KB est soutenu par NIH GM54544.

Partie 1: matériel et fournitures <ol><li> Un microscope de dissection est nécessaire pour une bonne dissections embryon; grossissement 40X est suggéré, avec oculaires 25X au maximum à augmenter le grossissement. </li><li> Pinces fines (numéro 5) sont requis pour la sélection manuelle des embryons et des devitellinization d&#39;embryons. </li><li> Équipement pour faire et modifier des aiguilles de verre fine est nécessaire. Tiré de verre aiguilles sont nécessaires pour la dissection. Nous préférons le verre solide pour la dissection (durent plus longtemps), mais la norme de tubes de verre à parois épaisses (diamètre extérieur 1-1,5 mm) fonctionne aussi bien. Certaines personnes utilisent des aiguilles de tungstène fine, électrolysée à la netteté voulue dans un bain de NaOH 1M en utilisant une batterie 10 Amp + autotransformateur. Aiguisé aiguilles de dissection de tungstène ont l&#39;avantage qu&#39;ils sont relativement résistants et durables. Aiguilles de verre creux (formé similaires à patcher électrodes pince) sont attachés aux tuyaux de plastique et utilisées lors de la dissection pour l&#39;aspiration et l&#39;expulsion des salines, ainsi que la livraison contrôlée de colle. Une variété d&#39;extracteurs de verre sont disponibles pour la fabrication d&#39;aiguilles de verre. Ordinateur programmé extracteurs peut être préréglé pour tirer une variété de formes et tailles. Le Brun-Flaming modèles (Sutter Instrument Co.) sont largement favorisées. Un microscope composé (20-40X objectif) devrait être disponible pour inspecter et modifier des électrodes avant d&#39;utiliser. </li><li> Deux types de solutions sont couramment utilisées pour le bain du tissus embryonnaires lors de la dissection: 1) «standard» (Jan et Jan, 1976a) ou &quot;standard modifié&quot; (Broadie, 2000) Salines, basée sur des solutions couramment utilisés pour l&#39;enregistrement dans d&#39;autres systèmes d&#39;invertébrés , et 2) &quot;hémolymphe-like» (HL) Salines (Stewart et al 1994), un compromis entre la solution saline standard et les concentrations ioniques mesurés dans l&#39;hémolymphe de drosophile. Il est à noter qu&#39;aucun de ces salines précisément imiter la composition chimique de l&#39;hémolymphe in vivo (Broadie, 2000). Salines standard contient (en mM): 135 NaCl, KCl 5, 4 MgCl 2, 1,8 CaCl2, 72 sucrose, 5 TES, pH 7,2. Les expérimentateurs peuvent également envisager une solution saline insectes préparés commercialement (insectes par exemple médias Schneider), qui est impropre à l&#39;enregistrement électrophysiologique, mais les plus susceptibles de protéger la santé des tissus exposés. </li></ol> Partie 2: Mise en scène de l&#39;embryon et dissection <ol><li> Pour la collecte d&#39;œufs optimale, de maintenir les jeunes (&lt;7 jours), de sexe masculin en bonne santé et mouches femelles (20-40) sur les frais des pots de pose de 2-3 jours. Pose des pots se composent généralement de 100 gobelets en plastique ml (Tri-Verser) perforé pour assurer une circulation d&#39;air suffisante, couvrant une plaque de gélose à 60 mm. Plaques de gélose contenant de pomme ou de jus de raisin trempé avec de l&#39;agar. Plusieurs recettes existent, mais un exemple est la suivante: 700 ml d&#39;eau, 25-30 grammes d&#39;agar, 300 ml de concentré de jus (de raisin ou de pomme), 0.5g méthyl paraben (p-hydroxymethylbenzoate), et le sucre 30g. Autoclave l&#39;eau et l&#39;agar, et séparément bouillir le p-hydroxymethylbenzoate avec le sucre et le jus. Mélanger l&#39;agar-agar et des solutions de jus et verser rapidement en 60 plaques mm, éliminer les bulles. </li><li> Avant la collecte d&#39;œufs, les mouches sont nourris avec de la pâte de levure (7 grammes de levure de boulanger dans 9 ml d&#39;eau stockée à 4 ° C) sur des plaques fraîches, changé au moins deux fois par jour pendant au moins deux jours. Au jour 3, un pot de bonne devrait produire 100 à 200 œufs par heure. Mieux la pose d&#39;oeuf est observé sur un pot maintenu sur le côté avec l&#39;agar-agar dans la plaque rayée. Beaucoup embryons seront fixées dans ou à côté de la rayure. </li><li> Pour collecter un grand nombre d&#39;embryons, le transfert des embryons en douceur de la plaque de gélose dans un panier à l&#39;aide d&#39;un pinceau. Un panier peut être faite avec un tube de 15 ou 50 ml centrifugeuse. Coupez le bas en laissant une surface suffisante pour piéger un écran (70 mailles um) entre le couvercle et le haut du tube. Rincez les embryons avec dH 2 0 et mis dans une boîte de Pétri des frais de 50% à 100% eau de javel pour enlever le chorion externe. Alternativement, les œufs peuvent être collectés soit individuellement à l&#39;aide de pinces fines, et dechorionated en les plaçant manuellement dans une goutte d&#39;eau de Javel. De-chorionation peut prendre de 30 secondes à 2 minutes (eau de Javel fraîche est beaucoup plus rapide), et doivent être surveillés sous le microscope de dissection. Retrait du chorion expose le brillant, la membrane transparente vitelline. Dechorionated oeufs doivent être brièvement rincés, ou placés dans une solution saline, l&#39;eau de Javel détruit rapidement les tissus devitellinized. </li><li> Il est essentiel de bien l&#39;étape embryons à l&#39;âge désiré de développement. La drosophile défini embryonnaire (1-17; Campos-Ortega et Hartenstein, 1985) ne sont pas utiles, comme tout développement fonctionnelle neuromusculaire survient dans 16 stade tardif ou le stade 17. Ainsi, les embryons sont mis en scène par heure de développement à 25 ° C dans un incubateur humidifié, en quelques heures après la fécondation, ou, plus fréquemment, après la ponte (AEL). Dans ces conditions, l&#39;embryogenèse dure 21 + / - 1 h à 25 ° C. </li><li> Oeufs à partir d&#39;un oeuf chronométrés-lay (1-2 hrs) qui sont d&#39;âge approprié sont considérées pour les prochains critères morphologiques. Après un lavage extensif en DH 2 O, les oeufs dechorionated sont placés dans une boîte de culture en plastique (sous dH 2 O) pour la visualisation. Mise en scène correcte peut être confirmé par des critères morphologiques en utilisant la lumière réfléchie avec un microscope à dissection (Campos-Ortega et Hartenstein, 1985). Mature embryons à un stade avancé (22-24h AEL) sur le point d&#39;incubation sont généralement reconnus par la segmentation de la cuticule et de la trachée gonflée. Les embryons peuvent également être génotypés avec des équilibreurs GFP et un microscope à dissection de fluorescence avec des filtres appropriés. </li><li> Pour la dissection, les embryons et de développement mis en scène dechorionated sont attachés (face dorsale vers le haut) sur une lamelle en une goutte de sérum physiologique (voir étape 1.5). Les embryons sont retirés de la membrane vitelline par ponction de la membrane vers la fin antérieure ou postérieure avec une micropipette de verre ou d&#39;une aiguille de tungstène, et puis doucement en libérant l&#39;embryon de la membrane. En préparation de la dissection, l&#39;embryon doit être placé sur la lamelle avec la surface dorsale jusqu&#39;à (côté dorsal jusqu&#39;à la dissection va exposer le système nerveux et neuromusculaire ventrale pour les expériences. Toutefois, l&#39;embryon ne peut évidemment être placé dans d&#39;autres façons de faciliter l&#39;accès à d&#39;autres les tissus.) </li><li> Embryons au stade précoce (&lt;16 hrs AEL) attacher directement à nettoyer les vitres ou verre recouverte de polylysine (poly-L-lysine bromhydrate; Broadie et Bate, 1993a, b). Âgés embryons (&gt; 16 heures AEL), suite à la formation de la cuticule, doivent être collés, idéalement à Sylgard-enduit (Dow Corning) lamelles. Une eau-polymériser la colle cyanoacrylate chirurgicaux ou vétérinaires est utilisé (Broadie et Bate, 1993c, d). La colle est livrée par le biais d&#39;une pipette d&#39;électrode petit verre (10-20 micromètres de diamètre pointe intérieure) attaché à un tube en caoutchouc avec le flux de la colle contrôlée par la pression bouche. Des précautions doivent être prises lors de la livraison de la colle, la colle polymérise en plus rapidement au contact de la saline. Notez également que le collage ne peut pas être effectuée dans une solution saline sans ou avec de faibles concentrations d&#39;ions divalents, comme la colle nécessite des ions divalents à polymériser. De petites quantités de colle sont d&#39;abord utilisées pour fixer fermement la tête et la queue. </li><li> Une fois les extrémités de l&#39;embryon sont collés à la lamelle, une incision est faite le long de la ligne médiane dorsale avec une électrode en tungstène de verre ou d&#39;une aiguille aiguisée. Ceci est rendu plus facile par la douceur de perforation de la ligne d&#39;incision avant d&#39;effectuer le montage final. Les côtés de l&#39;incision sont ensuite attachés à la lamelle avec plus de colle, pour doucement répandre la plate embryon. Les organes internes, y compris l&#39;intestin, des corps gras et, éventuellement, la trachée sont ensuite éliminés par aspiration avec une pipette de verre attaché au tuyau en caoutchouc (Broadie et Bate 1993a-c). L&#39;embryon doit maintenant être attachés à plat pour la lamelle, l&#39;épiderme vers le bas avec le système nerveux central ventrale, les nerfs périphériques, et la musculature somatique exposés pour l&#39;expérimentation. </li></ol>

Staging précis des embryons est essentielle en raison de la maturation rapide des propriétés fonctionnelles au cours du temps de seulement quelques heures. Plusieurs questions compliquent cette mise en scène. Premièrement, la plupart des chercheurs utilisent oeuf chronométré établit au embryons au stade, mais le temps de ponte peut varier énormément d&#39;un animal à des conditions différentes (Broadie et al. 1992). En particulier, les femelles sur un régime alimentaire restreint tendent à retenir les oeuf...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
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harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

Drosophile est un modèle de premier génétique pour l&#39;étude du développement embryonnaire et à la fois fonctionnelles en neurosciences. Traditionnellement, ces champs sont assez isolées les unes des autres, avec des histoires largement indépendantes et les communautés scientifiques. Cependant, l&#39;interface entre ces domaines généralement disparates sont les programmes de développement qui sous-tendent l&#39;acquisition de propriétés fonctionnelles électriques de signalisation et de la différenciation des synapses chimiques fonctionnels pendant les phases finales de la formation des circuits neuronaux. Cette interface est une zone cruciale pour l&#39;enquête. Chez la drosophile, ces phases de développement fonctionnel se produisent pendant une période de &lt;8 heures (à 25 ° C) pendant le dernier tiers de l&#39;embryogenèse. Cette période tardive de développement a été longtemps considérés comme insurmontables à cause enquête pour le dépôt d&#39;un dur, la cuticule épidermique imperméable. Une percée a été l&#39;avance de l&#39;application de l&#39;eau-de polymérisation de la colle chirurgicale qui peut être appliqué localement à la cuticule pour permettre une dissection contrôlée des embryons à un stade avancé. Avec une incision dorsale longitudinale, l&#39;embryon peut être posé à plat, ce qui expose la corde nerveuse ventrale et la musculature paroi du corps d&#39;une enquête expérimentale. Ce système a été largement utilisé pour isoler et caractériser des mutants génétiques qui altèrent la formation des synapses embryonnaire, et révéler ainsi les mécanismes moléculaires qui régissent la spécification et la différenciation des connexions synaptiques fonctionnelles synapse et propriétés de signalisation.

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Cette technique expose les neuromusculaire Drosophila embryonnaire pour l&#39;immunohistochimie ou l&#39;enregistrement électrophysiologiques. Il est utile pour étudier les événements précoces de développement neuromusculaire ou d&#39;effectuer l&#39;électrophysiologie dans des mutants qui ne peuvent pas éclore.

La récolte et la préparation Drosophile Les embryons destinés à l&#39;enregistrement électrophysiologique et autres procédures

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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Biology

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K vues.  University of Illinois. Cette technique expose les neuromusculaire Drosophila embryonnaire pour l'immunohistochimie ou l'enregistrement électrophysiologiques. Il est utile pour étudier les événements précoces de développement neuromusculaire ou d'effectuer l'électrophysiologie dans des mutants qui ne peuvent pas éclore.

Vidéo: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Harvesting Sperm and Artificial Insemination of Mice

Rodents of the genus Peromyscus (deer mice) are the most prevalent native North American mammals. Peromyscus species are used in a wide range of research including toxicology, epidemiology, ecology, behavioral, and genetic studies. Here they provide a useful model for demonstrations of artificial insemination. 

Methods similar to those displayed here have previously been used in several deer mouse studies, yet no detailed protocol has been published. Here we demonstrate the basic method of artificial insemination. This method entails extracting the testes from the rodent, then isolating the sperm from the epididymis and vas deferens. The mature sperm, now in a milk mixture, are placed in the female&#x2019;s reproductive tract at the time of ovulation. Fertilization is counted as day 0 for timing of embryo development. Embryos can then be retrieved at the desired time-point and manipulated.

Artificial insemination can be used in a variety of rodent species where exact embryo timing is crucial or hard to obtain. This technique is vital for species or strains (including most Peromyscus) which may not mate immediately and/or where mating is hard to assess. In addition, artificial insemination provides exact timing for embryo development either in mapping developmental progress and/or transgenic work. Reduced numbers of animals can be used since fertilization is guaranteed. This method has been vital to furthering the Peromyscus system, and will hopefully benefit others as well.

This protocol demonstrates methods for extracting sperm from the testes of males and then inseminating female mice. This procedure is useful when precise time is needed in developmental studies as well as transgenic work.

Le sperme de récolte et l&#39;insémination artificielle des souris

Rodents of the genus Peromyscus (deer mice) are the most prevalent native North American mammals.  Peromyscus species are used in a wide range of research ...

Recherche

Biologie

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 ...

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

Méthodes électrophysiologiques pour l&#39;enregistrement du potentiel synaptique de la JNM des larves de drosophile

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

Enregistrement électrophysiologique dans le Drosophile Embryon

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 ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

In vivo d&#39;embryons de drosophile Centrifugation

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Imagerie verticale de<em&gt; Drosophila</em&gt; Embryons

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

Stratégie de transfection optimisés pour l&#39;expression et l&#39;enregistrement électrophysiologique des recombinants canaux ioniques voltage-dépendants présents au HEK-293T

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Isolement et purification de la kinésine de Drosophila Embryons

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

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

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