This work was supported by NIH P20GM103643 to I. Meng.

1. Préparation des larves <ol><li> Pour analyser une larve souhaitée pour la locomotion ou la préférence de positionnement, de grandir larves dans des conditions standard pour l&#39;âge souhaité pour doser 10 à l&#39;aide alimentaire à la mouche norme 11. </li><li> Faites un filtre à mailles en étirant nylon sérigraphie de qualité maille sur un entonnoir. Fixez le maillage au niveau du col de l&#39;entonnoir avec une bande élastique. Placez l&#39;entonnoir dans un bêcher. </li><li> Pour collecter des larves pour l&#39;analyse, ramasser une spatule-plein de nourriture contenant la recherche de nourriture des larves de la bouteille de culture et laver avec de l&#39;eau du robinet RT sur le filtre de maille, la collecte de larves individuelle directement à partir de la maille avec un pinceau 10. </li></ol> 2. Préparation d&#39;Assay Tubes <ol><li> Pour se préparer à boucher les tubes à essai, faire bouillir avec soin un gel d&#39;agar à 4% et verser dans un plat de Petri à une profondeur de 1,5 cm. Cela fournira une fiche pour chaque côté du tube lorsque l&#39;animal est enfermé. La solution à 4% est suffisamment dense pour empêcher les larves de penetrating les bouchons. </li><li> Veiller à ce que les tubes sont propres et claires avant l&#39;insertion de larve. Si elles ne sont pas, cela peut bloquer les mouvements de cours d&#39;enregistrement. </li><li> Pour assurer larves ont suffisamment d&#39;humidité pour le mouvement, préparer un flacon souple contenant une petite quantité d&#39;eau chaude. Retourner le flacon, orienter soigneusement la sortie vers un évier et presser doucement pour expulser l&#39;eau du tube de ramassage. </li><li> Insérer la sortie de la bouteille dans le tube de dosage. Pressez la bouteille pour fournir de la vapeur d&#39;eau dans le tube à essai jusqu&#39;à ce qu&#39;une fine couche de la condensation apparaît sur la paroi du tube. Vidéo 1 représente une larve se déplaçant dans un tube avec une quantité appropriée de l&#39;humidité. </li><li> Pour sceller la larve dans le tube, enlever le gel d&#39;agar 1,5 cm d&#39;épaisseur de la boîte de Pétri et placez sur une surface de grillage pour permettre aux flux d&#39;air sous l&#39;agar de sorte qu&#39;un bouchon de gélose peut être inséré dans le tube. Appuyer sur une extrémité du tube à essai dans le gel deux fois pour insérer deux bouchons de gel dans une extrémité. </li><li> Avec un paintbprécipiter, placer une larve dans le tube d&#39;environ 1,5 pouces de profondeur et sceller le tube en appuyant sur la fin proche de la larve dans le gel d&#39;agar. La pression résultante va forcer le second bouchon de gel de gélose à partir de l&#39;extrémité opposée de l&#39;animal et sceller l&#39;intérieur du tube. </li><li> Placer les tubes dans le dispositif MB5 multi-Beam Drosophila Activity Monitor (DAM), et ajuster les positions des tubes de sorte que les animaux ne peuvent pas aller au-delà de la portée des capteurs. </li><li> Pour veiller à ce que dosage tubes ne glissent pas hors du cadre de lecture infrarouge de l&#39;appareil pendant le transport, maintenir le tube en place par la fixation d&#39;un anneau de mastic autour de chaque tube où il est en contact de l&#39;appareil d&#39;enregistrement. </li></ol> 3. mesure de l&#39;activité <ol><li> l&#39;activité d&#39;enregistrement dans un incubateur pour éviter des lectures erronées qui peuvent survenir en raison d&#39;ombres, les lumières fluorescentes ou aux variations de température dans le laboratoire. Voir la figure 5 pour un agencement proposé du système. </li><li> Définir un incubateur à 20 ° C (voir Figure 1). Pour éviter les faux enregistrements de perturbations dues à des sources lumineuses incandescentes pendant l&#39;enregistrement, éteindre les lumières fluorescentes de pépinières et d&#39;utiliser une source de lumière LED séparée lors de l&#39;enregistrement dans l&#39;incubateur. Effectuer un essai sans animaux pour assurer que les conditions d&#39;éclairage ne déclenche pas aberrante capteurs. Il devrait y avoir pas de données de mouvement enregistrées après ce test. </li><li> Permettre aux larves de acclimater à 20 ° C les paramètres de l&#39;incubateur pendant 5 min avant le début de l&#39;essai. </li><li> Pour définir le système de DAM à intervalles d&#39;enregistrement désirée (par exemple, 1 min), assurer le logiciel d&#39;enregistrement du système DAM est téléchargé sur l&#39;ordinateur hôte 12 et ouvrez le fichier du système de DAM avant de connecter la chambre d&#39;enregistrement à l&#39;interface Psiu. </li><li> Pour définir la fréquence souhaitée d&#39;enregistrement à laquelle les données seront sauvegardées, sélectionnez Préférences, puis cliquez dessus ou en dessous de l&#39;option d&#39;intervalle de lecture pour sélectionner différentes échelles de temps dans lequel les donnéesseront stockées. Par exemple, sélectionnez la lecture temps de pause de 1 sec à 1 h. </li><li> Pour sélectionner le paramètre variable pour l&#39;enregistrement de données, choisissez Préférences puis cochez les cases correspondantes dans la section Sortie type de données (comptes, mouvements, positions et vous habiterez, voir le tableau 1). Chaque paramètre fournit une analyse unique de l&#39;activité des larves à l&#39;intérieur du tube. </li><li> Pour commencer l&#39;enregistrement, connecter le moniteur à l&#39;unité d&#39;interface d&#39;alimentation (de Psiu) en utilisant le câble CAB6 téléphonique. Connectez le Psiu à une prise de courant. Une lumière verte indiquera un branchement approprié. </li><li> Connectez le Psiu par un câble Universal Serial Bus (USB) à un Macintosh ou PC sous Windows pour l&#39;enregistrement de données. Ouvrez le programme DAM SystemMB1v6x sur l&#39;ordinateur de démarrer automatiquement l&#39;enregistrement de données. </li><li> Après le temps de collecte souhaitée est terminée, sélectionnez Quitter, puis Quitter maintenant sur l&#39;écran de l&#39;activité de DAM; données seront ensuite automatiquement enregistrée sous Données système DAM. Prenez ce fichier de données (par exemple, surveiller 1) et faites glisser<html> dans un dossier distinct. Cela permettra de stocker les données brutes à partir du logiciel de DAM afin qu&#39;ils puissent être traitées par la suite. </li></ol> 4. Préparation des données pour le traitement (DAM FileScan) <ol><li> Pour traiter les données brutes en un format compréhensible ouvrir l&#39;application DAM FileScan et sélectionnez Sélectionner le dossier des données d&#39;entrée. Ensuite, choisissez le dossier de données et le fichier souhaité, et sélectionnez l&#39;option de numérisation. Enfin sélectionner la longueur de bin à la période de lecture souhaitée. Ce sera d&#39;organiser les données brutes en paramètres fixés par l&#39;opérateur et le programme va rapporter les données recueillies dans cette gamme particulière. </li><li> Sélectionnez le type de données de sortie à analyser le paramètre de mouvement souhaitée (nombre de moniteurs, se déplace, habitent). Dans le menu des lectures supplémentaires, sélectionnez somme dans le bac (ce qui est si la longueur du bac est plus grande que le système intervalle de lecture sélectionnée). </li><li> Nommez le fichier de manière appropriée et économisez. Le fichier sera stocké dans le même emplacement que le dossier de l&#39;étape 3.9. </li></ol>_title &quot;&gt; 5. Accès aux données d&#39;analyse <ol><li> Afin de recueillir des mouvements moyens, processus, les données brutes générées par l&#39;étape 4. Pour générer une table dans un tableur, ouvrez le fichier qui a été précédemment enregistrée (étape 4.3), et lorsque la fenêtre de l&#39;assistant d&#39;importation de texte apparaît, sélectionnez Terminer pour ouvrir les fichiers de données dans un format de tableur. Remarque: Selon le type de données analysées, la feuille de calcul sera lu différemment. Typiquement pour mesurer se déplace du moniteur ou chefs d&#39;accusation, la période de l&#39;étude sera enregistrée numériquement, tandis que chaque tube individuel de lecture recevra une lettre désigné. Lors de la visualisation des données dans le tableur, KZ colonnes représentent les fentes pour tubes à essai de 1 à 16, respectivement. Les lignes représentent les points de données collectés. </li><li> Par exemple, si se déplace ont été recueillies pendant 20 min à 1 min d&#39;intervalle, la moyenne des 20 lignes de calculer un nombre moyen de coups / minute et d&#39;autres mesures. </li></ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+disease+models+in+Drosophila.+melanogaster.and+the+role+of+the+fly+in+therapeutic+drug+discovery.">Human disease models in Drosophila. melanogaster.and the role of the fly in therapeutic drug discovery.</a> Pharmacol. Rev. 63 (2), 411-436 (2011).</li><li>Cattaert, D., Birman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blockade+of+the+central+generator+of+locomotor+rhythm+by+noncompetitive+NMDA+receptor+antagonists+in+Drosophila.larvae.">Blockade of the central generator of locomotor rhythm by noncompetitive NMDA receptor antagonists in Drosophila.larvae.</a> J. Neurobiol. 48 (1), 58-73 (2001).</li><li>Caldwell, J. C., Miller, M. M., Wing, S., Soll, D. R., Eberl, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+analysis+of+larval+locomotion+in+Drosophila.+hordotonal+organ+mutants.">Dynamic analysis of larval locomotion in Drosophila. hordotonal organ mutants.</a> PNAS. 100 (26), 16053-16058 (2003).</li><li>Nichols, C. D., Bechnel, J., Pandey, U. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+to+assay+Drosophila+behavior.">Methods to assay Drosophila behavior.</a> J. Vis. Exp. (61), (2012).</li><li>Godoy-Herrera, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+and+genetics+of+digging+behavior+in+Drosophila.+arvae.">The development and genetics of digging behavior in Drosophila. arvae.</a> Heredity. 56, 33-41 (1986).</li><li>Sinadinos, C., Cowan, C. M., Wyttenbach, A., Mudher, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+throughput+assays+of+locomotor+dysfunction+in+Drosophila.+larvae.">Increased throughput assays of locomotor dysfunction in Drosophila. larvae.</a> Journal of Neuroscience Methods. 203 (2), 325-334 (2012).</li><li>Homyk, T., Sheppard, D. 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Science Education Database Available from: <a target="_blank" href="https://www.jove.com/science-education-database/3/essentials-of-biology-1-yeast-drosophila-and-c-elegans">https://www.jove.com/science-education-database/3/essentials-of-biology-1-yeast-drosophila-and-c-elegans</a> (2015)</li><li>Woods, J. K., Kowalski, S., Rogina, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+the+spontaneous+locomotor+activity+in+Drosophila+melanogaster.">Determination of the spontaneous locomotor activity in Drosophila melanogaster.</a> J. Vis. Exp. (86), e51449 (2014).</li><li>Troncoso, B., Godoy-Herrera, R., Waldo, 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+development+of+larval+movement+patterns+in+Drosophila).">The development of larval movement patterns in Drosophila).</a> Heredity. 58 (1), 321-329 (1987).</li><li>Helfrich, C., Engelmann, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circadian+rhythm+of+the+locomotor+activity+in+Drosophila.+melanogaster.and+its+mutants+&#8216;sine+oculis&#8217;+and+&#8216;small+optic+lobes.">Circadian rhythm of the locomotor activity in Drosophila. melanogaster.and its mutants &#8216;sine oculis&#8217; and &#8216;small optic lobes.</a> Physiological Entomology. 8 (3), 257-272 (1983).</li><li>Johnson, A. W., Carder, W. 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The authors declare that they have no competing financial interests.

L&#39;activité des larves de Drosophila est influencée par divers facteurs, y compris génotype 8, 13 l&#39;âge et de la température ambiante 2. Bien que les méthodes vidéographiques puissants capables d&#39;analyse très détaillée ont été développés par ceux qui étudient la locomotion 5, ce niveau de détail peut être superflu pour ceux qui souhaitent pour déterminer les paramètres de base de l&#39;activité. La méthode décrite ici emploie un appareil qui est ...

The use of Drosophila as a genetic tool has transformed scientific knowledge of biological systems. Drosophila larvae have been used in a variety of studies including nociception1, development2 and as a model for the study of human disease genes3. Drosophila activity encompasses a range of behaviors that vary under different conditions including temperatures2, exposure to drugs4 and amongst different genotypes. Yet, despite the significant use of the larva as a model organism, a simple, standardized method to analyze larval activity has not been available. Presently, many larval locomotion studies employ sophisticated video analysis software5. While powerful, the complexity of such automated tools may discourage labs that are not already equipped to study locomotion from including analysis of informative activity parameters in their studies. In other current non-automated methods, such as the grid crawling analysis, motion is scored by a human observer, which introduces the possibility of subjectivity and limits throughput to one animal at a time6-7. A similar study used a 5-lane crawling assay, which measured the time it took larvae to travel a certain linear distance8. In such non-automated assays, displacement is measured but this does not account for non-linear travel between the start and end points. As discussed below, the method described here accounts for more of the actual larval movement, is objective, easy to operate, and offers robust throughput.
To easily study larval activity behavior without the compromise of accuracy, efficiency, or cost, this method employs the TriKinetics Drosophila Activity Monitor (DAM), a device often used to study adult activity. Using one device to study both adult and larval activity is cost-effective, and allows direct comparison of motion by animals at these two life stages. The system, featuring the manufacturer&#8217;s highest level of resolution, makes use of 17 infrared detection beams per assay tube, which record larval activity when sensor beams are broken within the 16 individual tubes. The system then automatically saves recorded information to a computer, making it available for manipulation with basic graphing software. The data obtained represents the beams that were broken by individual larvae (which can be converted into a rate), movement when a larva stays within a detector beam and the position of the animals within the assay chambers during a recording period (allowing one to calculate position preference). The system is efficient and relatively simple to operate, and brings highly reproducible basic activity analysis within the reach of any laboratory studying Drosophila larvae.
To demonstrate the power of this assay, data are presented that show its use to verify differences in activity resulting from varying ambient temperatures, as well as through the comparison of a mutant previously described as hypoactive (iav1)9 with a widely analyzed control (w1118).

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Drosophila Activity Monitor, Multibeam, 16 tubes, including wires</td>
			<td>TriKinetics Inc.&#160;</td>
			<td>MB5</td>
			<td></td>
		</tr>
		<tr>
			<td>Power Supply Interface for Activity Monitor&#160;</td>
			<td>TriKinetics Inc.&#160;</td>
			<td>PSIU24</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass 80 x 5 mm tubes for Activity Monitor (100)</td>
			<td>TriKinetics Inc.&#160;</td>
			<td>PGT 5x80</td>
			<td></td>
		</tr>
		<tr>
			<td>DAMsystemMB1v6x Data Acquisitions Software for Macintonsh OSX (Intel)</td>
			<td><a href="http://www.trikinetics.com/">www.trikinetics.com</a></td>
			<td></td>
			<td>free download</td>
		</tr>
		<tr>
			<td>DAMFileScan 108x software for Macintosh</td>
			<td><a href="http://www.trikinetics.com/">www.trikinetics.com</a></td>
			<td></td>
			<td>free download</td>
		</tr>
		<tr>
			<td>USB software (PSIUdrivers.zip).</td>
			<td><a href="http://www.trikinetics.com/">www.trikinetics.com</a></td>
			<td></td>
			<td>free download</td>
		</tr>
		<tr>
			<td>DAMSystem Notes 308</td>
			<td><a href="http://www.trikinetics.com/">www.trikinetics.com</a></td>
			<td></td>
			<td>free download</td>
		</tr>
		<tr>
			<td>Zeiss Stemi 2000C- Stereo Microscope</td>
			<td>Spectra Services</td>
			<td>SP-STEMI2000C-BS</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbon Dioxide</td>
			<td>Maine Oxy</td>
			<td></td>
			<td>anaesthesia</td>
		</tr>
		<tr>
			<td>Fly Pad</td>
			<td>Genesee</td>
			<td>59-114</td>
			<td>surface for sorting anaesthetized flies</td>
		</tr>
		<tr>
			<td>Small paint brush&#160;</td>
			<td>Winsor &#38; Newton</td>
			<td>#2 ROUND</td>
			<td>or similar, used for sorting anaesthetized flies</td>
		</tr>
		<tr>
			<td>Silk Screen Printing Mesh (160)</td>
			<td>msj-gallery.com</td>
			<td>SM160W63-3YD</td>
			<td>pore sized used in this protocol was ~ 0.1 mm</td>
		</tr>
		<tr>
			<td>Tegosept</td>
			<td>Genesee</td>
			<td>20-258</td>
			<td>preservative</td>
		</tr>
		<tr>
			<td>Ethanol (190proof)</td>
			<td>Pharmco</td>
			<td>111000190</td>
			<td>used to dissolve Tegosept</td>
		</tr>
		<tr>
			<td>6 oz Square Bottom Bottle (PP)</td>
			<td>Genesee</td>
			<td>32-130</td>
			<td></td>
		</tr>
		<tr>
			<td>&#34;Flugs&#34; for Plastic Fly bottles</td>
			<td>Genesee</td>
			<td>49-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Drosophila Vials, Wide (PS)</td>
			<td>Genesee</td>
			<td>32-117</td>
			<td></td>
		</tr>
		<tr>
			<td>Flugs for wide plastic vials</td>
			<td>Genesee</td>
			<td>49-101</td>
			<td></td>
		</tr>
		<tr>
			<td>Yellow Degerminated Corn Meal</td>
			<td>Gold Medal</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Drosophila agar</td>
			<td>LabScientific</td>
			<td>FLY 8020</td>
			<td></td>
		</tr>
		<tr>
			<td>Baker&#39;s Yeast - Red Star</td>
			<td>King Arthur Flour</td>
			<td>1270</td>
			<td></td>
		</tr>
		<tr>
			<td>Granulated Sugar - Extra Fine</td>
			<td>Domino</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

measurement of larval activity in the drosophila activity monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

La figure 1 montre les résultats d&#39;une étude de réponse à la température de contrôle des larves de troisième stade larvaire, 1118 w, à l&#39;aide du dispositif de surveillance pour détecter des différences dans la locomotion des larves à sept températures différentes. Les larves ont été lavées et placées dans le dispositif de l&#39;activité de DAM comme décrit ci-dessus, et placé dans un incubateur réglé à la température désirée. L&#39;appareil a ensuite été la...

Watch this Scientific Journal Video about Measurement of Larval Activity in the Drosophila Activity Monitor at JoVE.com

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Mesure de larves activité dans le<em&gt; Drosophila</em&gt; Moniteur d&#39;activité

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...

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Measurement of Larval Activity in the Drosophila Activity Monitor

9.3K vues.  University of New England. This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Vidéo: Measurement of Larval Activity in the Drosophila Activity Monitor

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 ...

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Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

Drosophile larvaire JNM Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Drosophila Larval NMJ Immunohistochemistry

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 31 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. Immunohistochemistry can be used to clearly image the components of the NMJ. In this article, we demonstrate how to use antibody staining to visualize the Drosophila larval NMJ.

Drosophila Larval NMJ Immunohistochemistry

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

Drosophile larvaire JNM immunohistochimie

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Dissection et la coloration des Drosophile Ovaires larvaires

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

Détection directe de l&#39;activité Acétate de formation de l&#39;acétate kinase Enzyme

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

Assaying the Kinase Activity of LRRK2 in vitro

Leucine Rich Repeat Kinase 2 (LRRK2) is a 2527 amino acid member of the ROCO family of proteins, possessing a complex, multidomain structure including a GTPase domain (termed ROC, for Ras of Complex proteins) and a kinase domain1. The discovery in 2004 of mutations in LRRK2 that cause Parkinson's disease (PD) resulted in LRRK2 being the focus of a huge volume of research into its normal function and how the protein goes awry in the disease state2,3. Initial investigations into the function of LRRK2 focused on its enzymatic activities4-6. Although a clear picture has yet to emerge of a consistent alteration in these due to mutations, data from a number of groups has highlighted the importance of the kinase activity of LRRK2 in cell death linked to mutations7,8. Recent publications have reported inhibitors targeting the kinase activity of LRRK2, providing a key experimental tool9-11. In light of these data, it is likely that the enzymatic properties of LRRK2 afford us an important window into the biology of this protein, although whether they are potential drug targets for Parkinson's is open to debate.
A number of different approaches have been used to assay the kinase activity of LRRK2. Initially, assays were carried out using epitope tagged protein overexpressed in mammalian cell lines and immunoprecipitated, with the assays carried out using this protein immobilised on agarose beads4,5,7. Subsequently, purified recombinant fragments of LRRK2 in solution have also been used, for example a GST tagged fragment purified from insect cells containing residues 970 to 2527 of LRRK212. Recently, Dani&euml;ls et al. reported the isolation of full length LRRK2 in solution from human embryonic kidney cells, however this protein is not widely available13. In contrast, the GST fusion truncated form of LRRK2 is commercially available (from Invitrogen, see table 1 for details), and provides a convenient tool for demonstrating an assay for LRRK2 kinase activity. Several different outputs for LRRK2 kinase activity have been reported. Autophosphorylation of LRRK2 itself, phosphorylation of Myelin Basic Protein (MBP) as a generic kinase substrate and phosphorylation of an artificial substrate - dubbed LRRKtide, based upon phosphorylation of threonine 558 in Moesin - have all been used, as have a series of putative physiological substrates including &alpha;-synuclein, Moesin and 4-EBP14-17. The status of these proteins as substrates for LRRK2 remains unclear, and as such the protocol described below will focus on using MBP as a generic substrate, noting the utility of this system to assay LRRK2 kinase activity directed against a range of potential substrates.

Assaying the Kinase Activity of LRRK2 in vitro

Leucine Rich Repeat Kinase 2 is a large multidomain kinase, mutations in which are the most common genetic cause of Parkinson's disease. Analysis of the kinase activity of this protein has proven to be a crucial tool in understanding the biology and dysfunction of this protein. In this paper, in vitro assaying of the kinase activity of LRRK2 and a selection of its mutants is described, providing an experimental system to examine phosphorylation of putative substrates and potential dysfunction of LRRK2 in disease.

Dosage de l&#39;activité kinase de LRRK2 In vitro</em

Leucine Rich Repeat Kinase 2 (LRRK2) is a 2527 amino acid member of the ROCO family of proteins, possessing a complex, multidomain structure including a ...

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 ...

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of genetic manipulations with this model has supplanted pharmacological approaches that are commonplace in other animal models and cell-based assays. Here we describe recent advances in a protocol that enables application of small molecules to the developing fruit fly embryo. The method details steps to overcome the impermeability of the eggshell while maintaining embryo viability. Eggshell permeabilization across a broad range of developmental stages is achieved by application of a previously described d-limonene embryo permeabilization solvent (EPS1) and by aging embryos at reduced temperature (18 &#176;C) prior to treatments. In addition, use of a far-red dye (CY5) as a permeabilization indicator is described, which is compatible with downstream applications involving standard red and green fluorescent dyes in live and fixed preparations. This protocol is applicable to studies using bioactive compounds to probe developmental mechanisms as well as for studies aimed at evaluating teratogenic or pharmacologic activity of uncharacterized small molecules.

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

Introduction of small molecules to the developing Drosophila embryo offers great potential for characterizing biological activity of novel compounds, drugs, and toxins as well as for probing fundamental developmental pathways. Methods described herein outline steps that overcome natural barriers to this approach, expanding the utility of the Drosophila embryo model.

Une méthode de perméabilisation de<em&gt; Drosophila</em&gt; Embryons pour dosages de petites molécules activité

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

Reconstitution fonctionnelle et Activité du Chan mesures de type sauvage purifiée et Mutant protéine CFTR

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

In Vivo Biosensor Tracks Non-apoptotic Caspase Activity in Drosophila

Caspases are the key mediators of apoptotic cell death via their proteolytic activity. When caspases are activated in cells to levels detectable by available technologies, apoptosis is generally assumed to occur shortly thereafter. Caspases can cleave many functional and structural components to cause rapid and complete cell destruction within a few minutes. However, accumulating evidence indicates that in normal healthy cells the same caspases have other functions, presumably at lower enzymatic levels. Studies of non-apoptotic caspase activity have been hampered by difficulties with detecting low levels of caspase activity and with tracking ultimate cell fate in vivo. Here, we illustrate the use of an ultrasensitive caspase reporter, CaspaseTracker, which permanently labels cells that have experienced caspase activity in whole animals. This in vivo dual color CaspaseTracker biosensor for Drosophila melanogaster transiently expresses red fluorescent protein (RFP) to indicate recent or on-going caspase activity, and permanently expresses green fluorescent protein (GFP) in cells that have experienced caspase activity at any time in the past yet did not die. Importantly, this caspase-dependent in vivo biosensor readily reveals the presence of non-apoptotic caspase activity in the tissues of organ systems throughout the adult fly. This is demonstrated using whole mount dissections of individual flies to detect biosensor activity in healthy cells throughout the brain, gut, malpighian tubules, cardia, ovary ducts and other tissues. CaspaseTracker detects non-apoptotic caspase activity in long-lived cells, as biosensor activity is detected in adult neurons and in other tissues at least 10 days after caspase activation. This biosensor serves as an important tool to uncover the roles and molecular mechanisms of non-apoptotic caspase activity in live animals.

In Vivo Biosensor Tracks Non-apoptotic Caspase Activity in Drosophila

To detect healthy cells in whole animals that contain low levels of caspase activity, the highly sensitive biosensor designated CaspaseTracker was generated for Drosophila. Caspase-dependent biosensor activity is detected in long-lived healthy cells throughout the internal organs of adult animals reared under optimized conditions in the absence of death stimuli.

In Vivo Biocapteur Tracks apoptotique non-activité Caspase chez la drosophile

Caspases are the key mediators of apoptotic cell death via their proteolytic activity. When caspases are activated in cells to levels detectable by ...