Ce travail a été soutenu par le NIH aux subventions NS27501 DKOD. Un appui supplémentaire pour ce travail à partir d&#39;une subvention HHMI professeur d&#39;DKOD.

Collection I. embryon de drosophile <ol><li> Préparer les plaques collecte des œufs <ol><li> Faire bouillir 200 ml dH 2 O avec 8 g d&#39;agar </li><li> Laisser refroidir à 50 ° C </li><li> Ajouter 2 ml de EtOH et 2 ml d&#39;acide acétique </li><li> Verser dans des boîtes de 35 mm de Petri en plastique, dessus </li><li> Refroidir et conserver dans contenant hermétique à 4 ° C </li><li> Donne environ 80 plaques </li></ol></li><li> Préparer la pâte de levure <ol><li> Dissoudre la levure active Fleishmans de cuisson dans de l&#39;eau pour former une pâte </li><li> Micro-ondes pendant 20-30 secondes pour tuer la levure (attention aux bouillante sur) </li><li> Stocker dans une seringue de 10 cc (sans aiguille) à 4 ° C pendant 2 semaines </li></ol></li><li> Mouches: Les adultes utilisés pour la collecte d&#39;œufs sont généralement plus productives entre 3 et 14 jours après son apparition, pourvu qu&#39;ils aient eu accès à des aliments frais. De nombreux stocks de mutants ont diminué de fertilité qui peut se manifester à un taux réduit de la ponte et un changement dans l&#39;âge où ils sont plus productifs. Généralement, plusieurs centaines d&#39;adultes font pour une bonne ponte de la population. </li><li> Procédure: Étendre une mince couche de levure pâte sur une plaque de collecte des œufs. Cela fournit un substrat favorable à la ponte des œufs. Transfert adulte vole à une bouteille de collecte des œufs propres et la plaque de collection de bandes à l&#39;embouchure de la bouteille. Ne laissez pas les mouches dans ces bouteilles plus longtemps que 3-4 heures que l&#39;humidité augmente et les mouches se coincer dans la pâte et sur les côtés de la bouteille. </li></ol> II. Dechorionation Embryon avec javellisant <ol><li> Mettre en place un entonnoir Büchner relié à un flacon de filtre attaché à un aspirateur. Placer un morceau de papier Whatman n ° 1 dans l&#39;entonnoir. Avec aspiration course, mouiller le papier avec de l&#39;eau distillée stérile. </li><li> Rincez embryons à partir de plaques de collecte sur le papier filtre avec un jet d&#39;eau à partir d&#39;un flacon de lavage. </li><li> Rincez-les en plusieurs volumes d&#39;eau stérile, éteignez l&#39;aspirateur, puis ajoutez un volume d&#39;entonnoir d&#39;eau de Javel à 50%. Laissez les œufs reposer pendant 5-10 minutes. Le chorions sera dissoute en 1-2 minutes, mais plus vous laissez les embryons dans l&#39;eau de javel plus de la levure contaminant seront détruits. Ramassez tous les adultes ou les larves qui ont rincé plat. </li><li> Rincez les embryons dans une boîte de Pétri stérile (culture de tissus non) avec de l&#39;eau distillée stérile. Beaucoup des embryons sera coller au fond du plat si elle n&#39;a pas été pré-mouillé. Rincez plusieurs fois à l&#39;eau stérile dans ce plat. </li><li> Examiner les embryons avec la lumière transmise à repérer embryons au stade mi-gastrula. </li></ol> III. Préparation des cultures d&#39;embryon unique <ol><li> Faire DDM1 (voir les recettes dans la section IV) </li><li> Versez de l&#39;eau hors plat du embryons juste avant la culture. Couvrir avec les médias embryons stérile. </li><li> Énoncer 3, 35 mm des boîtes de Pétri. Mettre en 4 lamelles rondes autoclavés dans chaque plat. Sur chaque lamelle, mettre 5 pi de milieu. Utilisez Bellco lamelles basse coverlips verre au plomb. Bellco biologique Verrerie 800-257-7043 Cat # 1943-00012 </li><li> Tirez quelques pipettes assez fine. Les pipettes nous utilisons sont tirés sur une Narishige PP-83 extracteur. Les dimensions exactes de la pipette ne sont pas cruciaux et nous avons l&#39;habitude de les tirer plus fine que nous voulons et les casser la pointe dans le même champ de vision comme l&#39;embryon d&#39;évaluer la taille. Vous ne voulez pas d&#39;aspirer tous les embryons d&#39;un seul coup, et ne voulons pas de votre bout si petit qu&#39;il faut 5 minutes pour aspirer les cellules. Visez entre les deux. Examiner les embryons à lumière transmise et d&#39;utiliser un tube d&#39;aspiration la bouche attachée à la pipette pour retirer le contenu de l&#39;embryon. Disperser les cellules sur la lamelle délicatement préparé par l&#39;expulsion des cellules aussi proche de la lamelle que possible. Vous voudrez peut-être examiner les lamelles à haute puissance et encore de disperser les cellules de la même manière. </li><li> Laissez reposer pendant les cellules ne sont plus que 10 minutes. Recouvrir le plat avec les médias et mis en couveuse, 4-5% CO 2 l&#39;environnement, si on utilise un milieu défini. Température entre 22-24 ° C. Nous utilisons régulièrement 23 ° C et une humidité de 95%. L&#39;humidité est importante, comme vous voulez éviter l&#39;évaporation. Vous pouvez utiliser une norme incubateur mammifères placés dans une chambre froide avec contrôle de température réglée à 23 ° C. </li></ol> IV. Moyennes DDM1 Définie pour la croissance des cultures <ol><li> Faire DMEM: Pour 100 ml de Ham F12/DME médias (DMEM) (Irvine Scientific # 9052). <ol><li> Ajouter 0,476 g Hepes (20mm) (Sigma # H-3375). Mélanger. </li><li> Ajouter 1,25 ml de L-glutamine 200 mM (Irvine Scientific # 9317). Mélanger. </li><li> Filtre de hotte avec 0,2 um acétate de filtre seringue (besoin de 2 filtres). </li><li> Le pH est 6,64 et OSM 288. </li><li> Faire des aliquotes de 10 ml dans des tubes de centrifugeuse de 15 ml. </li><li> Conserver à 4 ° C pendant pas plus de 2 semaines. Remarque: Toujours utiliser l&#39;eau filtrée et autoclavés font dans des conteneurs qui sont réservées pour les médias et les suppléments seulement. Ne mettez jamais une électrode de pH ou le barreau d&#39;agitation utilisé à des fins autres Media.To font DDM1: suppléments Ajouter énumérés ci-dessousau DMEM juste avant la culture. </li></ol></li><li> Pour 10 ml de DMEM ajouter: <ul><li> 100 ul de la transferrine </li><li> 100 ul Putrescine </li><li> 100 Sélénium ul </li><li> 100 ul de progestérone </li><li> 50 ul insuline </li></ul></li></ol>

<ol>
	<li>Rohrbough, J., O'Dowd, D. K., Baines, R. A., Broadie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+bases+of+behavioral+plasticity:+Establishing+and+modifying+synaptic+circuits+in+the+Drosophila+Genetic+System.">Cellular bases of behavioral plasticity: Establishing and modifying synaptic circuits in the Drosophila Genetic System.</a> J. Neurobiol. 54, 254-271 (2003).</li><li>O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-gated+currents+and+firing+properties+of+embryonic+Drosophila+neurons+grown+in+a+chemically+defined+medium.">Voltage-gated currents and firing properties of embryonic Drosophila neurons grown in a chemically defined medium.</a> J. Neurobiol. 27, 113-126 (1995).</li><li>Lee, D., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+excitatory+synaptic+transmission+mediated+by+nAChR+in+cultured+Drosophila+neurons.">Fast excitatory synaptic transmission mediated by nAChR in cultured Drosophila neurons.</a> J. Neurosci. 19, 5311-5321 (1999).</li><li>Lee, D., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclic-AMP-dependent+plasticity+at+excitatory+cholinergic+synapses+in+Drosophila+neurons:+Alterations+in+the+memory+mutant+dunce.">Cyclic-AMP-dependent plasticity at excitatory cholinergic synapses in Drosophila neurons: Alterations in the memory mutant dunce.</a> J. Neurosci. 20, 2104-2111 (2000).</li><li>Hodges, D. D., Lee, D., Boswell, K., Preston, C. F., Hall, L. M., O'Dowd, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=tipE+regulates+sodium-dependent+repetitive+firing+in+Drosophila+neurons.">tipE regulates sodium-dependent repetitive firing in Drosophila neurons.</a> Mol. Cell Neurosci. 19, 402-416 (2002).</li><li>Lee, D., Su, H., Dowd, O., K, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GABA+receptors+containing+Rdl+subunits+mediate+fast+inhibitory+synaptic+transmission+in+Drosophila+neurons.">GABA receptors containing Rdl subunits mediate fast inhibitory synaptic transmission in Drosophila neurons.</a> J. Neurosci. 23, 4625-4634 (2003).</li></ol>

Dans les cultures préparées à partir de la drosophile embryons au stade midgastrula les neurones proviennent de précurseurs neuroblastes, dont beaucoup divisent avant de différenciation in vitro. Ce système offre donc une occasion unique d&#39;étudier les facteurs génétiques et environnementaux importants dans des phases très précoces du développement neuronal (Rohrbaugh et al., 2003). Nous avons montré que les neurones cultivés dans un milieu simple défini différencier en neurones électriquement excitables qui a également...

Les cultures faites dans ce média doit être maintenu dans un incubateur à 5% de CO2 à environ 22-24 ° C. Nous utilisons une culture de tissus de mammifères standards incubateur placé dans une chambre froide et chauffée à 23 ° C. Toujours utilisé autoclavé eau filtrée et de faire dans des conteneurs qui sont réservées pour les médias et les suppléments seulement. Ne placez jamais un pH-mètre ou le barreau d&#39;agitation utilisé à d&#39;autres fins dans les médias.

preparation of neuronal cultures from midgastrula stage drosophila embryos

Cette vidéo illustre la procédure de mise cultures neuronales primaires d&#39;embryons de drosophile midgastrula scène. Les méthodes de collecte des embryons et leur dechorionation utilisant l&#39;eau de Javel sont démontrés. Aide d&#39;une pipette en verre attaché à un tube d&#39;aspiration bouche, nous illustrons l&#39;élimination de toutes les cellules issues d&#39;embryons unique. La méthode pour disperser les cellules d&#39;embryons de chaque dans un petit (5 l) goutte de milieu sur une lamelle de verre sans revêtement est démontrée. Une vue à travers le microscope à 1 heure après le placage illustre la densité de cellule préférée. La plupart des cellules qui survivent lorsqu&#39;elles sont cultivées en milieu défini sont neuroblastes qui divisent une ou plusieurs fois dans la culture avant d&#39;étendre les processus neuritique par 12-24 heures. Une vue à travers le microscope illustre le niveau de la croissance des neurites et la ramification attendus dans une culture saine à 2 jours in vitro. Les cultures sont cultivées dans un milieu à base de bicarbonate de simples définis, dans un 5% de CO 2 incubateur à 22-24 ° C. Processus neuritiques continuer à élaborer au cours de la première semaine de la culture et quand ils entrent en contact avec des neurites des cellules voisines, ils forment souvent des connexions synaptiques fonctionnelles. Les neurones de ces cultures d&#39;exprimer voltage-dépendants sodium, le calcium, et les canaux potassiques et sont électriquement excitables. Ce système de culture est utile pour étudier moléculaires des facteurs génétiques et environnementaux qui régulent la différenciation neuronale, excitabilité et la formation des synapses / fonction.

Watch this Scientific Journal Video about Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos at JoVE.com

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

Cette vidéo montre la préparation de cultures primaires de neurones d&#39;embryons de drosophile midgastrula scène. Vues des cultures vivantes montrent les cellules 1 heure après le placage et les neurones différenciés après 2 jours de croissance dans un milieu à base de bicarbonate défini. Les neurones sont électriquement excitables et forment des connexions synaptiques.

Préparation des cultures de neurones d&#39;embryons de drosophile Midgastrula scène

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

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8.7K vues.  University of California, Irvine (UCI). Cette vidéo montre la préparation de cultures primaires de neurones d'embryons de drosophile midgastrula scène. Vues des cultures vivantes montrent les cellules 1 heure après le placage et les neurones différenciés après 2 jours de croissance dans un milieu à base de bicarbonate défini. Les neurones sont électriquement excitables et forment des connexions synaptiques.

Vidéo: Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

Cultures neuronales primaires du cerveau des drosophiles stade tardif pupes

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

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Placing Growth Factor-Coated Beads on Early Stage Chicken Embryos

The neural tube expresses many proteins in specific spatiotemporal patterns during development. These proteins have been shown to be critical for cell fate determination, cell migration, and formation of neural circuits. Neuronal induction and patterning involve bone morphogenetic protein (BMP), sonic hedgehog (SHH), fibroblast growth factor (FGF), among others. In particular, the expression pattern of Fgf8 is in close proximity to regions expressing BMP4 and SHH. This expression pattern is consistent with developmental interactions that facilitate patterning in the telencephalon.

Here we provide a visual demonstration of a method in which an in ovo preparation can be used to test the effects of Fgfs in the formation of the forebrain. Beads are coated with protein and placed in the developing neural tube to provide sustained exposure. Because the procedure uses small, carefully placed beads, it is minimally invasive and allows several beads to be placed within a single neural tube. Moreover, the method allows for continued development so that embryos can be analyzed at a more mature stage to detect changes in anatomy and in neural patterning. This simple but useful protocol allows for real time imaging. It provides a means to make spatially and temporally limited changes to endogenous protein levels.

A variety of growth factors and proteins interact to induce cells to take on different cell fates during development. Here we demonstrate the use of an in ovo preparation to address possible interactions between different proteins in development by placing beads on E2.5 chick embryos.

Placer du facteur de croissance des billes recouvertes sur les premiers embryons de poulet scène

The neural tube expresses many proteins in specific spatiotemporal patterns during development.   These proteins have been shown to be critical for cell ...

In Ovo Electroporations of HH Stage 10 Chicken Embryos

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of molecular biological techniques, the chick has become a useful system in which to study gene regulation and function.  By electroporating DNA or RNA constructs into the developing chicken embryo, genes can be expressed or knocked down in order to analyze in vivo gene function.  Similarly, reporter constructs can be used for fate mapping or to examine putative gene regulatory elements.  Compared to similar experiments in mouse, chick electroporation has the advantages of being quick, easy and inexpensive.  This video demonstrates first how to make a window in the eggshell to manipulate the embryo.  Next, the embryo is visualized with a dilute solution of India ink injected below the embryo.  A glass needle and pipette are used to inject DNA and Fast Green dye into the developing neural tube, then platinum electrodes are placed parallel to the embryo and short electrical pulses are administered with a pulse generator.  Finally, the egg is sealed with tape and placed back into an incubator for further development.  Additionally, the video shows proper egg storage and handling and discusses possible causes of embryo loss following electroporation.

Chick in ovo electroporation is a technique which allows genetic manipulation of the avian embryo. Common applications of this technique include functional analysis of genes and putative enhancer elements. This video demonstrates neural tube electroporation in HH 10 chick embryos. Injection technique and proper egg handling are discussed.

En électroporations Ovo d&#39;embryons au stade de poulet 10 HH

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of ...

Neuronal Cell Cultures from Aplysia for High-Resolution Imaging of Growth Cones

Neuronal growth cones are the highly motile structures at the tip of axons that can detect guidance cues in the environment and transduce this information into directional movement towards the appropriate target cell. To fully understand how guidance information is transmitted from the cell surface to the underlying dynamic cytoskeletal networks, one needs a model system suitable for live cell imaging of protein dynamics at high temporal and spatial resolution. Typical vertebrate growth cones are too small to quantitatively analyze F-actin and microtubule dynamics. Neurons from the sea hare Aplysia californica are 5-10 times larger than vertebrate neurons, can easily be kept at room temperature and are very robust cells for micromanipulation and biophysical measurements. Their growth cones have very defined cytoplasmic regions and a well-described cytoskeletal system. The neuronal cell bodies can be microinjected with a variety of probes for studying growth cone motility and guidance. In the present protocol we demonstrate a procedure for dissection of the abdominal ganglion, culture of bag cell neurons and setting up an imaging chamber for live cell imaging of growth cones.

Aplysia californica neurons develop large growth cones in culture that are excellent for high-resolution imaging of growth cone motility and guidance. Here, we present a protocol for dissection and plating of Aplysia bag cell neurons as well as for setting up a chamber for live cell imaging.

Cultures de cellules neuronales de l&#39;aplysie pour l&#39;imagerie haute résolution des cônes de croissance

Neuronal growth cones are the highly motile structures at the tip of axons that can detect guidance cues in the environment and transduce this information ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

La préparation des Drosophile Les embryons destinés à Live-imagerie utilisant le protocole goutte suspendue

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Preparation of Aplysia Sensory-motor Neuronal Cell Cultures

The nervous system of the marine mollusk Aplysia californica is relatively simple, consisting of approximately 20,000 neurons. The neurons are large (up to 1 mm in diameter) and identifiable, with distinct sizes, shapes, positions and pigmentations, and the cell bodies are externally exposed in five paired ganglia distributed throughout the body of the animal. These properties have allowed investigators to delineate the circuitry underlying specific behaviors in the animal1. The monosynaptic connection between sensory and motor neurons is a central component of the gill-withdrawal reflex in the animal, a simple defensive reflex in which the animal withdraws its gill in response to tactile stimulation of the siphon. This reflex undergoes forms of non-associative and associative learning, including sensitization, habituation and classical conditioning. Of particular benefit to the study of synaptic plasticity, the sensory-motor synapse can be reconstituted in culture, where well-characterized stimuli elicit forms of plasticity that have direct correlates in the behavior of the animal2,3. Specifically, application of serotonin produces a synaptic strengthening that, depending on the application protocol, lasts for minutes (short-term facilitation), hours (intermediate-term facilitation) or days (long-term facilitation). In contrast, application of the peptide transmitter FMRFamide produces a synaptic weakening or depression that, depending on the application protocol, can last from minutes to days (long-term depression). The large size of the neurons allows for repeated sharp electrode recording of synaptic strength over periods of days together with microinjection of expression vectors, siRNAs and other compounds to target specific signaling cascades and molecules and thereby identify the molecular and cell biological steps that underlie the changes in synaptic efficacy.
An additional advantage of the Aplysia culture system comes from the fact that the neurons demonstrate synapse-specificity in culture4,5. Thus, sensory neurons do not form synapses with themselves (autapses) or with other sensory neurons, nor do they form synapses with non-target identified motor neurons in culture. The varicosities, sites of synaptic contact between sensory and motor neurons, are large enough (2-7 microns in diameter) to allow synapse formation (as well as changes in synaptic morphology) with target motor neurons to be studied at the light microscopic level.
In this video, we demonstrate each step of preparing sensory-motor neuron cultures, including anesthetizing adult and juvenile Aplysia, dissecting their ganglia, protease digestion of the ganglia, removal of the connective tissue by microdissection, identification of both sensory and motor neurons and removal of each cell type by microdissection, plating of the motor neuron, addition of the sensory neuron and manipulation of the sensory neurite to form contact with the cultured motor neuron.

Preparation of Aplysia Sensory-motor Neuronal Cell Cultures

Primary cultures of Aplysia sensory-motor neurons provide a model preparation for studying synapse formation and synaptic plasticity in vitro. This video demonstrates the identification and microdissection of sensory and motor neurons from Aplysia ganglia as well as the methods for establishing and maintaining sensory-motor neurons in culture.

Préparation des Aplysie Sensori-motrice cultures de cellules neuronales

The nervous system of the marine mollusk Aplysia californica is relatively simple, consisting of approximately 20,000 neurons.  The neurons are large (up ...

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 ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

Cultures de cellules primaires à partir Drosophile Embryons Gastrula

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...