Nous remercions les membres du laboratoire Gelfand qui ont contribué à l&#39;élaboration de ces protocoles. Recherche présentée dans cette publication a été soutenue par l&#39;Institut national de sciences générales médical des National Institutes of Health sous le numéro d&#39;attribution R01GM052111 de VIG et par basque ministère du gouvernement de l&#39;Éducation, des Universités et de la Recherche sous le numéro d&#39;attribution BFI-2011-295 à UdC.

Une. Cellules S2 de Drosophila <ol><li> Préparation des lamelles couvre-objet enduites ConA <ol><li> Placez les lamelles dans un rack en céramique et les laver par immersion de l&#39;acide chromique pendant 1 heure. [ATTENTION: l&#39;acide chromique est corrosif et peut provoquer une irritation des yeux, du nez, de la gorge et de la peau]. </li><li> Rincer abondamment avec de lamelles constamment courir dH 2 O pendant 30 minutes jusqu&#39;à ce que l&#39;acide est complètement délavé. </li><li> Laissez l&#39;air lamelles sec et les enrober de ConA (solution à 0,5 mg / ml dans dH 2 O) pendant 30 min. </li><li> Rincer lamelles avec dH 2 O pendant 15 minutes et laisser sécher à l&#39;air. Lamelles recouvertes ConA peuvent être stockés jusqu&#39;à 1 mois. </li></ol></li><li> Étalement des cellules <ol><li> Laisser les cellules S2 de croître de façon exponentielle en T25 ou T75 cm2 en fonction du nombre de cellules nécessaires pour l&#39;expérience. </li><li> Compter la densité cellulaire en utilisant un hémocytomètre. Ajouter une croissance ml moyenne (par exemple les insectes Xpress) dans une boîte de 35 mm de tissu de culture avec de la ConA lamelle revêtue, et le transfert en douceur ~ 1 x 10 5 cellules à l&#39;antenne. Cette densité cellulaire est optimale pour l&#39;imagerie parce que les procédés à partir de cellules différentes ne se chevauchent pas. </li><li> Afin d&#39;induire la formation de processus, immédiatement après l&#39;étalement ajouter 1 ml de milieu avec 5 uM CytoD à l&#39;antenne (à la concentration finale de 2,5 uM CytoD). Permettre le plein développement de procédés en incubant les cellules au moins pendant 2 heures à 25 ° C avant l&#39;imagerie. </li></ol></li></ol> 2. Drosophila primaire Neuron Culture <ol><li> Préparation pour la culture de neurones <ol><li> Préparer Drosophila milieu de croissance neuronale (milieu de Schneider supplémenté avec 20% de sérum de fœtus bovin, inactivé par la chaleur à 55 ° C pendant 30 min, 5 ug / ml d&#39;insuline, 100 ug / ml de pénicilline, 100 pg / ml de streptomycine, 10 pg / ml de tetracycline) . Ce soutienle milieu de plemented Schneider peut être conservé à 4 ° C pendant 6 mois. </li><li> Préparer la pomme flacons de gélose au jus de collecte de Drosophila embryons (22,5 g d&#39;agar-agar, 12,0 g de saccharose et 750 ml H 2 O dans un ballon de 2 litres, autoclave pendant 20 à 25 min, ajouter 250 ml de jus de pomme et verser ~ 10 ml dans chaque Drosophila flacon, après solidification, branchez flacons avec des boules de coton, et l&#39;envelopper d&#39;une pellicule de flacons en plastique pour aliments afin d&#39;éviter un séchage excessif de la nourriture). Le jus de pomme dans des flacons de gélose peut être conservé à 4 ° C dans des sacs en plastique bien fermés pendant 4 semaines. </li><li> Lavés à l&#39;acide manteau 25 mm lamelles de cercle avec ConA (voir détails dans le protocole pour les cellules S2 de drosophile) et stériliser les sous lumière UV pendant 10-15 min. Ces lamelles peuvent être stockés pendant un mois. </li></ol></li><li> Culture des neurones embryonnaires de Drosophila <ol><li> Gardez jeune adulte vole dans des flacons de gélose pomme de jus additionné de levure sèche de 1-2 jours avant la collecte des embryons fou la préparation de neurones. </li><li> Le jour de collecte, afin de synchroniser les embryons collectés, transfert mouches adultes à une pomme jus agar flacon pendant 1 heure précollecte à la lumière et jeter ces embryons. </li><li> Transfert vole vers un nouveau flacon de jus de gélose de pomme, de recueillir des embryons pendant 1 heure, et de transférer les mouches à un autre flacon pendant une heure, à la fois dans l&#39;obscurité pour augmenter le rendement de l&#39;embryon. </li><li> Retirez les mouches adultes à partir du jus de pomme agar flacon et laisser les embryons se développent encore 4 h à étapes 9-11 à la température ambiante (4-6 embryons h-vieux sont entre le stade 9-11). </li><li> Retirez les embryons en injectant de l&#39;eau dans des flacons et desserrant les embryons du jus de pomme agar avec un petit pinceau (au moins 10 embryons sont nécessaires pour une bonne préparation des neurones). </li><li> Transférer les embryons à un filet de nylon tamis cellulaire 100 um avec une pipette de transfert pour évacuer l&#39;eau (prélavage la pipette de transfert chez la drosophile tampon de lavage de l&#39;embryon, 0,4% de NaCl, 0,03% de TritonX-100). </li><li> Recueillir les embryons de la crépine en utilisant un petit pinceau, et les transférer dans un microtube de 1,5 ml remplie de 1 ml Drosophila lavage de l&#39;embryon (Note: nettoyer les fibres de gélose ou de coton à l&#39;aide du pinceau ou des embouts de pipette). Dans ce protocole, le microtube de 1,5 ml est livré avec une pastille jetable pilon correspondant. </li><li> Laver les embryons dans l&#39;embryon de drosophile lavage 3x. </li><li> Dechorionate les embryons dans une solution d&#39;eau de Javel frais 01h01 commerciale et 95% d&#39;éthanol pour 10 min [ATTENTION: l&#39;eau de Javel peut provoquer une irritation des yeux, du nez, de la gorge et de la peau]. </li><li> Passez à la hotte de culture de tissus, et laver les embryons 2x en milieu de Schneider complété. </li><li> Laissez 200-300 milieu de pi dans le tube avec les embryons, et doucement dissocier mécaniquement embryons dechorionated utilisant Kimble Chase pilons de granulés jetables qui ont été stérilisées par 70% d&#39;éthanol (10-15x moudre). </li><li> Centrifuger le mélange homogénéisé à 20 g pendant 2 min, et transfer le surnageant dans un tube frais. [Cette étape est facultative, elle aide à éliminer les débris de l&#39;embryon] </li><li> Centrifuger le surnageant à 550 x g pendant 2 min pour sédimenter les cellules. </li><li> Reprendre le culot dans 500 ul de cellules de moyennes et de spin de Schneider complété à nouveau à 550 g pendant 2 min; répéter le cycle de remise en suspension de filage fois. </li><li> Enfin remettre en suspension le culot dans 100 pl ~ supplémenté le milieu de Schneider et de la plaque les cellules sur une lamelle couvre-objet de 25 mm revêtu de ConA dans une boîte de Pétri de 35 mm stériles. Incuber pendant 10 min pour permettre la fixation des cellules. </li><li> Ajouter 2 ml de milieu de Schneider, complétée avec 5 uM CytoD pour submerger la lamelle avec des cellules attachées. </li><li> Incuber le plat dans une humidifié 25 ° C incubateur pendant une nuit avant l&#39;imagerie. </li></ol></li></ol>

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Les auteurs déclarent qu&#39;ils n&#39;ont aucun intérêt financier concurrents.

Nous présentons ici des protocoles détaillés pour étudier le transport de fret microtubules dépendante dans les cellules S2 de drosophile et de la culture de neurones primaires. Les processus et les neurites S2 sont des structures remplies par groupés microtubules tableaux qui servent de pistes pour le transport des organites. Deux stratégies sont généralement utilisées pour des organelles de l&#39;étiquette: la transfection de vecteurs codant pour une protéine fluoresce...

Cellules S2 de drosophile ont gagné en popularité de plus en plus pour l&#39;étude de nombreux processus cellulaires. Ces cellules ont été initialement obtenues à partir d&#39;embryons à un stade avancé de trypsinisées OregonR Drosophila melanogaster et maintiennent des caractéristiques de type macrophage 1. Cellules S2 de Drosophila offrent de nombreux avantages par rapport à d&#39;autres lignées cellulaires. Elles sont mises en culture à 25 ° C sans CO 2, et peuvent être visualisés pendant plusieurs heures à la température ambiante sans nécessiter de chauffage ou d&#39;échange de gaz. Plus important encore, les cellules S2 de Drosophila sont très sensibles à un traitement ARNi permet une grande efficacité de knock-down des protéines d&#39;intérêt. Les cellules S2 sont hautement transfectable et des lignées cellulaires stables peuvent être sélectionnés en moins de quatre semaines pour permettre l&#39;expression ectopique stable de protéines d&#39;intérêt. Les cellules S2 poussent normalement soit vaguement attachés mais pas étalés sur un ballon ou en suspension. Ils peuvent être amenés à se propager sur des lamelles prérevêtues avec une conca lectine végétalenavalin A (ConA). La périphérie des cellules de propagation est particulièrement mince et est idéal pour la microscopie de cellules vivantes donc. Des protocoles détaillés pour les cellules S2 de la culture, le traitement ARNi, l&#39;imagerie par immunomarquage direct et la lumière peuvent être trouvés dans la littérature de 2,3. Notre laboratoire a adopté cellules S2 en tant que système modèle pour étudier le transport du fret à base de microtubules. Cellules S2 de Drosophila traités avec un médicament qui dépolymérise ou rompt la F-actine, tels que la cytochalasine D (CytoD), se développent de longs processus remplis de microtubules uniformément polarisées 4 -10, ce qui en fait un système idéal pour étudier le mouvement des marchandises le long des réseaux de microtubules. Différents paramètres de transport dans ces processus (c.-à-trajectoires. Longueurs, vitesses, directivité etc) peuvent être facilement mesurés en utilisant un logiciel de traces de particules automatisé, DiaTrack (Semasopht: http://www.microscopy.info/Organization/Details/4423; Dr . Pascal Vallotton: http://www.csiro.au/en/Organisation-Structure/Divisions/Mathematics-Informatics-and-Statistics/PascalVallotton.aspx # a4). Ces propriétés rendent les cellules S2 un système attrayant pour étudier le transport de cargaison le long des microtubules dans les cellules de culture de tissu. Des expériences pilotes dans les cellules S2 peuvent être étendus à un modèle physiologique important des études de transport de marchandises dans la drosophile neurones primaires. Similaire à des cellules S2, les neurones mis en culture à partir d&#39;embryons dissociées sont perméables aux petites molécules telles que des colorants et des inhibiteurs, et l&#39;imagerie haute résolution en temps réel du transport des organites peuvent être effectuées dans des neurones à la température ambiante. En utilisant la drosophile fonds génétiques différents, nous pouvons examiner la fonction des gènes dans les neurones primaires par KO (mutations génétiques perte de fonction), knockdown (injection d&#39;ARNdb ou expression) ou l&#39;expression ectopique (mouches transgéniques). Plus précisément, la fragmentation des filaments d&#39;actine en utilisant le traitement CytoD n&#39;empêche pas la croissance des neurites, mais plutôt qu&#39;ils poussent plus vite, et ainsi nous pouvons étudier microtubules dependent le transport des organites dans les neurones CytoD traités sans l&#39;influence de filaments d&#39;actine 11. Par conséquent, les cultures primaires de neurones combinent les avantages des cellules de culture de tissus et de voler la génétique, ce qui en fait un excellent système pour étudier le transport de fret dans un système pertinent physiologique 10,12. Depuis plusieurs maladies neurodégénératives familiales pourraient être causés par ou associées à des défauts de transport de marchandises (par exemple de la maladie de Parkinson 13, la maladie de Huntington 14, la maladie d&#39;Alzheimer 15,16, la sclérose latérale amyotrophique / ALS 17, HMN7B et Perry syndrome 18,19, et Ataxie spinocérébelleuse type 5/SCA5 20), les cultures primaires de neurones peuvent être utiles dans l&#39;analyse des effets de mutations spécifiques qui causent ces maladies sur le transport des microtubules dépendant. Enfin, les propriétés importantes de transport des microtubules dépendant sont bien conservées entre les mammifères et les mouches, mais <em&gt; Culture génétique et cellulaire chez la drosophile est moins cher et prend du temps, ce qui justifie l&#39;utilisation de cellules de drosophile en culture pour l&#39;étude des aspects essentiels de transport des organites dans des conditions normales et pathologiques.

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			<td height="61" style="height:61px;width:124px;">Powder Insect cell medium (Schneider&#8217;s)</td>
			<td style="width:133px;">Sigma&#160;</td>
			<td style="width:179px;">S9895</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Insect &#8211;Xpress medium</td>
			<td style="width:133px;">Fisher</td>
			<td style="width:179px;">BW12730Q</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:124px;">Insulin</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:179px;">I6634</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:124px;">Penicillin</td>
			<td style="width:133px;">Research Products International</td>
			<td style="width:179px;">P93000</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:124px;">Streptomycin</td>
			<td style="width:133px;">Research Products International</td>
			<td style="width:179px;">S62000</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Tetracycline hydrochloride</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:179px;">T7660-5G</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Fetal bovine serum</td>
			<td style="width:133px;">Atlanta Biologicals</td>
			<td style="width:179px;">S11150</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:124px;">Concanavalin A</td>
			<td style="width:133px;">Sigma</td>
			<td style="width:179px;">C2010</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:124px;">Cytochalasin D</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">IC15077105</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">LysoTracker Red DND-99</td>
			<td style="width:133px;">Molecular Probes</td>
			<td style="width:179px;">L7528</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">100 &#181;m Cell strainer</td>
			<td style="width:133px;">Fisher Scientific</td>
			<td style="width:179px;">22363549</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">25 mm Circle cover glass</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">48380 080</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:124px;">Drosophila Vials</td>
			<td style="width:133px;">VWR</td>
			<td style="width:179px;">89092-730</td>
		</tr>
		<tr height="81">
			<td height="81" style="height:81px;width:124px;">Disposable pellet pestles with matching microtubes</td>
			<td style="width:133px;">Kimble Chase</td>
			<td style="width:179px;">749520-0000</td>
		</tr>
	</tbody>
</table>

organelle transport in cultured drosophila cells: s2 cell line and primary neurons.

Des cellules S2 de Drosophila étalées sur une lamelle couvre-objet en présence de tous les longs processus non ramifiées de forme de médicaments de l&#39;actine-dépolymérisation de microtubules uniformément remplis de polarisation. Organites se déplacent le long de ces processus par des moteurs de microtubules. Facile d&#39;entretien, de sensibilité à la protéine ARNi knock-down et procédure efficace pour créer des lignées cellulaires stables rendre les cellules S2 de drosophile un système modèle idéal pour étudier le transport de marchandises par imagerie en temps réel. Les résultats obtenus avec des cellules S2 peuvent être en outre appliqués à un système plus physiologiquement pertinente: transport axonal dans les neurones primaires en culture à partir d&#39;embryons de Drosophila dissociées. Des neurones en culture se développent de longues neurites remplis de microtubules groupés, très similaires aux processus S2. Comme dans les cellules S2, organelles dans des neurones en culture peuvent être visualisées soit par des colorants fluorescents spécifiques à l&#39;organelle ou en utilisant des marqueurs d&#39;organelles fluorescentes codées par l&#39;ADN injecté dans des embryons précoces ou exprimé dans transgenic vole. Par conséquent, le transport des organites peut être facilement enregistré dans les neurones en culture sur des lamelles de verre en utilisant l&#39;imagerie vivant. Nous décrivons ici les procédures pour la culture et la visualisation de transport de marchandises dans les cellules S2 de drosophile et les neurones primaires. Nous croyons que ces protocoles font deux systèmes accessibles pour les laboratoires qui étudient le transport de fret.

Cellules S2 de drosophile attachés sur la propagation de la lamelle ConA revêtu dans un plat rond de forme de &quot;crêpe&quot; (Figures 1A et 1C). Cependant, lorsque les cellules étalées sur des lamelles couvre-S2, en présence d&#39;CytoD et autorisés à se propager pendant 2-3 heures, ils forment de longs processus minces remplis par les microtubules parallèles (figures 1B et 1D). Ces microtubules ont une polarité uniforme avec plus-...

Watch this Scientific Journal Video about Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons. at JoVE.com

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Cellules S2 de drosophile et des neurones en culture sont de grands systèmes pour l&#39;imagerie du transport motorisé organite in vivo. Nous décrivons ici des protocoles détaillés pour la culture des deux types de cellules, leur imagerie et d&#39;analyse de transport.

Organite Transport en culture d&#39;<em&gt; Drosophila</em&gt; Cellules: S2 Cell Line et neurones primaires.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

organelle-transport-cultured-drosophila-cells-s2-cell-line-primary

Research

JoVE Journal

Biology

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

13.6K vues.  Feinberg School of Medicine, Northwestern University.  Cellules S2 de drosophile et des neurones en culture sont de grands systèmes pour l'imagerie du transport motorisé organite in vivo. Nous décrivons ici des protocoles détaillés pour la culture des deux types de cellules, leur imagerie et d'analyse de transport. 

Vidéo: Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

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 ...

Recherche

Biologie

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Imagerie en temps réel du noyau dense de vésicules cultures primaires neurones de l&#39;hippocampe

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Preparation of Drosophila S2 cells for Light Microscopy

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

Preparation of Drosophila S2 cells for Light Microscopy

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

Préparation des Drosophile Les cellules S2 pour la microscopie optique

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

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

Adénovirus suppression génétique des molécules de signalisation dans les cultures de fibroblastes de souris embryonnaires primaires

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

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 ...

Isolation and Primary Culture of Rat Hepatic Cells

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology and other related subjects. The method based on two-step collagenase perfusion for isolation of intact hepatocytes was first introduced by Berry and Friend in 1969 1 and, since then, has undergone many modifications. The most commonly used technique was described by Seglenin 1976 2. Essentially, hepatocytes are dissociated from anesthetized adult rats by a non-recirculating collagenase perfusion through the portal vein. The isolated cells are then filtered through a 100 &mu;m pore size mesh nylon filter, and cultured onto plates. After 4-hour culture, the medium is replaced with serum-containing or serum-free medium, e.g. HepatoZYME-SFM, for additional time to culture. These procedures require surgical and sterile culture steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol, which allow consistently in the generation of viable hepatocytes in large numbers.

Primary hepatocytes provide a valuable tool to evaluate biochemical, molecular, and metabolic functions in a physiologically relevant experimental system. We describe a reliable protocol for rat in situ liver perfusion, which consistently generates viable hepatocytes up to 1.0 &times; 108 cells per preparation with cell viability between 88 ~ 96%.

L&#39;isolement et la culture primaire de cellules de foie de rat

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Imagerie Ca locale<sup&gt; 2+</sup&gt; Signaux de cellules de mammifères cultivées

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Generating a &#34;Humanized&#34; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper distribution of the genetic material requires that sister kinetochores on every chromosome become bioriented by attaching to microtubules from opposite spindle poles before progressing into anaphase. However, erroneous, non-bioriented attachment states are common and cellular pathways exist to both detect and correct such attachments during cell division. The process by which improper kinetochore-microtubule interactions are destabilized is referred to as error correction. To study error correction in living cells, incorrect attachments are purposely generated via chemical inhibition of kinesin-5 motor, which leads to monopolar spindle assembly, and the transition from mal-orientation to biorientation is observed following drug washout. The large number of chromosomes in many model tissue culture cell types poses a challenge in observing individual error correction events. Drosophila S2 cells are better subjects for such studies as they possess as few as 4 pairs of chromosomes. However, small molecule kinesin-5 inhibitors are ineffective against Drosophila kinesin-5 (Klp61F). Here we describe how to build a Drosophila cell line that effectively replaces Klp61F with human kinesin-5, which renders the cells sensitive to pharmacological inhibition of the motor and suitable for use in the cell-based error correction assay.

Generating a &quot;Humanized&quot; Drosophila S2 Cell Line Sensitive to Pharmacological Inhibition of Kinesin-5

This protocol describes how to generate a Drosophila S2 cell line that is sensitive to small molecule inhibitors of kinesin-5. The use of these cells in a cell-based error correction assay is also outlined.

Génération d&#39;un «humanisé»<em&gt; Drosophila</em&gt; S2 Cell Line Sensible à pharmacologique inhibition de Kinesin-5

Kinetochores are large protein-based structures that assemble on centromeres during cell division and link chromosomes to spindle microtubules. Proper ...

Glucose Uptake Measurement and Response to Insulin Stimulation in In Vitro Cultured Human Primary Myotubes

Skeletal muscle is the largest glucose deposit in mammals and largely contributes to glucose homeostasis. Assessment of insulin sensitivity of muscle cells is of major relevance for all studies dedicated to exploring muscle glucose metabolism and characterizing metabolic alterations. In muscle cells, glucose transporter type 4 (GLUT4) proteins translocate to the plasma membrane in response to insulin, thus allowing massive entry of glucose into the cell. The ability of muscle cells to respond to insulin by increasing the rate of glucose uptake is one of the standard readouts to quantify muscle cell sensitivity to insulin. Human primary myotubes are a suitable in vitro model, as the cells maintain many features of the donor phenotype, including insulin sensitivity. This in vitro model is also suitable for the test of any compounds that could impact insulin responsiveness. Measurements of the glucose uptake rate in differentiated myotubes reflect insulin sensitivity.
In this method, human primary muscle cells are cultured in vitro to obtain differentiated myotubes, and glucose uptake rates with and without insulin stimulation are measured. We provide a detailed protocol to quantify passive and active glucose transport rates using radiolabeled [3H] 2-deoxy-D-Glucose ([3H]2dG). Calculation methods are provided to quantify active basal and insulin-stimulated rates, as well as stimulation fold.

Glucose Uptake Measurement and Response to Insulin Stimulation in In Vitro Cultured Human Primary Myotubes

In this method, human primary muscle cells are cultured in vitro to obtain differentiated myotubes and glucose uptake rates are measured. We provide a detailed protocol to quantify rates in basal and insulin-stimulated states using radiolabeled [3H] 2-deoxy-D-Glucose.

Mesure de l&#39;absorption de glucose et réponse à la stimulation de l&#39;insuline dans<em&gt; In Vitro</em&gt; Myotubes primaires humains cultivés

Skeletal muscle is the largest glucose deposit in mammals and largely contributes to glucose homeostasis. Assessment of insulin sensitivity of muscle ...

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

Électrophysiologie germes entiers de cellules entérochromaffines murins cultivés primaires

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...