We thank members of the Kamiyama Lab for comments on the manuscript. This work was supported by an NIH R01 NS107558 (to M.I., K.B., and D.K.).

1. Equipment and Supplies
<ol>
	<li>Materials for collecting embryos and training adults to lay eggs
	<ol>
		<li>Prepare the filtration apparatus by severing a 50 mL tube and cutting open a hole in the cap to set a mesh filter with pores of 100 &#181;m (Table of Materials) in between the tube and the cap. 
		NOTE: Alternatively, cell strainers with pores of 100 &#181;m (Table of Materials) can be used for the filtration step of embryo collection.</li>
		<li>Make agar plates with grape agar premix (Table of Materials) according to the listed instructions. Briefly, gently stir 1 packet of the powder mix into 500 mL of room temperature (RT, 23 &#176;C) dH2O and microwave the dissolved mixture to vigorous boil. After cooling down to 70&#8722;75 &#176;C, pour the mixture into Petri dishes (60 mm). After the agar is solidified, store plates at 4 &#176;C.</li>
		<li>Prepare yeast paste by mixing active dry yeast (Table of Materials) and water to a paste consistency, and keep at 4 &#176;C.</li>
		<li>Use egg-collection cages (for 60 mm Petri dish, Table of Materials) that provide sufficient air flow.</li>
	</ol>
	</li>
	<li>Preparation of dissection needles and dye injection micropipettes
	<ol>
		<li>Prepare dye injection micropipette and dissection needle from the same capillary tubing with an inner diameter of 0.6 mm and an outer diameter of 1.2 mm (Table of Materials). Pull the capillary tubing by a micropipette puller at 7% from 170 V maximum output (Table of Materials) to create a sharp needle with a taper of ~0.4 cm in length.</li>
		<li>For dye injection, adjust the micropipette with a micropipette beveler (Table of Materials) by a bubble beveling technique described in instrument&#39;s manual.
		<ol>
			<li>In short, soak the grinder with a wetting agent (Table of Materials) to prevent the water from &#39;dragging&#39; the needle tip. Place the needle on the micropipette clamp at 25&#8722;30&#176; and lower the tip onto two-thirds of the radius out from the center of the beveling surface. Grind the needle while a syringe with tubing pushes air into the needle, to ensure that the micropipette will be clear of glass shavings.</li>
			<li>Mark the micropipette with a fine-tip permanent marker to indicate the position of the opening at the tip after beveling as it is challenging to locate the narrow opening of the micropipette that is formed at an angle.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Preparation for Embryo Collection
<ol>
	<li>Ensure that the adult flies (20&#8722;40 wild-type Canton-S or white flies), males and females, are maintained in young (&#60;7 days) and healthy conditions for the ideal egg collection. 
	NOTE: To stimulate egg-laying, flies are trained in their egg collection cage a couple of days prior to egg collection on agar plates streaked with yeast paste at least once every day.</li>
</ol>
3. Embryo Staging
<ol>
	<li>Allow the flies to lay eggs overnight (or at least 15 h) at RT to collect the embryos at 15 h AEL, i.e., stage 1618, to view dendritogenesis of the aCC and RP3 motor neurons. In the morning, collect the plate with the eggs. 
	NOTE: The embryos at 15 h AEL will have a distinct 4-chamber gut18. For imaging different stages follow their specific morphological criteria and aging conditions.</li>
	<li>To collect the embryos, dechorionate the eggs laid on the plate with 50% bleach for 5 min.</li>
	<li>Once the chorions have cleared, pour the contents of the plate through the filtration apparatus or cell strainer to isolate the embryos. Using a squeeze bottle of water, dilute the bleach left on the plate and gather as many embryos as possible by decanting the mixture into the filter.</li>
	<li>Wash the embryos on the filter 3&#8722;4x with more water or until the bleach odor dissipates. Remove the filter from the apparatus and wash the embryos onto another clean plate with water. Decant the water from the new plate that the embryos are on.</li>
	<li>Prepare a glass slide by covering it with two layers of vinyl tape in the center, forming a rectangle. Cut a rectangular pool out of the tape using a razor blade. Place a thin strip of double-sided tape towards the upper end of the pool, this is where the embryos will be placed as shown in Figure 1.</li>
	<li>Using fine forceps, individually select 5&#8722;10 embryos at 15 h AEL and place them on the double-sided tape with the dorsal side facing up. Add insect Ringer&#39;s saline19 to the dissection pool to protect the embryos from desiccation (Figure 1).</li>
</ol>
4. Dissection and Staining
<ol>
	<li>Using a glass needle under a dissecting microscope (Table of Materials), cut through the midline of a single embryo at its surface from its posterior to its anterior end. Then drag the embryo out from the vitelline membrane from the tape onto the glass (boxed in Figure 1). Take care not to damage the interior tissues of the embryo.</li>
	<li>Flip the epithelial tissues from the center and attach the epidermal edge onto the surface of the glass slide (Figure 1, inset).</li>
	<li>Using a tube-connected needle with a tip opening of ~300 &#181;m (prepared by breaking the thin tip of a dissection needle), aspirate or blow air to detach and remove the dorsal longitudinal tracheal trunks as well as any remaining guts.</li>
	<li>Use 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) to fix the embryos for 5 min at RT. Wash the embryos 3x with PBS.</li>
	<li>Stain the embryos with 1 &#181;L of anti-horseradish peroxidase antibody conjugated with cyanine 3 dye (anti-HRP Cy3) (Table of Materials) in 200 &#181;L of PBS for 1 h. Wash the embryos with PBS 3x after staining. 
	NOTE: The dye of anti-HRP can be changed based on the lipophilic dyes of choice for injection.</li>
</ol>
5. Filling of the Injection Micro-pipette
<ol>
	<li>Heat lipophilic dyes (5 mg/mL of DiO or DiD, Table of Materials) to 60 &#176;C in a 1:10 mixture of ethanol:vegetable oil before use.</li>
	<li>Prepare an oil-dissolved dye slide for the injection micropipette. Place the micropipette into the capillary holder (Figure 2, #1). Using the micromanipulator (Table of Materials), adjust the micropipette to be over the dye slide. Then, adjust the stage to place the micropipette onto the dye (Figure 2, #2).</li>
	<li>To fill up the micropipette, use a microinjector (Table of Materials) (Figure 2, #3). Collect the dye in the micropipette by setting the Pi (injection pressure) between 200&#8722;500 hPa (hectopascal), the Ti (injection time) between 0.1&#8722;0.5 s and Pc (compensation pressure) to 0 hPa for 5 min (Figure 2, #4).</li>
	<li>Once the dye has been collected, remove the dye slide and place the sample onto the microscope stage. Next, increase the Pc to a range of 20&#8722;60 hPa before lowering the micropipette into the sample to prevent contamination of PBS by capillary action.</li>
</ol>
6. Dye Injection into Neurons
<ol>
	<li>Locate the embryo in the center using the microscope with 10x objective lens (Table of Materials) and align the micropipette with the embryo. 
	NOTE: The size of the dye droplet can be adjusted by changing the Pi or the size of the opening of the micropipette tip. The droplet should be 10&#8722;20 &#181;m, which is approximately the width of 1 muscle.</li>
	<li>Change the objective lens to a water-immersion 40x lens (Table of Materials) and submerge the lens into PBS to see the embryo.
	<ol>
		<li>Use fluorescence microscopy to check the neuronal morphology marked by anti-HRP Cy3 and determine the injection site.</li>
		<li>During injection, use brightfield microscopy to see the dye droplet. When the embryo is in focus, change the position of the micropipette to make gentle contact with the tip of the axon of interest (e.g., aCC, RP3).</li>
		<li>Drop the dye in a right abdominal (A2&#8722;A6) hemi-segment at the neuromuscular junction of aCC or RP3 (Figure 3) with either DiD or DiO, by using the neurons marked by anti-HRP Cy3. Using the hand control (mouse; Figure 2, 5) release the dye and remove the micropipette after dropping the dye with the micromanipulator and move onto the next injection site. 
		NOTE: Unlike other dyes (e.g., Lucifer yellow, calcein) which spread into neighboring cells through gap junctions, lipophilic dyes associate with cell membranes and do not transfer to neighbors. Due to the relatively large size of the dye droplet, however, this technique also results in labeling of the partnering muscles (Figure 3A).</li>
	</ol>
	</li>
	<li>Incubate the sample at RT for 1 h after dye-drop before imaging. 
	NOTE: The protocol can be paused here before mounting, and the sample can be kept at 4 &#176;C overnight. Lipophilic dyes can also be delivered using iontophoresis, if an intracellular direct-coupled (DC) amplifier is readily available20.</li>
</ol>
7. Imaging with a Confocal Microscope
<ol>
	<li>Remove the double-sided tape and vinyl tape from the glass slide with the help of forceps.</li>
	<li>Prepare a cover slip (22 x 22 mm2 No.1 cover glass) with a small amount of vacuum grease (Table of Materials) at the four corners and carefully place on the sample, avoiding air bubbles. Remove any excess PBS using task wipers.</li>
	<li>Push down the cover slip to adjust the working distance between the objective lens and the sample. Completely seal the edges of the cover slip with nail polish.</li>
	<li>Image at 10x and 100x magnification using a confocal microscope.</li>
	<li>Use ImageJ software for processing raw images from the microscope (Table of Materials). 
	NOTE: Observation must begin within 10 min after mounting for the best images. Otherwise, at RT, the dye will spread to sites adjacent to the injection site creating unwanted background for imaging. To slow down the diffusion of dye, the sample can be stored at 4 &#176;C for a couple of hours.</li>
</ol>

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A., Hartenstein, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The embryonic development of Drosophila melanogaster. , (1985).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Ringer's+solution.">Drosophila Ringer's solution.</a> Cold Spring Harbor Protocols. 2007 (4), (2007).</li><li>Rickert, C., Kunz, T., Harris, K. -. L., Whitington, P., Technau, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Labeling+of+single+cells+in+the+central+nervous+system+of+Drosophila+melanogaster.">Labeling of single cells in the central nervous system of Drosophila melanogaster.</a> Journal of Visualized Experiments. 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The authors have nothing to disclose.

The use of dye labeling for studying neuronal morphology has several advantages over genetic cell-labeling techniques. The dye labeling technique can minimize the amount of time needed for labeling and imaging the morphologies of motor neurons. The dye labeling process is quite fast as it takes less than 2 h and enables us to define the outline of neuronal projections. As an alternative, one can visualize the aCC motor neuron by choosing a GAL4 line that expresses the yeast GAL4 transcription factor in aCC, and crossing ...

The embryonic motor neuron system of Drosophila offers a powerful experimental model to analyze the mechanisms underlying the development of the central nervous system (CNS)1,2,3. The motor neuron system is amenable to biochemical, genetic, imaging, and electrophysiological techniques. Using the techniques, genetic manipulations and functional analyses can be carried out at the level of single motor neurons2,4,5,6.
During early development of the nervous system, neuroblasts divide and generate a large number of glia and neurons. The spatiotemporal relationship between the delamination and the gene expression profile of neuroblasts has been previously investigated in detail7,8,9. In the case of the motor neuron system, the formation of embryonic neuromuscular junction (NMJ) has been extensively studied using the aCC (anterior corner cell), RP2 (raw prawn 2), and RP5 motor neurons2,10. For instance, when the RP5 motor neuron forms a nascent synaptic junction, the pre-synaptic and post-synaptic filopodia are intermingled11,12,13. Such direct cellular communication is vital to initiate the NMJ formation. Contrary to what we know about the peripheral nerve branches, our knowledge of how motor dendrites initiate synaptic connectivity within the CNS is still primitive.
In this report, we present a technique that allows retrograde labeling of motor neurons in embryos by means of micropipette-mediated delivery of lipophilic dyes. This technique enables us to trace the 38 motor neurons innervating each of the 30 body wall muscles in a hemi-segment at 15 h after egg laying (AEL)14. By using this technique, our group has thoroughly investigated numerous gain-of-function/loss-of-function alleles15,16,17. We have recently unraveled the molecular mechanisms that drive initiation of motor dendrite connectivity and demonstrated that a Dscam1-Dock-Pak interaction defines the site of dendrite outgrowth in the aCC motor neuron17. In general, this technique is adaptable for the phenotypic analysis of any embryonic motor neurons in wild type or mutant strains, enhancing our ability to provide new insights into the functional design of the Drosophila nervous system.

<table><tbody><tr><td>10x objective lens</td><td>Nikon</td><td></td><td>Plan</td></tr><tr><td>40x water-immersion lens</td><td>Nikon</td><td></td><td>NIR Apo</td></tr><tr><td>Capillary tubing</td><td>Frederick Haer&amp;amp;Co</td><td>27-31-1</td><td></td></tr><tr><td>Confocal microscope</td><td>Andor</td><td>N/A</td><td>Dragonfly Spinning disk confocal unit</td></tr><tr><td>Cover glass</td><td>Corning</td><td></td><td>22x22 mm Square #1</td></tr><tr><td>DiD</td><td>ThermoFisher</td><td>V22886</td><td></td></tr><tr><td>DiI</td><td>ThermoFisher</td><td>V22888</td><td></td></tr><tr><td>DiO</td><td>ThermoFisher</td><td>V22887</td><td></td></tr><tr><td>Dissecting microscope</td><td>Nikon</td><td>N/A</td><td>SMZ-U</td></tr><tr><td>Double Sided Tape</td><td>Scotch</td><td>665</td><td></td></tr><tr><td>Dow Corning High-Vacuum Grease</td><td>Fisher Sci.</td><td>14-635-5D</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11252-20</td><td></td></tr><tr><td>Egg collection cage</td><td>FlyStuff</td><td>59-100</td><td></td></tr><tr><td>FemtoJet 5247</td><td>Eppendorf</td><td>discontinued</td><td>FemtoJet 4i (Cat No. 5252000021)</td></tr><tr><td>ImageJ</td><td>NIH</td><td></td><td>Image processing software</td></tr><tr><td>Micromanipulator</td><td>Sutter</td><td>MP-225</td><td></td></tr><tr><td>Micropipette beveler</td><td>Sutter</td><td>BV-10-B</td><td></td></tr><tr><td>Needle puller</td><td>Narishige</td><td>PC-100</td><td></td></tr><tr><td>Nutri-Fly Grape Agar Powder Premix Packets</td><td>FlyStuff</td><td>47-102</td><td></td></tr><tr><td>Nylon Net Filter</td><td>Millipore</td><td></td><td></td></tr><tr><td>Paraformaldehyde 16% Solution, EM grade</td><td>Electron Microscopy Sciences</td><td>15710</td><td>Any EM grades</td></tr><tr><td>PBS</td><td>Roche</td><td>11666789001</td><td>Sold on sigmaaldrich, boxed 10x solution</td></tr><tr><td>Photo-Flo 200</td><td>Kodak</td><td>146 4510</td><td>Wetting agent</td></tr><tr><td>Upright fluorescence microscope</td><td>Nikon</td><td>N/A</td><td>Eclipse Ci with a LED light source</td></tr><tr><td>Vinyl Electrical Tape</td><td>Scotch</td><td>6143</td><td></td></tr><tr><td>VWR Cell Strainers</td><td>VWR</td><td>10199-659</td><td></td></tr><tr><td>Yeast</td><td>FlyStuff</td><td>62-103</td><td>Active dry yeast (RED STAR)</td></tr></tbody></table>

retrograde tracing of drosophila embryonic motor neurons using lipophilic fluorescent dyes

We describe a technique for retrograde labeling of motor neurons in Drosophila. We use an oil-dissolved lipophilic dye and deliver a small droplet to an embryonic fillet preparation by a microinjector. Each motor neuron whose membrane is contacted by the droplet can then be rapidly labeled. Individual motor neurons are continuously labeled, enabling fine structural details to be clearly visualized. Given that lipophilic dyes come in various colors, the technique also provides a means to get adjacent neurons labeled in multicolor. This tracing technique is therefore useful for studying neuronal morphogenesis and synaptic connectivity in the motor neuron system of Drosophila.

A representative image of the aCC and RP3 motor neurons is shown in Figure 3C to demonstrate the multicolor labeling of motor neurons at 15 h AEL. Their dendritic morphologies are largely invariant between embryos. The staining pattern obtained with anti-HRP antibody is shown in gray. A small droplet of DiO or DiD was deposited on the NMJ of muscle 1 or 6/7, respectively. Figure 4 demonstrates the capability to quantitatively measure the phenoty...

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Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

We describe a method for retrograde tracing of the Drosophila embryonic motor neurons using lipophilic fluorescent dyes.

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

We describe a technique for retrograde labeling of motor neurons in Drosophila. We use an oil-dissolved lipophilic dye and deliver a small droplet to an ...

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6.1K Views.  University of Georgia. We describe a method for retrograde tracing of the Drosophila embryonic motor neurons using lipophilic fluorescent dyes.

Video: Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

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.

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

Neuroscience

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Biology

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

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

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila larvae, and fish, which all engage in undulatory movements (like crawling or swimming) as their primary mode of locomotion. However, a more sophisticated understanding of the individual motor neuron (MN) specification&#8212;at least in terms of informing disease therapies&#8212;demands an equally tractable system that better models the complex appendage-based locomotion schemes of vertebrates. The adult Drosophila locomotor system in charge of walking meets all of these criteria with ease, since in this model it is possible to study the specification of a small number of easily distinguished leg MNs (approximately 50 MNs per leg) both using a vast array of powerful genetic tools, and in the physiological context of an appendage-based locomotion scheme. Here we describe a protocol to visualize the leg muscle innervation in an adult fly.

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Here we describe a protocol to visualize the axonal targeting with a florescent protein in adult legs of Drosophila by fixation, mounting, imaging, and post-imaging steps.

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila ...

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of embryogenesis, these organelles undergo dramatic movements powered by actin-based cytoplasmic streaming and motor-driven trafficking along microtubules. The development of a multitude of small, organelle-specific fluorescent probes (FPs) makes it possible to visualize a wide range of different lipid-containing structures in any genotype, allowing live imaging without requiring a genetically encoded fluorophore. This protocol shows how to inject vital dyes and molecular probes into Drosophila embryos to monitor the trafficking of specific organelles by live imaging. This approach is demonstrated by labeling lipid droplets (LDs) and following their bulk movement by particle image velocimetry (PIV). This protocol provides a strategy amenable to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the ER, and for tracking the motion of individual LDs along microtubules. Using commercially available dyes brings the benefits of separation into the violet/blue and far-red regions of the spectrum. By multiplex co-labeling of organelles and/or cytoskeletal elements via microinjection, all the genetic resources in Drosophila are available for trafficking studies without the need to introduce fluorescently tagged proteins. Unlike genetically encoded fluorophores, which have low quantum yields and bleach easily, many of the available dyes allow for rapid and simultaneous capture of several channels with high photon yields.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of ...

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

Retrograde Labeling of Drosophila Motor Neurons Using Fluorescent Lipophilic Dyes

<a href='https://appjove.unicartagenaproxy.elogim.com/v/60716/retrograde-tracing-drosophila-embryonic-motor-neurons-using'>Source: Inal, M. A. et. al., Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes. J. Vis. Exp. (2020)</a>This video demonstrates the method of retrograde labeling of Drosophila motor neurons using lipophilic dyes. The dye enters the axon, travels retrogradely, and labels the neurons green, allowing detailed visualization of their morphology and projections under a confocal microscope.

Source: Inal, M. A. et. al., Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes. J. Vis. Exp. (2020)This video ...

JoVE Encyclopedia of Experiments

Neuroimaging

Developmental & Regenerative Neuroimaging

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells

<a href='https://appjove.unicartagenaproxy.elogim.com/v/63375/studying-membrane-protein-trafficking-drosophila-photoreceptor-cells'>Source: Wagner, K. et al., Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins.&#160;J. Vis. Exp. (2022).</a>This video demonstrates the visualization of membrane protein trafficking in Drosophila photoreceptor cells using eGFP-tagged TRPL proteins. It details the preparation of transgenic flies for imaging, the capture of fluorescence images of photoreceptor cells, and the quantification of TRPL trafficking by measuring relative eGFP fluorescence in the rhabdomeres.

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

Summary

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Chapters in this video

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Electrophysiological Recording in the Drosophila Embryo

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Retrograde Labeling of Drosophila Motor Neurons Using Fluorescent Lipophilic Dyes

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells