Agradecemos a la Sra. V. Lephart para el mantenimiento de poblaciones de moscas Este trabajo fue financiado por los Institutos Nacionales de Salud de subvención (GM62916) y la National Science Foundation (MCB0817107).

El siguiente protocolo para la preparación de la calabaza politénicos cromosoma es una adaptación del procedimiento descrito en Johansen et al. (2009). 1. Cultura de tercer estadio las larvas de Drosophila Con el fin de obtener óptimos cromosomas politénicos de alta calidad de las preparaciones squash, las condiciones de cultivo con poca gente son esenciales (es decir, el lugar alrededor de 20 huevos moscas hembra en un estándar de 4 botellas vuelan &quot;y el cambio de una botella nueva cada día). Elija el más gordo de los individuos de la primera cosecha de la escalada tercera larvas mientras que todavía están vagando, pero justo antes de la pupación. De manera rutinaria la cultura a 21 ° C, pero 18 ° C se producen mayores cromosomas que pueden ser más adecuados para ciertos fines, como por ejemplo, cuando la banda / regiones interbandas necesidad de ser visualizadas en alta resolución. 2. Materiales politénicos de squash <ul><li> Drummond pinzas de disección (2) </li><li> Placas de Petri (60 x 15 mm) </li><li> Frosted portaobjetos de microscopio (Fisher N º 12-544-3) (poli-lisina) </li><li> 22 x 22 mm N º 15 cubreobjetos (Fisher No. 12-520B) (recubierto con Sigmacote; Sigma # SL2) </li><li> 22 x 40 mm N º 15 cubreobjetos (Fisher No. 12-530B) </li><li> Kim-toallitas </li><li> Microscopio de contraste de fase con el objetivo de 20X </li><li> Dewar pequeña (por ejemplo, termo o botella termo) </li><li> Unas pinzas largas </li><li> Hojas de afeitar </li><li> Coplin Jar </li><li> De goma-Maid o en la bandeja de Tupperware (o la bandeja de cierre hermético equivalente) </li><li> Parafilm (cortadas en cuadrados de 22 mm) </li><li> 175 g de peso </li></ul> 3. Fijadores y soluciones <ol><li> Acciones de formaldehído 5X. Prepare una solución fresca de 0,74 g. paraformaldehído en 4,0 ml de dH 2 O y con 28 l de KOH 1N. Calentar a 65 ° C para disolver el paraformaldehído y luego almacenar en hielo. </li><li> Fijador 1. Prepare una solución fresca de 0,5 ml de PBS 10X, 50μl Triton X-100, 3,45 ml dH 2 O y 1,0 ml de una reserva de formaldehído 5X. Caliente para dispersar a la Triton X-100 y su uso dentro de una hora. </li><li> Fijador 2. Prepare una solución fresca de 1,5 ml de dH 2 O, 2,5 ml de ácido acético glacial, y 1,0 ml de una reserva de formaldehído 5X y su uso dentro de una hora. </li><li> Solución de ácido Lactoacetic. Prepare una solución de ácido láctico de 1 ml, 2 ml de dH 2 O, y 3 ml de ácido acético. </li></ol> 4. Cromosomas politénicos de squash preparación: <ol><li> Enjuague con agua las larvas y la transferencia de PBS en una placa de cultivo de tejidos para la disección. </li><li> Agarre la punta de la boca ganchos con un par de pinzas, sujete el cuerpo aproximadamente 2 / 3 del camino hacia abajo con la otra pareja, y tirar de la boca ganchos para las glándulas salivales están expuestos. Separar las glándulas salivales del cerebro y los ojos-antenal discos, y diseccionar la grasa corporal y los tejidos asociados a otros de las glándulas. </li><li> Añadir 200 microlitros de un fijador para el primer pozo de una diapositiva de la depresión y dos y 200 a 300 microlitros de fijador y 2 a dos. La transferencia de un par de glándulas salivales en un momento en que un fijador en un bien e incubar la cantidad de tiempo requerido para el epítopo objetivo, por lo general unos 1-2 minutos. (Nota: algunos epítopos, como la mayoría de las modificaciones de las histonas, puede requerir 5 minutos o más la fijación.) </li><li> Pinzas mediante la transferencia de las glándulas salivales de fijador y 2 en 2 y se incuba durante dos minutos. </li><li> Transferencia de las glándulas de 10-30 l de solución de ácido Lactoacetic en un cubreobjetos Sigmacoted limpio. Baje suavemente el portaobjetos del microscopio recubiertos con polilisina sobre el cubreobjetos y, sin aplicar presión vertical, levante el cubreobjetos para las glándulas se encuentran entre el portaobjetos y un cubreobjetos el. Inmediatamente facilitar la lisis celular y la difusión de los cromosomas con cuidado agarrando el cubreobjetos con unas pinzas en un borde y suavemente mueve un poco hacia atrás y hacia delante, tratando de minimizar cualquier presión vertical sobre el tejido. También puedes utilizar el lado de goma de un lápiz o un dedo para mover suavemente el cubreobjetos de ida y vuelta. Tenga en cuenta que cualquier retraso en el movimiento de la cubreobjetos de ida y vuelta es probable que disminuya la difusión de los brazos cromosómicos ya que tienden a ser más rígidos con la exposición a la solución de ácido Lactoacetic. Enturbiamiento de la solución es a menudo una buena indicación de la separación de células. Golpeando suavemente el cubreobjetos oblicua (para evitar la presión vertical) con la cara de goma de un lápiz un par de veces también puede ayudar en la difusión de los cromosomas. </li><li> Inmediatamente examinar el tejido bajo un microscopio de contraste de fase con un objetivo de 20 o 40 veces para determinar si los cromosomas están bien extendidas. </li><li> Cuando la difusión de los cromosomas es suficiente, establezca la diapositiva con la parte cubreobjetos abajo en una pila de agua limpia Kim-toallitas. Coloque un segundo Kim-borrar en la parte superior y aplanar los cromosomas, colocando el pulgar sobre el lugar donde se coloca el cubreobjetos y presionar con firmeza, evitando cualquier movimiento horizontal del cubreobjetos que se cortelos cromosomas. </li><li> Examinar el portaobjetos de nuevo bajo el microscopio para determinar si la preparación es adecuada para el propósito previsto. Si los cromosomas se han movido de manera significativa a aplastar, usar menos volumen de solución ácida Lactoacetic en su preparación posterior. Repita los pasos 4.1-4.8 hasta un número suficiente de diapositivas adecuadas se han obtenido. Lugar 175 pesos g en el cubreobjetos de las diapositivas. </li><li> Llenar un pequeño Dewar (por ejemplo, termo) con nitrógeno líquido. Con una pinza larga, baño caer en N 2 líquido hasta que deje de hervir, retire de diapositivas, e inmediatamente utilizar el borde de una hoja de afeitar limpia en una esquina para voltear de la cubreobjetos. Si de diapositivas se puede utilizar de inmediato, colocar en un frasco Coplin con PBS a 4 ° C. Cualquier forma de recolectar la diapositiva en una jarra Coplin llena de etanol al 95%. Examinar el portaobjetos una vez que el hielo se ha sublimado lejos de confirmar que el tejido adherido a la diapositiva y no quedarse con el cubreobjetos. Repita con el resto de las diapositivas hasta que todas las diapositivas se almacenan en PBS (para su uso dentro de varias horas) o etanol (para un almacenamiento más largo plazo). </li></ol> Resultados representante El primer paso crítico para la obtención de politénicos de alta calidad de la preparación de squash es hacer crecer las larvas de grasa con núcleos grandes glándulas salivales. El segundo es una buena técnica con el procedimiento de difusión, que puede tomar un poco de práctica. Un consejo para el éxito de la mejora se está extendiendo para encontrar el volumen mínimo de solución de ácido lactoacetic durante el paso de aplastamiento. Esto promoverá la suficiente difusión de los cromosomas sin generar excesivas fuerzas de transmisión que puede lavarse los brazos cromosómicos de distancia. Es también digno de mención que cualquier retraso en el movimiento de la cubreobjetos de ida y vuelta se reducirá la difusión de los brazos cromosómicos, ya que se vuelven más rígidos cuando son expuestos a la solución de ácido Lactoacetic. Si todo va bien, debería haber numerosas y difundir los cromosomas politénicos. La figura 1 muestra un ejemplo de este tipo de preparación, el doble etiquetado con un marcador para las regiones interbandas en verde y con un colorante que tiñe las regiones con bandas de color azul. Si es insuficiente la difusión se obtiene, los cromosomas se verá como bolitas, como se muestra en la Figura 2. Por otro lado, si mucho la difusión que ha ocurrido, el chomosomes será demasiado delgada y se extendió en algunos casos, fragmentado en pedazos pequeños, como se muestra en la Figura 3. <img src="/files/ftp_upload/1748/1748fig1.jpg" alt="Figura 1" /> Figura 1. Preparación de calabaza politénicos doble etiquetado con anticuerpos contra el JIL-1 histona H3S10 quinasa (en rojo) y Hoechst (azul). <img src="/files/ftp_upload/1748/1748fig2.jpg" alt="Figura 2" /> Figura 2. Politénicos calabaza con la difusión suficiente. <img src="/files/ftp_upload/1748/1748fig3.jpg" alt="Figura 3" /> Figura 3. Calabaza politénicos con demasiada difusión.

<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>

La inclusión de los ácidos acético y láctico en los protocolos convencionales de squash fijación facilita tanto la resolución interbandas y brazo cromosómico de propagación, pero, lamentablemente, algunos epítopos no sobreviven a este tratamiento. Un ejemplo de este tipo es un epítopo H3S10ph (Cai et al., 2008). Dado que el tratamiento con ácido también tiene la desventaja de que apaga la fluorescencia intrínseca de las buenas prácticas agrarias de etiquetado proteínas, DiMario et al. (2...

preparation of drosophila polytene chromosome squashes for antibody labeling

Drosophila ha sido durante mucho tiempo un sistema modelo preferido para el estudio de la relación entre la estructura de la cromatina y la regulación de genes debido a las ventajas citológico siempre por el gigante de los cromosomas politénicos de las glándulas salivales de las larvas. En este tejido se someten a los cromosomas de muchas rondas de replicación en la ausencia de división celular que da lugar a unas 1000 copias. El ADN permanece alineado después de cada ciclo de replicación de los cromosomas que resulta muy ampliado que proporcionan una oportunidad única para relacionar la morfología de la cromatina con la localización de proteínas específicas. En consecuencia, ha habido un alto nivel de interés en la definición de las modificaciones epigenéticas presentes en genes diferentes y en diferentes etapas del proceso de transcripción. Una herramienta importante para estos estudios es el etiquetado de los cromosomas politénicos con anticuerpos frente a la enzima, el factor de transcripción, o modificación de las histonas de interés. Este protocolo de vídeo muestra la técnica de squash utilizados en el laboratorio para preparar Johansen Drosophila cromosomas politénicos para el etiquetado de anticuerpos.

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

Este protocolo de vídeo muestra la técnica de squash utilizados en el laboratorio para preparar Johansen<em&gt; Drosophila</em&gt; Cromosomas politénicos para el etiquetado de anticuerpos.

Preparación de los cromosomas politénicos de Drosophila Calabazas para el Etiquetado de anticuerpos

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 vistas.  Iowa State University. Este protocolo de vídeo muestra la técnica de squash utilizados en el laboratorio para preparar Johansen Drosophila Cromosomas politénicos para el etiquetado de anticuerpos.

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.

Preparación de cultivos neuronales a partir de embriones de Drosophila Midgastrula Etapa

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

Investigación

Biología

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 preparación de los Drosophila Los embriones para Live-Imaging utilizando el protocolo de gota

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.

Marcaje fluorescente de Drosophila Estructuras del Corazón

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.

Preparación de embriones para Microscopía Electrónica de la Drosophila Tubo cardíaco embrionario

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.

La disección en vivo de Drosophila Los embriones: métodos simplificados para la detección colecciones mutantes tinción con anticuerpos

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.

Preparación de desarrollo y de adultos Drosophila Los cerebros y retinas de imágenes en 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.

Preparación de Drosophila Células S2 para microscopía de luz

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.

Preparación de los adultos Drosophila Ojos de seccionamiento fino y análisis microscópico

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.

El estudio de puesto de control mitótico, ilustrando comportamiento dinámico de proteínas cinetocoro y el movimiento de cromosomas en vivo<em&gt; Drosophila</em&gt; Los embriones Sincitial

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 Preparación de las células cultivadas

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