcuni casi frammentato in piccoli pezzi, come mostrato nella Figura 3. <img src="/files/ftp_upload/1748/1748fig1.jpg" alt="Figura 1" /> Figura 1. Preparazione zucca polytene doppio etichettati con anticorpi al JIL-1 istone H3S10 chinasi (in rosso) e Hoechst (blu). <img src="/files/ftp_upload/1748/1748fig2.jpg" alt="Figura 2" /> Figura 2. Zucca politene con diffusione insufficiente. <img src="/files/ftp_upload/1748/1748fig3.jpg" alt="Figura 3" /> Figura 3. Zucca politene con troppa diffusione.

<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;inclusione degli acidi acetico e lattico in protocolli convenzionali fissazione da squash facilita sia la risoluzione interbanda e braccio cromosomico diffusione ma purtroppo alcuni epitopi non sopravvivono a questo trattamento. Un esempio di tale epitopo è H3S10ph (Cai et al., 2008). Poiché il trattamento acido ha anche lo svantaggio che disseta la fluorescenza intrinseca di proteine ​​GFP-tagged, DiMario et al. (2006) ha recentemente sviluppato una base di formaldeide &quot;acid-free tecn...

preparation of drosophila polytene chromosome squashes for antibody labeling

Drosophila è stata a lungo un sistema di preferiti modello per studiare la relazione tra struttura della cromatina e regolazione dei geni per i vantaggi citologico fornite dal cromosomi giganti ghiandola salivare polytene larve del terzo stadio. In questo tessuto i cromosomi subiscono molti cicli di replicazione in assenza di divisione cellulare dando luogo a circa 1000 copie. Il DNA rimane allineata dopo ogni ciclo replicativo conseguente cromosomi notevolmente allargata che forniscono una opportunità unica per correlare morfologia della cromatina con la localizzazione di specifiche proteine. Di conseguenza, c&#39;è stato un elevato livello di interesse nella definizione delle modificazioni epigenetiche presenti in geni diversi e in diverse fasi del processo di trascrizione. Un importante strumento per tali studi è l&#39;etichettatura dei cromosomi politenici con anticorpi per l&#39;enzima, fattore di trascrizione o la modifica degli istoni di interesse. Questo protocollo video illustra la tecnica di zucca utilizzata in laboratorio per preparare Johansen Drosophila cromosomi politenici per l&#39;etichettatura degli anticorpi.

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

Questo protocollo video illustra la tecnica di zucca utilizzata in laboratorio per preparare Johansen<em&gt; Drosophila</em&gt; Cromosomi politenici per l&#39;etichettatura degli anticorpi.

Preparazione dei cromosomi di Drosophila Zucche politene per l&#39;etichettatura di anticorpi

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

Research

JoVE Journal

Biology

31.0K Views.  Iowa State University. Questo protocollo video illustra la tecnica di zucca utilizzata in laboratorio per preparare Johansen Drosophila Cromosomi politenici per l'etichettatura degli anticorpi.

Video: 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.

Preparazione delle Culture neuronale da Midgastrula fase embrioni di Drosophila

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

Ricerca

Biologia

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 preparazione di Drosophila Gli embrioni per Live-Imaging Utilizzando il protocollo goccia

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.

Etichette fluorescenti di Drosophila Cuore Strutture

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.

Preparazione di embrioni per la microscopia elettronica del Drosophila Tubo cuore embrionale

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.

Dissezione di vivere Drosophila Embrioni: Metodi semplificati per lo screening Collezioni Mutant dalla colorazione anticorpale

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.

Preparazione di Sviluppo e degli adulti Drosophila Brains e retina per l&#39;imaging dal vivo

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.

Preparazione del Drosophila Cellule S2 per la microscopia ottica

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.

Preparazione di adulti Drosophila Occhi di sezionamento sottile e analisi microscopica

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.

Studiare Checkpoint mitotica illustrando un comportamento dinamico delle proteine ​​cinetocoro e Motion cromosoma in Living<em&gt; Drosophila</em&gt; Embrioni sinciziale

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).

Cromosoma Preparazione da cellule in coltura

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