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1. Preparazione dei reagenti <ol><li> Preparare 1x buffer Dissection a 128 mM NaCl, CaCl 2 4mm, 4 mm MgCl 2, 2 mM KCl, HEPES 5 mm e 36 mm saccarosio. Il pH della soluzione a 7,2 e filtro sterilizzare. </li><li> Preparare il fissativo con diluizione 1:1 del 16% di formaldeide e Buffer Dissection 1x. </li><li> Preparare 1x PBT utilizzando 1x soluzione tampone fosfato (PBS) a pH di 7,2 e Triton X-100 in un rapporto 1:100. </li><li> Preparare il 5% di albumina sierica bovina (BSA) con 0,01% Na azide in 1x PBS. </li></ol> 2. Preparazione per la dissezione <ol><li> Sylgard piatti vengono utilizzati per la dissezione. Preparare i piatti versando 184 Sylgard base di elastomeri di silicone e curare miscela agente in una capsula di Petri. Lasciare il composto a solidificare completamente asciutti prima dell&#39;uso. </li><li> Prima della dissezione, raccogliere un paio pulito # 5 pinze, un paio di forbici pulito lama dritta Vännäs, e perni. </li></ol> 3. Dissezione di 3 Larve ° Instar <ol><li> Raccogliere vagare 3 ° larve instar di un genotipo specifico su una piastra di Petri. </li><li> Lavare le larve in acqua deionizzata per sbarazzarsi del cibo. </li><li> Inserire una larva sul piatto sylgard, con l&#39;alto lato dorsale. Perno la larva alle sue estremità anteriori e posteriori. </li><li> Mettere una goccia di tampone Dissection 1x sulla larva appuntato. Utilizzando bene nip forbici la larva sulla linea mediana dorsale verso la fine posteriore. Utilizzando le forbici, tagliare la cuticola larvale dalla fine posteriore fino alla fine anteriore. </li><li> Mettere una goccia di nuovo tampone Dissection 1x sul larva sezionato. Utilizzando uno spillo, scegliere un bordo laterale della cuticola vicino al centro della larva e perno la cuticola. Usando un altro pin, pin il secondo bordo laterale della cuticola, come mostrato in figura. Due pin sono usati su entrambi i lati laterali. Vedi figura sotto. </li><li> Rimuovere con attenzione l&#39;intestino, corpi grassi, e la trachea, lasciando solo il cervello con i due lobi ottici e gangli ventrali con i nervi segmentali allegato. Il resto della procedura è fatto sul piatto sylgard. </li></ol> 4. Fissazione della larva Dissected <ol><li> Incubare la larva sezionato in correzione per 30 minuti, sostituendo la correzione ogni 15 minuti. </li><li> Dopo la fissazione, lavare la larva in 1x PBT per 30 minuti, sostituendo il buffer ogni 10 minuti. E &#39;importante notare che la larva deve essere sempre in un buffer. </li></ol> 5. La colorazione degli anticorpi della Larva Dissected <ol><li> Preparare l&#39;anticorpo primario diluendo alla concentrazione adeguata nel 5% BSA con Na azide in 1x PBT. Concentrazioni ottimali variano per gli anticorpi diversi. </li><li> Sostituire l&#39;ultimo 1x PBT sciacquare con la soluzione preparata anticorpo primario e incubare in camera umida a 4 ° C durante la notte. La camera umida è costruito da rivestimento di un piatto di plastica con kim-salviette imbevute di acqua. </li><li> Il giorno seguente, lavare la larva sezionato in 1x PBT per 30 minuti, sostituendo il buffer ogni 10 minuti. </li><li> Preparare l&#39;anticorpo secondario diluito alla concentrazione appropriata nel 5% BSA con Na azide in 1x PBT, e incubare la larva con la soluzione di anticorpo secondario a 25 ° C per un&#39;ora. Avvolgere la camera umida in un foglio per impedire alla luce, come gli anticorpi secondari sono sensibili alla luce. </li><li> Dopo l&#39;incubazione, lavare la larva in 1x PBT per 30 minuti, sostituendo il buffer ogni 10 minuti. </li></ol> 6. Montaggio e visualizzazione del Larva Dissected <ol><li> Prima di montare la larva sulla diapositiva, preparare la slitta diffondendo due linee verticali di smalto nel mezzo, con uno spazio sufficiente per una copertura antiscivolo per sedersi (vedi schema). Lasciare asciugare completamente smalto. </li><li> Mettere una goccia di tampone di montaggio nello spazio tra le linee verticali smalto per unghie sulla diapositiva (vedi schema). Il vetrino è pronto per il montaggio. <img src="/files/ftp_upload/2128/2128.6.2.1.jpg" alt="Figura 1 passo 6,2" /> <img src="/files/ftp_upload/2128/2128.6.2.2.jpg" alt="Figura 2 passo 6,2" /></li><li> Dopo l&#39;ultimo risciacquo 1x PBT, rimuovere delicatamente i perni laterali. Fare attenzione a non strappare la cuticola. </li><li> Rimuovere con attenzione i due perni rimanenti e raccogliete la larva con una pinza e la larva posto orizzontalmente sul vetrino preparato. Assicurarsi che la larva non è capovolto ed è orientato a lato ventrale. </li><li> Delicatamente luogo uno slittamento di copertura sulla parte superiore e sigillare i bordi con smalto per unghie (vedi schema). La larva è ora pronto per la visualizzazione al microscopio a fluorescenza. <img src="/files/ftp_upload/2128/2128.6.5.jpg" alt="Figura 1 passo 6,5" /></li></ol> 7. Rappresentante Risultati <img src="/files/ftp_upload/2128/2128fig1.jpg" alt="Figura 1" /> Figura 1: Una immagine rappresentativa del larvale nervi segmentali da un tipo selvaggio larvA utilizzando anticorpi contro la proteina proteina sinaptica stringa vescicola cisteina (CSP, Zinsmaier et al. 1994). Si noti che i nervi segmentali sono uniformemente colorati. Bar = 20μm <img src="/files/ftp_upload/2128/2128fig2.jpg" alt="Figura 2" /> Figura 2: Una immagine rappresentativa del larvale nervi segmentale da una larva proteina mutante motore utilizzando anticorpi contro la proteina proteina sinaptica stringa vescicola cisteina (CSP). Si noti che i nervi segmentali mostrano accumuli di massa che macchia brillante per il CSP (frecce).

<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>

Nessun conflitto di interessi dichiarati.

I nervi larvale Drosophila segmentale sono un potente sistema per lo studio dei meccanismi di trasporto assonale. Se il 3 ° instar dissezione larvale viene eseguita correttamente, tutte le componenti del sistema nervoso centrale, PNS, e tessuti associati, come i muscoli e NMJs, possono essere esaminate all&#39;interno di una singola larva. Abbiamo adattato questo protocollo da quella pubblicata da Hurd e Saxton (1996). Questo protocollo può essere adattato anche a test altri anticorpi neuronali, ma...

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

Drosophila melanogaster sta emergendo come un potente sistema modello per studiare lo sviluppo e la funzione del sistema nervoso, in particolare a causa della sua genetica e conveniente genoma completamente sequenziato. Inoltre, il sistema nervoso larvale è un sistema modello ideale per studiare i meccanismi del trasporto assonale, come i nervi larvale segmentale contenere fasci di assoni con i loro corpi cellulari situato all&#39;interno del cervello e le loro terminazioni nervose finale lungo la lunghezza del corpo. Qui si descrive la procedura per la visualizzazione di proteine ​​delle vescicole sinaptiche all&#39;interno larvale nervi segmentale. Se fatto correttamente, tutte le componenti del sistema nervoso, con relativi tessuti come i muscoli e NMJs, rimane intatto, integro e pronto per essere visualizzato. 3 ° larve instar portatori di mutazioni diverse sono sezionati, fisso, incubate con anticorpi delle vescicole sinaptiche, visualizzati e confrontati con le larve wild-type. Questa procedura può essere adattata per diversi anticorpi diversi sinaptica o neuronale e cambiamenti nella distribuzione di una varietà di proteine ​​può essere facilmente osservata entro larvale nervi segmentale.

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

Larve di Drosophila melanogaster fornire un sistema modello ideale per studiare i meccanismi di trasporto assonale in nervi larvale segmentale. Utilizzando questa procedura, 3 ° larve instar portatori di mutazioni varie può essere paragonato a larve wild-type.

Visualizzazione dei nervi larvali segmentale in 3 Rd Instar Drosophila I preparativi larvali

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

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Research

JoVE Journal

Neuroscience

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

10.2K Views.  SUNY-University at Buffalo. Larve di Drosophila melanogaster fornire un sistema modello ideale per studiare i meccanismi di trasporto assonale in nervi larvale segmentale. Utilizzando questa procedura, 3 ° larve instar portatori di mutazioni varie può essere paragonato a larve wild-type.

Video: 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.

Nella visualizzazione in vivo delle vescicole sinaptiche In Drosophila assoni larvali segmentale

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

Ricerca

Neuroscienze

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.

Mappatura e applicazione di Enhancer-trappola Espressione Flippase in larvali e adulti Drosophila SNC

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.

Dissezione e immunoistochimica di larvale, pupa e adulto<em&gt; Drosophila</em&gt; Retine

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.

Proencefalo registrazione elettrofisiologica in larvale 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 ...

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.

Analisi funzionale del circuito di alimentazione larvale 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.

Immagini dal vivo di<em&gt; Drosophila</em&gt; larvali neuroblasti

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.

Perché Quantificazione Matters: Caratterizzazione dei fenotipi al<em&gt; Drosophila</em&gt; Larvale Giunzione neuromuscolare

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.

Utilizzando canali lineari Agarose per studiare<em&gt; Drosophila</em&gt; Comportamento larvale Crawling

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.

Cella singola risoluzione fluorescenza Live Imaging di Drosophila orologi circadiani nel cervello larvale cultura

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.