Ce protocole est actuellement utilisé dans notre laboratoire et a été affiné au fil des années par de nombreuses personnes, y compris les Drs David Sharp, Mijung Kwon, Patrizia SOMMI, Dhanya Cheerambathur. Nous remercions le Dr Bill Sullivan (UCSC) qui nous a fourni d&#39;excellents conseils sur la manipulation et la micro-injection d&#39;embryons de drosophile tôt quand notre travail dans ce système a été lancé (réf. 13). Nous remercions tous les membres du laboratoire Scholey. Notre travail sur la mitose chez la drosophile est soutenu par NIH GM55507.

Recettes: Plaques Jus de raisin: <ul><li> 5.5g bacto agar </li><li> 14,5 g de dextrose ou de glucose </li><li> 7,15 g de saccharose </li><li> 45 ml de concentré de jus de raisin (jus 100%). </li><li> 204,5 ml d&#39;H 2 0 </li><li> 625 ul de NaOH 10N </li></ul> Mélanger tous les ingrédients et les micro-ondes jusqu&#39;à ébullition. Ajouter 2,8 ml d&#39;acide mélange (mix d&#39;acide: 20,9 ml d&#39;acide propionique, 2,1 ml d&#39;acide phosphorique, 27 ml de H 2 0) Mélangez et versez sur 35 boîtes de Pétri mm. Laisser solidifier à température ambiante pendant un ou deux jours. Si les plaques ne vont pas être utilisés rapidement, sceller avec du parafilm et de garder à 4 ° C (laisser s&#39;équilibrer à température ambiante avant de l&#39;utiliser). Pâte de levure: Dissolvez environ une cuillère à café de levure (Sigma YSC2, levure de Saccharomyces cerevisiae type II) dans l&#39;eau pour former une pâte épaisse. Placer une petite quantité de ce sur chaque assiette de jus de raisin, juste avant utilisation. G-PEM mémoire tampon: <ul><li> 80 mM de Na-PIPES, pH 6,9 </li><li> MgCl2 1 mM </li><li> 1 mM d&#39;EGTA </li><li> 1 mM GTP </li></ul> Tampon d&#39;injection: <ul><li> 150 mM de K-aspartate </li><li> 10 mM de K-phosphate </li><li> 20 mM imidazole, pH 7,2 </li></ul> Colle Heptane: Dérouler ruban adhésif double collant et le placer dans une bouteille 100ml, ajouter environ 50 ml d&#39;heptane, sceller la bouteille, et le rock pendant plusieurs jours. Lors de l&#39;utilisation, ajouter heptane si la colle est trop épaisse. Chambre de déshydratation: Prenez une boîte de Petri de 100 mm, mettre une partie d&#39;un plat de 35 mm à l&#39;intérieur pour faire une «table» et d&#39;ajouter Drierite (sulfate de calcium anhydre) autour de lui afin que la hauteur des Drierite n&#39;est pas plus élevé que la «table» et couvrir. La lamelle avec des embryons sera mis sur cette &quot;table&quot; pour déshydratation avant l&#39;injection. C&#39;est une bonne idée d&#39;utiliser au moins quelques Drierite indiquant et le changer quand il a changé de couleur. Protocole: Ce protocole peut être utilisé pour l&#39;injection de pratiquement tous les réactifs solubles dans l&#39;embryon de drosophile syncytial: Par exemple, soit une protéine fluorescente pour l&#39;observation ou un inhibiteur de la protéine cible ou les deux. Obtenir tout est prêt: 1 .- 2 à 3 jours avant l&#39;expérience, faire des plaques de jus de raisin et de mettre en place jeter en cage: Prendre une bouteille en plastique (6 oz, à fond rond de scientifique appliquée drosophile), couper deux petits trous (environ 1cm 2) sur les côtés opposés du bouteille et remplissez avec un morceau de coton (ce qui permettra une circulation d&#39;air), appuyez sur les mouches dans la bouteille et couvrir avec une assiette de jus de raisin. Gardez à 25 ° C (ou température ambiante), les mouches devraient définir les embryons pendant environ 10 jours. 2 .- Au moins un jour avant l&#39;expérience tirez aiguilles pour l&#39;injection, on utilise une aiguille Kopf / pipette extracteur de modèle 720 et minces filaments de verre à paroi (World Precision Instruments tw100f). Pour l&#39;injection de tubuline fluorescente, gardez aiguilles à 4 ° C pour empêcher la température induite par polymérisation dans l&#39;aiguille. Préparation de la tubuline: Remarque: Tous les travaux avec la tubuline devrait être fait à 4 ° C pour empêcher la polymérisation. <ol><li> Prenez la tubuline étiquetés hors du congélateur et le mettre sur la glace pendant 5 à 15 minutes. </li><li> Diluer avec le G-PEM tampon (4 à 10 fois selon l&#39;intensité requise). </li><li> Centrifuger à 13k rpm à 4 ° C pendant 10-15 minutes. </li><li> Charge de 1 à 2 ul dans l&#39;aiguille. </li></ol> Anticorps ou d&#39;un inhibiteur chimique: Préparer aiguilles comme pour la tubuline, en utilisant une concentration appropriée (ce peut être nécessaire de tester). L&#39;anticorps peut être dans le tampon d&#39;injection, le G-PEM tampon ou PBS. Si un autre tampon est nécessaire, ce tampon devra être injectée comme un contrôle pour s&#39;assurer que ce tampon provoque pas de dégâts sur son propre. Remarque: la tubuline inhibiteur et fluorescentes peuvent être mélangés et injectés ensemble, si les concentrations le permettent. Sinon, injecter la tubuline fluorescente et attendre 5 à 10 minutes avant l&#39;injection de l&#39;inhibiteur. Lorsque vous effectuez une double injection, commencer avec plus d&#39;embryons, car moins d&#39;embryons va récupérer normalement. Collecte d&#39;embryons: <ol><li> Mettez une plaque de raisin jus de nouvelles sur la cage laïcs. </li><li> Retirez cette plaque au bout d&#39;une heure, il s&#39;agit de la première collection de la journée et ne sont souvent pas très bon. Il peut être mis au rebut. </li><li> Changer la plaque toutes les heures pour garder les embryons de collecte pour une heure chacune. </li></ol> Préparation de la lamelle: <ol><li> Mettre une lamelle de 50 x 22mm sur un côté d&#39;une lame de microscope. Bande des quatre coins de la lame de sorte qu&#39;il ne bouge pas. </li><li> En utilisant un coton-tige, placez une couche de colle heptane en une seule ligne sur la lamelle (la colle ne doit pas être visqueux et doit sécher en quelques secondes, sinon ajouter plus heptane). </li><li> Placer un morceau dedoubles ruban adhésif sur la lame vers le côté de la lamelle. </li></ol> Préparation de l&#39;embryon: Remarque: Les embryons doivent être imagés environ 2 heures après le début de la collecte, afin de commencer les étapes suivantes laissant suffisamment de temps pour les terminer dans ce laps de temps. <ol><li> Avec un pinceau imbibé soigneusement ramasser les embryons de la plaque de jus de raisin et le placer sur un ruban adhésif double glissière. </li><li> Utilisation de la partie externe de la pince à épiler, rouler les embryons au cours du ruban adhésif double collant jusqu&#39;à la chorion (la membrane externe) Séjours ouverts. </li><li> Ramassez l&#39;embryon par la doucement rouler sur le chorion de sorte qu&#39;il colle à la pince à épiler et le placer sur la colle heptane sur la lamelle avec le côté long du parallèle embryon pour le côté long de la lamelle. Réglez hausse de 10 à 20 embryons dans une rangée. </li><li> Retirer la lamelle et la placer dans la chambre de déshydratation pour les 3 à 8 minutes (cela dépend de l&#39;humidité de la pièce et la quantité à injecter). </li><li> Placer sur une enceinte métallique avec de la graisse à vide. </li><li> Couvrir les embryons avec 700 huile halocarbone pour éviter de nouvelles déshydratation. Les embryons sont maintenant prêtes pour l&#39;injection. </li></ol> L&#39;injection d&#39;embryons: <ol><li> Trouvez les embryons dans un objectif 16x. </li><li> Déplacer les embryons loin, et de trouver l&#39;aiguille sans déplacer le plan focal. </li><li> Centre de l&#39;aiguille et le déplacer en place sans la déplacer en direction X ou Y. </li><li> Si l&#39;aiguille n&#39;est pas ouvert, placez le bord de la lamelle dans le champ de vision, mais pas là où l&#39;aiguille sera, et abaisser l&#39;aiguille au même plan focal. Très soigneusement déplacer la lamelle jusqu&#39;à ce qu&#39;il frappe l&#39;aiguille et la douceur des pauses de l&#39;ouvrir. Si l&#39;aiguille est ouvert, passez à l&#39;étape 5. (Aiguilles peuvent être ouvertes avec de l&#39;acide fluorhydrique, avant le remplissage). </li><li> Mettez les embryons en vue (mais pas là où l&#39;aiguille sera), abaisser l&#39;aiguille dans l&#39;huile et assurez-vous d&#39;obtenir des gouttes de liquide bien de l&#39;aiguille. </li><li> Soigneusement mais sûrement passer l&#39;embryon dans l&#39;aiguille et injecter une goutte dans l&#39;embryon et l&#39;embryon se déplacer loin. (Ou suivez les instructions de votre appareil d&#39;injection). </li><li> Après que tous les embryons ont été injectés, ils sont prêts pour l&#39;observation sur un microscope confocal. </li></ol> Imagerie: Image des embryons sur un microscope confocal avec un objectif de 60 ou 100X: Notes: <ol><li> L&#39;intervalle de temps sur une série laps de temps devrait être au plus de quelques secondes, depuis la mitose chez l&#39;embryon est très rapide, selon le processus exact d&#39;être étudiée. </li><li> Une série en 3D ou un seul plan peut être acquis en fonction du procédé d&#39;intérêt. </li><li> Pour les inhibiteurs à action rapide, le transfert du microscope injection microscope d&#39;imagerie doit être rapide. </li></ol><img src="/files/ftp_upload/1382/1382fig1.jpg" alt="Figure 1" /> Figure 1: Étude de la mitose dans la drosophile embryon syncytial. Noyaux dans l&#39;embryon précoce subissent des divisions rapides sans cytokinèse, lors des cycles de 10 à 13 former des noyaux d&#39;une monocouche au niveau du cortex. Une. Schéma de l&#39;embryon précoce avec des noyaux à cortex. Embryon peut exprimer la GFP et / ou DP protéines marquées et peuvent être microinjectés avec d&#39;autres protéines marquées et / ou inhibiteurs. Imagerie B. laps de temps de l&#39;embryon exprimant la GFP-tubuline et de la DP-histones. Plot C. pôle-pôle de séparation pour les cycles 11 , 12 et 13 dans un embryon de type sauvage. Longueur de la broche expositions des périodes de durée isométrique et des périodes d&#39;allongement, qui sont très cohérents et reproductibles. Dynamique des microtubules D. broche. L&#39;injection d&#39;une faible concentration de tubuline conduit à la rhodamine taches formées d&#39;un sous-unités de tubuline années, ce qui peut être utilisée pour étudier la dynamique des microtubules.

<ol>
	<li>Brust-Mascher, I., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitotic+spindle+dynamics+in+Drosophila.">Mitotic spindle dynamics in Drosophila.</a> Int Rev Cytol. 259, 139-172 (2007).</li><li>Duffy, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GAL4+system+in+Drosophila:+a+fly+geneticist's+Swiss+army+knife.">GAL4 system in Drosophila: a fly geneticist's Swiss army knife.</a> Genesis. 34, 1-15 (2002).</li><li>Morris, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+methods+for+analyzing+the+functions+of+kinesins+in+early+embryos.">Microinjection methods for analyzing the functions of kinesins in early embryos.</a> Methods Mol Biol. 164, 163-172 (2001).</li><li>Brust-Mascher, I., Sommi, P., Cheerambathur, D. K., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin-5-dependent+poleward+flux+and+spindle+length+control+in+Drosophila+embryo+mitosis.">Kinesin-5-dependent poleward flux and spindle length control in Drosophila embryo mitosis.</a> Mol Biol Cell. 20, 1749-1762 (2009).</li><li>Brust-Mascher, I., Civelekoglu-Scholey, G., Kwon, M., Mogilner, A., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Model+for+anaphase+B:+role+of+three+mitotic+motors+in+a+switch+from+poleward+flux+to+spindle+elongation.">Model for anaphase B: role of three mitotic motors in a switch from poleward flux to spindle elongation.</a> Proc Natl Acad Sci U S A. 101, 15938-15943 (2004).</li><li>Brust-Mascher, I., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubule+flux+and+sliding+in+mitotic+spindles+of+Drosophila+embryos.">Microtubule flux and sliding in mitotic spindles of Drosophila embryos.</a> Mol Biol Cell. 13, 3967-3975 (2002).</li><li>Cheerambathur, D. K., Brust-Mascher, I., Civelekoglu-Scholey, G., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+partitioning+of+mitotic+kinesin-5+crosslinkers+between+microtubule-bound+and+freely+diffusing+states.">Dynamic partitioning of mitotic kinesin-5 crosslinkers between microtubule-bound and freely diffusing states.</a> J Cell Biol. 182, 421-428 (2008).</li><li>Cheerambathur, 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=Quantitative+analysis+of+an+anaphase+B+switch:+predicted+role+for+a+microtubule+catastrophe+gradient.">Quantitative analysis of an anaphase B switch: predicted role for a microtubule catastrophe gradient.</a> J Cell Biol. 177, 995-1004 (2007).</li><li>Kwon, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chromokinesin,+KLP3A,+dives+mitotic+spindle+pole+separation+during+prometaphase+and+anaphase+and+facilitates+chromatid+motility.">The chromokinesin, KLP3A, dives mitotic spindle pole separation during prometaphase and anaphase and facilitates chromatid motility.</a> Mol Biol Cell. 15, 219-233 (2004).</li><li>Sharp, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+coordination+of+three+mitotic+motors+in+Drosophila+embryos.">Functional coordination of three mitotic motors in Drosophila embryos.</a> Mol Biol Cell. 11, 241-253 (2000).</li><li>Sharp, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+bipolar+kinesin,+KLP61F,+cross-links+microtubules+within+interpolar+microtubule+bundles+of+Drosophila+embryonic+mitotic+spindles.">The bipolar kinesin, KLP61F, cross-links microtubules within interpolar microtubule bundles of Drosophila embryonic mitotic spindles.</a> J Cell Biol. 144, 125-138 (1999).</li><li>Sharp, D. J., Rogers, G. C., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoplasmic+dynein+is+required+for+poleward+chromosome+movement+during+mitosis+in+Drosophila+embryos.">Cytoplasmic dynein is required for poleward chromosome movement during mitosis in Drosophila embryos.</a> Nat Cell Biol. 2, 922-930 (2000).</li><li>Sharp, D. J., Yu, K. R., Sisson, J. C., Sullivan, W., Scholey, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antagonistic+microtubule-sliding+motors+position+mitotic+centrosomes+in+Drosophila+early+embryos.">Antagonistic microtubule-sliding motors position mitotic centrosomes in Drosophila early embryos.</a> Nat Cell Biol. 1, 51-54 (1999).</li><li>Waterman-Storer, C. M., Desai, A., Bulinski, J. C., Salmon, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+speckle+microscopy,+a+method+to+visualize+the+dynamics+of+protein+assemblies+in+living+cells.">Fluorescent speckle microscopy, a method to visualize the dynamics of protein assemblies in living cells.</a> Curr Biol. 8, 1227-1230 (1998).</li></ol>

Ce protocole est relativement simple, cependant, chaque étape requiert la pratique de s&#39;assurer que les embryons ne sont pas endommagés. Expériences de contrôle minutieux toujours besoin d&#39;être fait pour s&#39;assurer que des résultats fiables sont obtenus. Une bonne façon de commencer ce n&#39;est par la micro-injection de tubuline tampon ou la rhodamine dans des embryons de contrôle exprimant la GFP-tubuline afin de s&#39;assurer que la mitose se déroule normalement dans tous les cycles à la surface ...

microinjection techniques for studying mitosis in the drosophila melanogaster syncytial embryo

Ce protocole décrit l&#39;utilisation de l&#39;embryon de Drosophila melanogaster pour l&#39;étude de la mitose syncytial 1. Drosophile a la génétique utile avec un génome séquencé, et il peut être facilement entretenu et manipulé. Il existe de nombreux mutants mitotique, et les mouches transgéniques exprimant fonctionnelle par fluorescence (par exemple GFP) - protéines mitotiques étiquetés ont été et sont générés. L&#39;expression du gène ciblé est possible en utilisant le système GAL4/UAS 2. La drosophile embryon précoce effectue mitoses multiples très rapidement (durée du cycle cellulaire, ≈ 10 min). Il est bien adapté pour l&#39;imagerie de la mitose, parce que pendant les cycles de 10-13, les noyaux se divisent rapidement et de manière synchrone, sans intervenir cytokinèse à la surface de l&#39;embryon dans une seule monocouche juste en dessous du cortex. Ces noyaux se divisent rapidement probablement utiliser la machinerie mitotique même que d&#39;autres cellules, mais ils sont optimisés pour la vitesse, le checkpoint est généralement admis de ne pas être rigoureux, permettant l&#39;étude des protéines mitotique dont l&#39;absence entraînerait l&#39;arrêt du cycle cellulaire dans les cellules avec un checkpoint forte . Embryons exprimant des protéines GFP étiqueté ou microinjectés avec des protéines marquées par fluorescence peut être facilement imagée de suivre la dynamique en direct (Fig. 1). En outre, les embryons peuvent être microinjectés avec fonction de blocage des anticorps ou des inhibiteurs de protéines spécifiques à étudier l&#39;effet de la perte ou la perturbation de leur fonction 3. Ces réactifs peuvent diffuser dans tout l&#39;embryon, atteignant plusieurs fuseaux pour produire un gradient de concentration de l&#39;inhibiteur, ce qui entraîne à son tour dans un gradient de défauts comparables à une série allélique de mutants. Idéalement, si la protéine cible est marqué par fluorescence, le gradient d&#39;inhibition peut être visualisé directement 4. Il est supposé que la plus forte phénotype est comparable à l&#39;phénotype nul, mais il est difficile d&#39;exclure formellement la possibilité que les anticorps peuvent avoir des effets dominante dans de rares cas, des contrôles afin rigoureuse et une interprétation prudente doit être appliquée. Plus loin du site d&#39;injection, la fonction des protéines n&#39;est que partiellement perdue permettant d&#39;autres fonctions de la protéine cible à devenir évidents.

Watch this Scientific Journal Video about Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo at JoVE.com

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Ce protocole décrit l&#39;utilisation de micro-injection et l&#39;imagerie à haute résolution dans le<em&gt; Drosophila melanogaster</em&gt; Embryon syncytial pour étudier la mitose.

Techniques de microinjection pour l&#39;étude de la mitose dans les Drosophila melanogaster Embryon syncytial

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

microinjection-techniques-for-studying-mitosis-drosophila

Research

JoVE Journal

Biology

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

14.0K vues.  University of California, Davis. Ce protocole décrit l'utilisation de micro-injection et l'imagerie à haute résolution dans le Drosophila melanogaster Embryon syncytial pour étudier la mitose.

Vidéo: Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

Dissection de SNC larvaire chez Drosophila melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Recherche

Biologie

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Suivi vidéo haute résolution de la locomotion chez les adultes Drosophila melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

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

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

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

Surveillance de la fonction cardiaque chez les larves Drosophila melanogaster Pour les études physiologiques

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

Patch Clamp et des techniques de perfusion pour étudier les canaux ioniques Exprimé en Xenopus Ovocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

Etudier Checkpoint mitotique en illustrant le comportement dynamique des protéines Kinetochore et Motion chromosomique chez Vivre<em&gt; Drosophila</em&gt; Les embryons syncytial

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Mesure de la durée de vie en<em&gt; Drosophila melanogaster</em

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

Imagerie à travers le cas de la nymphe<em&gt; Drosophila melanogaster</em

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular biology has been greatly fueled by the discovery of noncoding RNAs as major players in post-transcriptional regulation. Such a revolutionary perspective has been accompanied and triggered by the development of powerful technologies for profiling short RNAs expression, both at the high-throughput level (genome-wide identification) or as single-candidate analysis (steady state accumulation of specific species). Although several state-of-art strategies are currently available for dosing or visualizing such fleeing molecules, Northern Blot assay remains the eligible approach in molecular biology for immediate and accurate evaluation of RNA expression. It represents a first step toward the application of more sophisticated, costly technologies and, in many cases, remains a preferential method to easily gain insights into RNA biology. Here we overview an efficient protocol (Enhanced Northern Blot) for detecting weakly expressed microRNAs (or other small regulatory RNA species) from Drosophila melanogaster whole embryos, manually dissected larval/adult tissues or in vitro cultured cells. A very limited amount of RNA is required and the use of material from flow cytometry-isolated cells can be also envisaged.

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The aim of this publication is to visualize and discuss the operative steps of an Enhanced Northern Blot protocol on RNA extracted from Drosophila melanogaster embryos, cells, and tissues. This protocol is particularly useful for the efficient detection of small RNA species.

Amélioration de la détection du Nord Blot des petits ARN espèces en<em&gt; Drosophila melanogaster</em

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular ...

Generation of Transgenic Hydra by Embryo Microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Hydra is used as a model for the study of regeneration, pattern formation, and stem cells. However, research efforts have been hampered by lack of a reliable method for gene perturbations to study molecular function. The development of transgenic methods has revitalized the study of Hydra biology1. Transgenic Hydra allow for the tracking of live cells, sorting to yield pure cell populations for biochemical analysis, manipulation of gene function by knockdown and over-expression, and analysis of promoter function. Plasmid DNA injected into early stage embryos randomly integrates into the genome early in development. This results in hatchlings that express transgenes in patches of tissue in one or more of the three lineages (ectodermal epithelial, endodermal epithelial, or interstitial). The success rate of obtaining a hatchling with transgenic tissue is between 10% and 20%. Asexual propagation of the transgenic hatchling is used to establish a uniformly transgenic line in a particular lineage. Generating transgenic Hydra is surprisingly simple and robust, and here we describe a protocol that can be easily implemented at low cost.

Generation of Transgenic Hydra by Embryo Microinjection

Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi.