SG wird durch Mittel aus der State University of New York at Buffalo und von John R. Oishei Foundation unterstützt.

1. Vorbereitung der Reagenzien <ol><li> Bereiten 1x Dissection Buffer mit 128 mM NaCl, 4mm CaCl 2, 4mm MgCl 2, 2 mM KCl, 5 mM HEPES und 36mm Sucrose. PH der Lösung auf 7,2 und Filter zu sterilisieren. </li><li> Bereiten Sie das Fixiermittel mit 1:1-Verdünnung von 16% Formaldehyd und 1x Dissection Buffer. </li><li> Bereiten 1x PBT mit 1x Phosphate Buffered Saline (PBS) bei einem pH von 7,2 und Triton X-100 in einer 1:100-Verhältnis. </li><li> Bereiten Sie 5% Rinderserumalbumin (BSA) mit 0,01% Na in 1x PBS-Azid. </li></ol> 2. Vorbereitung für die Präparation <ol><li> Sylgard Gerichte sind für die Präparation verwendet. Bereiten Sie die Gerichte, die von Gießen Sylgard 184 Silicon-Elastomer-Basis und Härter-Gemisch in eine Petrischale. Lassen Sie die Mischung vollständig zu festigen und vor der Verwendung trocken. </li><li> Vor der Dissektion, sammeln ein sauberes Paar Nr. 5 Pinzette, ein sauberes Paar gerader Klinge Vannas Schere und Stifte. </li></ol> 3. Dissection der 3. Larvenstadium <ol><li> Sammeln Sie wandern 3 rd Larvenstadium von einem bestimmten Genotyp auf eine Petrischale. </li><li> Waschen Sie die Larven in deionisiertem Wasser, um loszuwerden, das Essen. </li><li> Legen Sie eine Larve auf dem Sylgard Gericht, mit dem Rücken nach oben. Pin die Larve an seinem vorderen und hinteren Enden. </li><li> Geben Sie einen Tropfen 1x Dissection Buffer auf die merken Larve. Mit einer feinen Schere nip die Larve auf der dorsalen Mittellinie in der Nähe des hinteren Endes. Mit der Schere, schneiden Sie die Larvencuticula vom hinteren Ende bis zum vorderen Ende. </li><li> Geben Sie einen Tropfen neuer 1x Dissection Buffer auf den sezierten Larve. Mit einem Stift, wählen Sie einen seitlichen Rand der Kutikula der Nähe der Mitte der Larve und Pin die Nagelhaut. Mit anderen Pin, pin zweiten seitlichen Rand der Kutikula, wie in der Abbildung dargestellt. Zwei Stifte werden auf jeder seitlichen Seite verwendet. Siehe Abbildung unten. </li><li> Entfernen Sie vorsichtig den Darm, Fettkörper, und der Luftröhre, so dass nur das Gehirn mit den beiden optischen Lappen und ventralen Ganglien mit der segmentalen Nerven angebracht. Der Rest des Verfahrens beruht auf der Sylgard Schüssel getan. </li></ol> 4. Die Fixierung der Dissected Larva <ol><li> Inkubieren seziert Larve in fix für 30 Minuten, indem das Update alle 15 Minuten. </li><li> Nach der Fixierung, spülen Sie die Larve in 1x PBT für 30 Minuten mit dem Austausch der Puffer alle 10 Minuten. Es ist wichtig zu beachten, dass die Larve sollte immer in einem Puffer werden. </li></ol> 5. Antikörper-Färbung der Dissected Larva <ol><li> Bereiten Sie den primären Antikörper durch Verdünnen auf die geeignete Konzentration in 5% BSA mit Na-Azid in 1x PBT. Optimale Konzentrationen für verschiedene Antikörper unterscheiden. </li><li> Ersetzen Sie die letzte 1x PBT mit dem vorbereiteten primärer Antikörper-Lösung spülen, und inkubieren in einer feuchten Kammer bei 4 ° C über Nacht. Die feuchte Kammer wird durch das Aufbringen einer Plastikschale mit kim-Tücher in Wasser eingeweicht gebaut. </li><li> Am folgenden Tag, spülen Sie die seziert Larve in 1x PBT für 30 Minuten mit dem Austausch der Puffer alle 10 Minuten. </li><li> Bereiten Sie den sekundären Antikörper durch Verdünnen auf die geeignete Konzentration in 5% BSA mit Na-Azid in 1x PBT und inkubieren die Larve mit dem sekundären Antikörper-Lösung bei 25 ° C für eine Stunde. Wickeln Sie den feuchten Kammer in Folie, um Licht zu verhindern, als Sekundärantikörper lichtempfindlich sind. </li><li> Nach der Inkubation spülen die Larve in 1x PBT für 30 Minuten mit dem Austausch der Puffer alle 10 Minuten. </li></ol> 6. Montage und Visualisierung der Dissected Larva <ol><li> Vor der Montage der Larve auf dem Objektträger, bereiten Sie das Bild durch die Verbreitung zwei vertikale Linien von Nagellack in der Mitte, mit einem Raum genug für ein Deckglas zu sitzen (siehe Grafik). Lassen Sie den Nagellack trocknen. </li><li> Geben Sie einen Tropfen der Montage-Puffer in den Raum zwischen den vertikalen Nagellack Linien auf der Folie (siehe Diagramm). Die Folie ist nun bereit zur Montage. <img src="/files/ftp_upload/2128/2128.6.2.1.jpg" alt="Abbildung 1 Schritt 6.2" /> <img src="/files/ftp_upload/2128/2128.6.2.2.jpg" alt="Abbildung 2 Schritt 6.2" /></li><li> Nach dem letzten 1x PBT spülen, entfernen Sie vorsichtig die seitlichen Zapfen. Achten Sie darauf, nicht reißen die Nagelhaut. </li><li> Entfernen Sie vorsichtig die beiden verbleibenden Pins und holen die Larve mit einer Pinzette und Ort der Larve horizontal auf die vorbereitete Folie. Vergewissern Sie sich die Larve nicht umgedreht wird und orientiert sich Bauchseite auf. </li><li> Gently Ort ein Deckglas auf und die Ränder mit Nagellack (siehe Diagramm). Die Larve ist nun bereit für die Visualisierung unter einem Fluoreszenzmikroskop. <img src="/files/ftp_upload/2128/2128.6.5.jpg" alt="Abbildung 1 Schritt 6.5" /></li></ol> 7. Repräsentative Ergebnisse <img src="/files/ftp_upload/2128/2128fig1.jpg" alt="Abbildung 1" /> Abbildung 1: Ein repräsentatives Bild der Larven segmentalen Nerven von einem Wildtyp-Larveine using Antikörper gegen den synaptischen Vesikel Protein Cystein-String-Protein (CSP, Zinsmaier et al. 1994). Beachten Sie, dass die segmentalen Nerven reibungslos gefärbt sind. Bar = 20 um <img src="/files/ftp_upload/2128/2128fig2.jpg" alt="Abbildung 2" /> Abbildung 2: Ein repräsentatives Bild der Larven segmentalen Nerven von einem Motor-Protein-Mutanten Larve mit Antikörpern gegen den synaptischen Vesikel Protein Cystein-String-Protein (CSP). Beachten Sie, dass die segmentalen Nerven massiven Ansammlungen, die hell für CSP (Pfeile) Flecken zeigen.

<ol>
	<li>Gunawardena, S., Goldstein, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+Axonal+Transport+and+Neuronal+Viability+by+Amyoid+Precursor+Protein+Mutations+in+Drosophila.">Disruption of Axonal Transport and Neuronal Viability by Amyoid Precursor Protein Mutations in Drosophila.</a> Neuron. 32 (2), 389-401 (2001).</li><li>Gunawardena, S., Her, L. S., Brusch, R. G., Laymon, R. A., Niesman, I. R., Gordesky-Gold, B., Sintasath, L., Bonini, N. M., Goldstein, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+axonal+transport+by+loss+of+huntingtin+or+expression+of+pathogenic+polyQ+proteins+in+Drosophila.">Disruption of axonal transport by loss of huntingtin or expression of pathogenic polyQ proteins in Drosophila.</a> Neuron. 40 (1), 1-2 (2003).</li><li>Hurd, D. D., Saxton, W. 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+mutations+cause+motor+neuron+disease+phenotypes+by+disrupting+fast+axonal+transport+in+Drosophila.">Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila.</a> Genetics. 144 (3), 1075-1085 (1996).</li><li>Zinsmaier, K. E., Eberle, K. K., Buchner, E., Walter, N., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paralysis+and+early+death+in+cysteine+string+protein+mutants+of+Drosophila.">Paralysis and early death in cysteine string protein mutants of Drosophila.</a> Science. 263, 977-980 (1994).</li></ol>

Die Drosophila Larve segmentalen Nerven sind ein leistungsfähiges System, Mechanismen in axonalen Transport zu studieren. Wenn die 3 rd instar Larven Dissektion korrekt durchgeführt wird, können alle Komponenten des ZNS, PNS, und die damit verbundenen Gewebe, wie die Muskeln und NMJs, innerhalb einer einzigen Larve untersucht werden. Wir haben dieses Protokoll von der von Hurd und Saxton (1996) veröffentlicht angepasst. Dieses Protokoll kann auch angepasst werden andere neuronale Antikörper-Test...

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visualization of larval segmental nerves in 3rd instar drosophila larval preparations

Drosophila melanogaster ist ein leistungsfähiges Modell für die Untersuchung der Entwicklung und Funktion des Nervensystems, vor allem wegen seiner günstigen Genetik und vollständig sequenzierten Genoms entstehen. Darüber hinaus ist die Larve des Nervensystems ein ideales Modellsystem, um Mechanismen der axonalen Transport Studie als Larven segmentalen Nerven Bündel von Axonen mit ihrem Zellkörper im Gehirn und die Nerven-Terminals endet entlang der Länge des Körpers befindet enthalten. Hier beschreiben wir das Verfahren zur Visualisierung der synaptischen Vesikel Proteine ​​innerhalb Larven segmentalen Nerven. Wenn es richtig gemacht, alle Komponenten des Nervensystems, zusammen mit der dazugehörigen Gewebe wie Muskeln und NMJs, bleiben intakt und unbeschädigt, und bereit, visualisiert werden. 3 rd Larvenstadium, die verschiedene Mutationen seziert werden, fest, inkubiert mit synaptischen Vesikel Antikörper, visualisiert und im Vergleich zum Wildtyp-Larven. Dieser Vorgang kann für mehrere verschiedene synaptischen oder neuronalen Antikörpern und Veränderungen in der Verteilung einer Vielzahl von Proteinen angepasst werden können innerhalb von Larven segmentalen Nerven beobachtet werden.

Watch this Scientific Journal Video about Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations at JoVE.com

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster-Larven bieten ein ideales Modellsystem, um die Mechanismen der axonalen Transport innerhalb Larven segmentalen Nerven zu untersuchen. Mit diesem Verfahren können 3 rd Larvenstadium, die verschiedene Mutationen zu Wildtyp-Larven verglichen werden.

Visualisierung von Larven segmentalen Nerven in 3 Rd Instar Drosophila Larven Vorbereitungen

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

visualization-larval-segmental-nerves-3rd-instar-drosophila-larval

Research

JoVE Journal

Neuroscience

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

10.2K Aufrufe.  SUNY-University at Buffalo. Drosophila melanogaster-Larven bieten ein ideales Modellsystem, um die Mechanismen der axonalen Transport innerhalb Larven segmentalen Nerven zu untersuchen. Mit diesem Verfahren können 3 rd Larvenstadium, die verschiedene Mutationen zu Wildtyp-Larven verglichen werden.

Serie: Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

In-vivo-Visualisierung von synaptischen Vesikeln Innerhalb Drosophila Larven Segmentale Axone

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Forschung

Neurowissenschaften

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

Mapping und Anwendung von Enhancer-trap Flippase Expression in Larven und Erwachsene Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

Dissektion und Immunhistochemie von Larven, Puppen und Erwachsenenbildung<em&gt; Drosophila</em&gt; Netzhäute

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Vorderhirn Elektrophysiologische Aufzeichnung im Larven Zebrafisch

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

Funktionelle Analyse der Larvenfütterung Circuit in Drosophila</i

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Live-Imaging von<em&gt; Drosophila</em&gt; Larven Neuroblasten

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Warum Quantifizierung von Bedeutung: Charakterisierung von Phänotypen bei der<em&gt; Drosophila</em&gt; Larval Neuromuskuläre Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

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 ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

Einzellige Auflösung Fluoreszenz Imaging von Drosophila zirkadiane Uhren in Larven Gehirn Kultur Leben

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.