Wir möchten Astha Maltare zur Erzeugung der Larven kriechen Daten danken. Wir möchten Dr. Nicholas Lanson Jr. für das Geben seiner Anmerkungen zum Manuskript danken. Diese Arbeit wurde von der Robert-Packard Zentrum für ALS an der Johns Hopkins (bis UBP) und die Amyotrophe Lateralsklerose Association (UBP) unterstützt; und R01MH083689 von den National Institutes of Mental Health (CDN).

A. Larven Crawling-Test 1. Larven Sammlung <ol><li> Richten Sie eine 8 Unzen Flasche fliegt (10-15 + 10-15 Männchen Weibchen). </li><li> Lassen Fliegen legen ihre Eier für 24 Stunden, dann klare Flasche fliegt. (Übertragen Sie die Erwachsenen in eine neue Flasche und wiederholen wie nötig). </li><li> Inkubieren Flasche für 3-4 Tage, oder bis zum dritten Larvenstadium sichtbar sind. </li><li> In 50 bis 100 ml 20% Saccharose in die Flasche mit Larven und lassen Sie sich für 20 Minuten. Die Larven werden auf der Oberfläche schwimmen. </li><li> Sammle Larven mit einem 25 ml serologische Pipette mit der Spitze abgeschnitten, und in einen Gitterkorb. </li><li> Waschen Sie Larven im Gitterkorb zweimal mit deionisiertem H 2 O. Die Larven sind jetzt bereit für die Experimente. </li></ol> 2. Um Larven mit Medikamenten behandeln <ol><li> Verwenden Sie eine Bürste, um die gewünschte Anzahl der Larven zu einem 5-ml-Becher, die eine Lösung von 5% Saccharose + Droge transportieren. </li><li> Lassen Larven ernähren sich mindestens 15 Minuten.</li><li> Gießen Medikament behandelten Larven in einen Gitterkorb und spülen. Sie sind jetzt einsatzbereit. </li></ol> 3. Bewegungsapparat Assay (Messung insgesamt zurückgelegte Entfernung oder Körperwand Kontraktionen) <ol><li> Mit einer Bürste, einzelne Larve bis ein Transport: <ol><li> 15 cm Petrischale mit 2% Agarose (zuvor gegossen und ausgehärtet) über Millimeterpapier mit einer 0,2 cm 2 Rasters. <ol><li> Count Anzahl der Gitterlinien in 1 Minute überschritten. </li></ol></li><li> Nun von einem Glas Dissektion Schale mit einer verdünnten Lösung Hefepaste. <ol><li> Count Peristaltik Kontraktionen (von vorne nach hinten voller Bewegung = 1 Kontraktion) in einer Minute unter Beobachtung unter einem Dissektionsmikroskop. </li></ol></li></ol></li><li> Wiederholen, bis die gewünschte Anzahl von Larven gezählt worden. </li></ol> B. Schnelle iterative negativen Geotaxis (RING) Protokoll Dieser Test wurde ursprünglich debeschrieben durch Gargano et al 5. <ol><li> Sammeln frisch geschlüpften erwachsenen männlichen Fliegen unter leichtem CO 2-Betäubung und in einen Standard-Fläschchen mit Nahrungsmitteln (+ Arzneimittel oder Lebensmittel). </li><li> Halten Sie Fliegen bei Raumtemperatur (auf der Tischplatte. ~ 22 ° C) für 2-3 Tage auf die zur Verwertung von CO 2 (und Akkumulation von Steady-State-Plasmaspiegel gegebenenfalls) zu ermöglichen. </li><li> Übertragen Sie etwa 25 Fliegen ohne Betäubung zu Polystyrol vorbereiteten Fläschchen. </li><li> Montieren Fläschchen mit Fliegen in den RING Apparatur (Abbildung 1). </li><li> Erlauben Sie fliegt in die Umwelt, ungestört, für 15-20 Minuten akklimatisieren. </li><li> Während dieser Zeit Platz Digitalkamera ~ 1 m vor dem Gerät (auf einer Plattform, wenn notwendig, das Zentrum der Linse auf halber Höhe des Vials ausrichten), Fokus und Zoom der Kamera auf das Gerät, und einen Timer einstellen, um 3,0 Sekunden. </li><li> Entfernen Sie vorsichtig von der RING-Apparat zu halten mit der linken Hand, um nicht zu disturb die Fliegen, und halten Sie den Timer mit der rechten Hand. </li><li> Scharf tippen Sie auf das Gerät nach unten auf der Oberfläche der Bank dreimal, sicherzustellen, dass das Leitungswasser hart genug, um knock down all die Fliegen auf den Boden der Fläschchen ist. </li><li> Gleichzeitig mit Fertigstellung des dritten Hahn, starten Sie die 3 Sekunden Countdown-Timer. </li><li> Bei 3 Sekunden ein Bild aufnehmen. </li><li> Setzen Sie den Timer für 1 Minute und starten. Während dieser Zeit setzen Sie die Kamera und Fokus auf das Gerät, und stellen Sie einen anderen Kanal des Timers für drei Sekunden. </li><li> Nach 1 Minute, wiederholen Sie die Schritte 1,7-1,10 </li><li> Nach insgesamt 5-6 Versuche, laden Sie Bilder auf einen Computer und nutzen Sie Ihre Lieblings-Bildbetrachter zu öffnen, zu punkten und die durchschnittliche Höhe kletterten für jedes Fläschchen. </li><li> Statistische Analysen auf Ihrer verschiedenen Gruppen einen Vergleich der mittleren Höhe geklettert. </li></ol> C. Balz und Paarung Assay <ol><li> Als erstes am Morgen, klar, gut-produzierenden Flaschen FLIes verwendet werden soll. </li><li> Im Laufe des Tages (alle 3-4 Stunden), sammeln neu entstandenen sexuell naive Männchen und Weibchen: <ol><li> Platzieren Sie Männchen einzeln in Fläschchen oder Röhrchen mit Medium. </li><li> Platzieren Sie 5-6 Weibchen zusammen pro Fläschchen / Tube. </li><li> Isolieren Sie gesammelt Fliegen bei 25 ° C unter 12 Stunden Licht / Dunkel für 5 Tage. </li><li> Übertragen Sie eine Frau in die Kammer eines Gegenrad. </li><li> Übertragen Sie ein Männchen in die Kammer eines Gegenrad. </li><li> Beachten Paar unter einem Dissektionsmikroskop für die folgenden Verhaltensweisen: <ol><li> Orientierung (die männlichen orientiert sich an den weiblichen) </li><li> Tapping (klopft das Männchen das Weibchen) </li><li> Flügel Lied (das männliche erstreckt und schwingt einen Flügel) </li><li> Lecken (die männlichen leckt die weiblichen Genitalien) </li><li> Curling (das Männchen seinen Bauch rollt unter sich) </li><li> Kopulation Versuch (Curling Tätigkeit beim Versuch, den weiblichen montieren) </li></ol></li></ol></li><li> Beachten Sie für 10 Minuten oder bis zum erfolgreichen Kopulation, in Anbetracht der Zeit, zu der jedes Verhalten auftritt (Latenzzeit), (die gesamte Zeit in Balzverhalten bis Kopulation (die Brautwerbung Index zu berechnen), sowie die Anzahl der Paare, die erfolgreich ein bestimmtes Verhalten auszuführen engagiert Frequenz). 100% der Wildtyp-Paare werden in der Regel innerhalb von 5 Minuten zu paaren. </li><li> Berechnen Balz ein Index (CI) durch Division der Zeitaufwand bei der Balz durch die gesamte Zeit bis Kopulation unterteilt. Für Wildtyp-Paaren dies in der Regel im Bereich zwischen 0,6 bis 0,8. </li></ol> D. Repräsentative Ergebnisse Crawling-Assay Normale Wildtyp-Larve wird wandern ~ 3 cm / min und Ausstellungsflächen ~ 40-50 Körperwand Kontraktionen in einer Minute. Wir haben kürzlich eine Drosophila-Modell FUS / TLS-bezogenen amyotrophe Lateralsklerose, die Larven kriechen Defekt, verminderte Lebensdauer und Erwachsenen klettern Beeinträchtigung 6 zeigt entwickelt. Wir gezielte Expression of Wildtyp und mutierte Formen von FUS / TLS zu den Motoneuronen (OK-371-GAL4-Treiber) und führte eine Larven-Crawling-Assay. Wie unten dargestellt, Wildtyp-Larven bis zu 12 cm zu kriechen, während die Expression von Wildtyp-FUS verringerte die Larven kriechen Fähigkeit, etwa 6 cm. Tiere Ausdruck des ALS-Mutation R521C in FUS / TLS zeigen eine sehr starke Beeinträchtigung in ihrem kriechenden Bewegung (Abbildung 1), Crawling nur etwa 1 cm / min. Negativ Geotaxis RING-Assay Junge Wildtyp-erwachsenen Fliegen sollten einen mittleren Steighöhe von ~ 4-5 cm in einem 3-Sekunden-Zeit (die Zeit zwischen 3 Sekunden und kann eingestellt werden, um verschiedene Stämme oder Aktivität aufnehmen, um eine gegebene durchschnittliche Höhe für eine gegebene Definition Dehnung / Behandlung). Fliegen, die am Boden bleiben, wird ein Wert von 0 zugewiesen. Es ist nicht ratsam, mehr als 25 Fliegen pro Fläschchen zu verwenden, da wird es dann schwierig, die Lage jeder einzelnen Fliege an Höhe messen zu bestimmen. No Desensibilisierung hat bei eingerichtet, um die 6 aufeinander folgenden Versuchen Abstand von 1 Minute auseinander, dass wir beschäftigt beobachtet. Es ist wichtig, nicht die Prüfung Polystyrol Fläschchen in diesem Assay nach den ersten Sätzen von Daten verwertet werden gesammelt, weil neue Fliegen in Glasfläschchen verwendet nicht im gleichen Maße steigt, wie in frische Röhrchen. <img alt="1" src="/files/ftp_upload/3795/3795fig1.jpg" /> Abbildung 1. Setup für das RING-Assay. Die Digitalkamera wird ca. 1 m vor dem Gerät mit Fliegen in den Polystyrol-Röhrchen platziert; konzentrieren und zoomen Sie die Kamera auf das Gerät, und einen Timer einstellen, um 3,0 Sekunden. <img alt="2" src="/files/ftp_upload/3795/3795fig2.jpg" /> 2. Repräsentative Daten aus der Larven Crawling-Assay unter Verwendung Fliegen ektopisch UAS-FUS WT und UAS-FUS R521C unter der Kontrolle einer motoneuronalen Treiber (OK371-Gal4). <pclass = &quot;jove_content&quot;&gt;

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Wir haben nichts zu offenbaren.

Drosophila Verhalten wird streng von genetischen und umweltbedingten Faktoren reguliert. Wir und andere haben vorher beschriebenen Assays hier verwendet, um Daten zu sammeln, um Gene in Bezug auf Verhalten und zu menschlichen neurodegenerativen Erkrankungen in Drosophila 19.05 modelliert fliegen zu untersuchen. Für das Crawling-Assay ist eine sorgfältige Auswahl der 3 rd Larven ein entscheidender Schritt. Wenn die Behandlung mit einem Arzneimittel, dauert es 10-15 Minuten (oder ...

<table border="1"><tbody><tr><td> Name des Reagenzes </td><td> Firma </td><td> Katalog-Nummer </td></tr><tr><td> Saccharose </td><td> Fisher Scientific </td><td> S5-500 </td></tr><tr><td> Agarose </td><td> Invitrogen </td><td> 16500-500 </td></tr><tr><td> 6 Unzen Drosophila Flasche </td><td> Genesee Scientific </td><td> 32-130 </td></tr><tr><td> Pinsel (# 1) </td><td> Ted Pella, Inc. </td><td> 11859 </td></tr><tr><td> Fly Nahrungsbestandteile </td><td></td><td></td></tr><tr><td> Maismehl </td><td> Fisher Scientific </td><td> NC9109741 </td></tr><tr><td> Agar </td><td> Genesee Scientific </td><td> 66-104 </td></tr><tr><td> Melasse </td><td> Fisher Scientific </td><td> NC9349176 </td></tr><tr><td> Propionsäure </td><td> Acros </td><td> 14930-0010 </td></tr><tr><td> Tegosept </td><td> Spitze </td><td> 20-258 </td></tr><tr><td> Ethanol </td><td> FischerWissenschaftlich </td><td> BP2818-4 </td></tr><tr><td> Hefe </td><td> Genesee Scientific </td><td> 62-107 </td></tr></tbody></table>

methods to assay drosophila behavior

Drosophila melanogaster, der Fruchtfliege, wurde genutzt, um molekulare Mechanismen von einer breiten Palette von menschlichen Krankheiten wie Krebs, Herz-Kreislauferkrankungen und verschiedenen neurologischen Erkrankungen ein zu studieren. Wir haben einfache und robuste Verhaltensstörungen Assays zur Bestimmung Larven Fortbewegung, erwachsenen Steigfähigkeit (RING-Assay) und Werbeverhalten von Drosophila optimiert. Diese Verhaltens-Assays sind allgemein anwendbar für die Untersuchung der Rolle von genetischen und umweltbedingten Faktoren auf Fly Verhalten. Larven kriechen Fähigkeit zuverlässig für die Bestimmung frühzeitig Veränderungen in den kriechenden Fähigkeiten von Drosophila-Larven und auch für die Prüfung Wirkung von Medikamenten oder Erkrankungen des Menschen Genen (in transgenen Fliegen) auf ihrer Fortbewegung verwendet werden. Die Larven kriechen Assay wird immer anwendbar, wenn die Expression oder Abschaffung eines Gens verursacht Letalität im Puppenstadium oder adulte Stadien, da diese Fliegen nicht bis ins Erwachsenenalter überleben kann, wo sie sonst könnte beurteilt werden. Dieses BASICssay kann auch in Verbindung mit hellem Licht oder Stress zusätzliche Verhaltensreaktionen in Drosophila-Larven untersucht werden. Balzverhalten wurde weithin verwendet, um genetische Basis der sexuellen Verhaltens zu untersuchen, und kann auch verwendet werden, um die Aktivität und die Koordination zu untersuchen, ebenso wie Lernen und Gedächtnis werden. Drosophila Balzverhalten der Austausch von verschiedenen Sinnesreizen einschließlich visueller, auditiver und chemosensorischen Signale zwischen Männern und Frauen, die zu einer komplexen Reihe von gut charakterisierten Motor Verhaltensweisen die ihren Höhepunkt in erfolgreiche Kopulation führen. Traditionelle Erwachsenen klettern Assays (negative Geotaxis) sind langwierig, arbeitsintensiv und zeitaufwendig, mit signifikanten Unterschiede zwischen den einzelnen Studien 2-4. Die schnelle iterative negativen Geotaxis (RING)-Assay 5 hat viele Vorteile gegenüber größerem Umfang eingesetzten Protokolle und bietet eine reproduzierbare, sensibel und mit hohem Durchsatz Ansatz zur Erwachsenen-und Bewegungsapparat negativ Geotaxis b quantifizierenehaviors. In der RING-Assay kann mehrere Genotypen oder medikamentöse Behandlungen getestet gleichzeitig unter Verwendung der großen Anzahl von Tieren, mit der High-Throughput-Ansatz macht es zugänglicher für Screening-Experimenten.

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Methods to Assay Drosophila Behavior

Drosophila melanogaster ist eine genetisch und auf der Verhaltensebene praktische Zwecke geeigneten Modells, das verwendet wurde, um die molekularen und zellulären Grundlagen von vielen wichtigen biologischen Prozessen seit über einem Jahrhundert ein zu verstehen. Drosophila wurde auch genutzt, um Einblicke in die genetischen Grundlagen der Fliege Verhalten zu gewinnen.

Methoden zum Test Drosophila Verhalten

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular ...

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Neuroscience

Methods to Assay Drosophila Behavior

42.7K Aufrufe.  Louisiana State University Health Sciences Center. Drosophila melanogaster ist eine genetisch und auf der Verhaltensebene praktische Zwecke geeigneten Modells, das verwendet wurde, um die molekularen und zellulären Grundlagen von vielen wichtigen biologischen Prozessen seit über einem Jahrhundert ein zu verstehen. Drosophila wurde auch genutzt, um Einblicke in die genetischen Grundlagen der Fliege Verhalten zu gewinnen.

Serie: Methods to Assay Drosophila Behavior

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

Einzel Sensillum Recordings in der Insekten Drosophila melanogaster Und Anopheles gambiae</em

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

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Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the first born neurons remain closer to the ventricle while the last born neurons migrate past the first born neurons towards the surface of the brain1. In addition to neuronal migration2, a key process for normal cortical function is the regulation of neuronal morphogenesis3. While neuronal morphogenesis can be studied in vitro in primary cultures, there is much to be learned from how these processes are regulated in tissue environments.
We describe techniques to analyze neuronal migration and/or morphogenesis in organotypic slices of the cerebral cortex4,6. A pSilencer modified vector is used which contains both a U6 promoter that drives the double stranded hairpin RNA and a separate expression cassette that encodes GFP protein driven by a CMV promoter7-9. Our approach allows for the rapid assessment of defects in neurite outgrowth upon specific knockdown of candidate genes and has been successfully used in a screen for regulators of neurite outgrowth8. Because only a subset of cells will express the RNAi constructs, the organotypic slices allow for a mosaic analysis of the potential phenotypes. Moreover, because this analysis is done in a near approximation of the in vivo environment, it provides a low cost and rapid alternative to the generation of transgenic or knockout animals for genes of unknown cortical function. Finally, in comparison with in vivo electroporation technology, the success of ex vivo electroporation experiments is not dependant upon proficient surgery skill development and can be performed with a shorter training time and skill.

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

To conduct a rapid assessment of the function of genes in the development of cerebral cortex, we describe methods involving the ex vivo electroporation of plasmids co-expressing inhibitory RNA (RNAi) and GFP in murine embryonic cortex. This protocol is amenable to the study of various aspects of neurodevelopment such as neurogenesis, neuronal migration and neuronal morphogenesis including dendrite and axon outgrowth.

Methoden zur Untersuchung der neuronalen Morphogenese:<em&gt; Ex-vivo-</em&gt; RNAi Elektroporation in embryonale Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the ...

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.
The Drosophila larva is an established model system to study thermal nociception, a sensory response to potentially harmful temperatures that is evolutionarily conserved across species1,2. The advantages of Drosophila for such studies are the relative simplicity of its nervous system and the sophistication of the genetic techniques that can be used to dissect the molecular basis of the underlying biology4-6 In Drosophila, as in all metazoans, the response to noxious thermal stimuli generally involves a "nocifensive" aversive withdrawal to the presented stimulus7. Such stimuli are detected through free nerve endings or nociceptors and the amplitude of the organismal response depends on the number of nociceptors receiving the noxious stimulus8. In Drosophila, it is the class IV dendritic arborization sensory neurons that detect noxious thermal and mechanical stimuli9 in addition to their recently discovered role as photoreceptors10. These neurons, which have been very well studied at the developmental level, arborize over the barrier epidermal sheet and make contacts with nearly all epidermal cells11,12. The single axon of each class IV neuron projects into the ventral nerve cord of the central nervous system11 where they may connect to second-order neurons that project to the brain.
Under baseline conditions, nociceptive sensory neurons will not fire until a relatively high threshold is reached. The assays described here allow the investigator to quantify baseline behavioral responses or, presumably, the sensitization that ensues following tissue damage. Each assay provokes distinct but related locomotory behavioral responses to noxious thermal stimuli and permits the researcher to visualize and quantify various aspects of thermal nociception in Drosophila larvae. The assays can be applied to larvae of desired genotypes or to larvae raised under different environmental conditions that might impact nociception. Since thermal nociception is conserved across species, the findings gleaned from genetic dissection in Drosophila will likely inform our understanding of thermal nociception in other species, including vertebrates.

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.

Lokale und globale Methoden zur Beurteilung der thermischen Nozizeption in<em&gt; Drosophila</em&gt; Larven

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

Optogenetische Stimulation von Escape Verhalten in<em&gt; Drosophila melanogaster</em

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

A Single-fly Assay for Foraging Behavior in Drosophila

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster. In a light-tight box illuminated by red light that is invisible to fruit flies, a camera linked to custom data acquisition software monitors the position of six flies simultaneously. Each fly is confined to walk in individual arenas containing a food odor at the center. The testing arenas rest on a porous floor that functions to prevent odor accumulation. Latency to locate the odor source, a metric that reflects olfactory sensitivity under different physiological states, is determined by software analysis. Here, we discuss the critical mechanics of running this behavioral paradigm and cover specific issues regarding fly loading, odor contamination, assay temperature, data quality, and statistical analysis.

A Single-fly Assay for Foraging Behavior in Drosophila

In this video article, we describe an automated assay to measure the effect of hunger or satiety on olfactory dependent food search behavior in the adult fruit fly Drosophila melanogaster.

Ein Single-fly-Assay für Sammelverhalten in<em&gt; Drosophila</em

For many animals, hunger promotes changes in the olfactory system in a manner that facilitates the search for appropriate food sources. In this video ...

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy and cheap methods to distinguish between individuals in a trial. A standard technique for this is CO2 anaesthesia and then labelling or wing clipping each fly. However, this is invasive and has been shown to affect behavior. Other techniques have used coloration to identify flies. This article presents a simple and non-invasive method for labelling Drosophila that allows them to be individually identified within experiments, using food coloring. This method is used in trials where two males compete to mate with a female. Dyeing allowed quick and easy identification. There was, however, some difference in the strength of the coloration across the three species tested. Data is presented showing the dye has a lower impact on mating behavior than CO2 in Drosophila melanogaster. The impact of CO2 anaesthesia is shown to depend on the species of Drosophila, with D. pseudoobscura and D. subobscura showing no impact, whereas D. melanogaster males had reduced mating success. The dye method presented is applicable to a wide range of experimental designs.

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

This protocol describes a simple, cost effective way to individually identify Drosophila or other insects. Demonstration data investigating mating success across three species of Drosophila show that this method is comparable or better than the use of CO2 anaesthesia.

Färben Insekten für Verhaltenstests: die Paarungsverhalten von narkotisierten<em&gt; Drosophila</em

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Linear Agarose Kanäle zu studieren<em&gt; Drosophila</em&gt; Larval Krabbeln Verhalten

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

Ein einfacher Weg zu messen Veränderungen in Reward-seeking Verhalten verwenden<em&gt; Drosophila melanogaster</em

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...

Quantification of Drosophila Grooming Behavior

Drosophila grooming behavior is a complex multi-step locomotor program that requires coordinated movement of both forelegs and hindlegs. Here we present a grooming assay protocol and novel chamber design that is cost-efficient and scalable for either small or large-scale studies of Drosophila grooming. Flies are dusted all over their body with Brilliant Yellow dye and given time to remove the dye from their bodies within the chamber. Flies are then deposited in a set volume of ethanol to solubilize the dye. The relative spectral absorbance of dye-ethanol samples for groomed versus ungroomed animals are measured and recorded. The protocol yields quantitative data of dye accumulation for individual flies, which can be easily averaged and compared across samples. This allows experimental designs to easily evaluate grooming ability for mutant animal studies or circuit manipulations. This efficient procedure is both versatile and scalable. We show work-flow of the protocol and comparative data between WT animals and mutant animals for the Drosophila type I Dopamine Receptor (DopR).

This protocol describes a scalable individual grooming assay technique in Drosophila that yields robust, quantitative data to measure grooming behavior. The method is based on comparing the difference in dye accumulation on the bodies of un-groomed versus groomed animals over a set period of time.

Quantifizierung von Drosophila Grooming Behavior

Drosophila grooming behavior is a complex multi-step locomotor program that requires coordinated movement of both forelegs and hindlegs. Here we present a ...

High-throughput Analysis of Locomotor Behavior in the Drosophila Island Assay

Advances in next-generation sequencing technologies contribute to the identification of (candidate) disease genes for movement disorders and other neurological diseases at an increasing speed. However, little is known about the molecular mechanisms that underlie these disorders. The genetic, molecular, and behavioral toolbox of Drosophila melanogaster makes this model organism particularly useful to characterize new disease genes and mechanisms in a high-throughput manner. Nevertheless, high-throughput screens require efficient and reliable assays that, ideally, are cost-effective and allow for the automatized quantification of traits relevant to these disorders. The island assay is a cost-effective and easily set-up method to evaluate Drosophila locomotor behavior. In this assay, flies are thrown onto a platform from a fixed height. This induces an innate motor response that enables the flies to escape from the platform within seconds. At present, quantitative analyses of filmed island assays are done manually, which is a laborious undertaking, particularly when performing large screens.
This manuscript describes the &#34;Drosophila Island Assay&#34; and &#34;Island Assay Analysis&#34; algorithms for&#160;high-throughput, automated data processing and quantification of island assay data. In the setup, a simple webcam connected to a laptop collects an image series of the platform while the assay is performed. The &#34;Drosophila Island Assay&#34; algorithm developed for the open-source software Fiji processes these image series and quantifies, for each experimental condition, the number of flies on the platform over time. The &#34;Island Assay Analysis&#34; script, compatible with the free software R, was developed to automatically process the obtained data and to calculate whether treatments/genotypes are statistically different. This greatly improves the efficiency of the island assay and makes it a powerful readout for basic locomotion and flight behavior. It can thus be applied to large screens investigating fly locomotor ability, Drosophila models of movement disorders, and drug efficacy.

High-throughput Analysis of Locomotor Behavior in the Drosophila Island Assay

The island assay is a relatively new, cost-effective assay that can be used to evaluate the basic locomotor behavior of Drosophila melanogaster. This manuscript describes algorithms for automatic data processing and objective quantification of island assay data, making this assay a sensitive, high-throughput readout for large genetic or pharmacological screens.