Name: visualize drosophila leg motor neuron axons through the adult cuticle
Uploaded: 2018-10-30T14:30:00.000+00:00
Duration: 8 min 33 s
Description: Watch this Scientific Journal Video about Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle at JoVE.com

We thank Robert Renard for preparing fly food medium. This work was supported by an NIH grant NS070644 to R.S.M. and funding from the ALS Association (#256), FRM (#AJE20170537445) and ATIP-Avenir Program to J.E.

1. Leg Dissection and Fixation
<ol>
	<li>Take a glass multi-well plate and fill appropriate number of wells with 70% ethanol. Add 15&#8211;20 CO2-anesthetized flies (of either sex and any age) to each well and by using a brush, gently dab the flies into the ethanol solution until flies are fully submerged. 
	NOTE: This step is to remove the hydrophobicity of the cuticle. Do not wash for more than 1 min, because this increases auto-fluorescence of the cuticle.</li>
	<li>Rinse the flies 3 times with 0.3% nonionic surfactant detergent solution in 1x phosphate buffered saline (PBS). Keep the flies in this solution for at least 10 min. 
	NOTE:&#160;Legs are better fixed when including detergent, which is likely to increase penetration of the fixative inside the leg.</li>
	<li>Place a multi-well plate on the ice along with a tube of 4% paraformaldehyde (PFA). 
	NOTE:&#160;The preparation of fresh 4% PFA from a 16% PFA ethanol free stock solution is critical.</li>
	<li>Use forceps to remove the head and abdomen of the flies without damaging the thoracic segment or the legs. 
	NOTE: Removing the abdomen makes it easier to hold the fly and dissect the legs.</li>
	<li>Dissect the legs from the thoracic segment with forceps and place the legs in the wells containing 4% PFA. For this, push gently but firmly at the coxa-thorax junction using the tip of fine forceps until the leg detaches.</li>
	<li>Fix the legs in 4% PFA overnight at 4 &#176;C (approximately 20 h total).</li>
	<li>Wash the legs with 0.3% nonionic surfactant detergent solution in 1x PBS, 5x for 20 min each.</li>
	<li>Replace the washing buffer with mounting medium. Keep the leg in mounting medium for at least a day prior to mounting to allow for its complete penetration into the leg. 
	NOTE: If the mounting medium is highly viscous, dilute the mounting medium to 80% with 1x PBS because the sudden change in viscosity can cause the cuticle to collapse and damage the overall structure of the leg. Depending on the Gal4 driver, flies can be preserved for 1 to 3 weeks at 4 &#176;C in mounting media until mounting.</li>
</ol>
2. Leg Mounting
<ol>
	<li>Place approximately 20 &#181;L of 70% glycerol on the left-hand side of a microscope slide and cover with a square 22 x 22 mm2 coverslip (Figure 2A, B). Pipette about 10 &#181;L of mounting medium along a line parallel to and at a small distance from the right edge of this coverslip. Add another 30 &#181;L of mounting medium further on the right side of the slide (Figure 2C). 
	NOTE: When applying a coverslip over the legs (see below) the mounting medium will gently spread under the coverslip and around the legs without displacing them.</li>
	<li>Using fine forceps lift the leg from the solution and gently put it on the mounting medium near the left coverslip. Do the same for each leg and align them from top to bottom (Figure 2D).
	<ol>
		<li>Lift and transfer the legs in a drop of medium held between both tips of the forceps. Orient the legs in two ways: external side up or down.</li>
	</ol>
	</li>
	<li>Once all the legs (up to 6&#8211;8 legs can be mounted) are properly aligned, put a second coverslip over the legs such that this coverslip rests slightly on the previously placed coverslip (Figure 2E) to allow for space between the coverslip and the tissue and to prevent the legs from getting damaged (Figure 2G). 
	NOTE: Alternatively, use sticker wells or orthodontic wax to create space between the coverslip and the slide.</li>
	<li>Use a nail polish to secure the position of the coverslips at each corner (Figure 2F). 
	NOTE: The tissue is now ready to image.</li>
</ol>
3. Imaging
<ol>
	<li>Set up a single track using the 488 nm laser and two detectors simultaneously to obtain both the GFP fluorescence and cuticle auto-fluorescence. Using the range control or the slider display in the spectral window of the light path window of the software, set the detection range of one detector (detector 1) between 498&#8211;535 nm and that of the other one (detector 2) between 566-652 nm. 
	NOTE: Typically, the detector 1 should be a GaAsP detector and the detector 2 a photomultiplier tube. The 498-535 channel optimally detects GFP signal but also detects auto fluorescence from the cuticle. However, the 566&#8211;652 channel detects only the auto fluorescence from the cuticle (Figure 3A).</li>
	<li>Use a 20X or 25X oil immersion objective, 1024 x 1024 pixels resolution, 12-bit depth, and set Z spacing as 1 &#181;m. Set pixel dwell time as approximately 1.58 &#181;s, and average 2 frames/image. Use objectives with high numerical aperture.</li>
	<li>Set all other settings, depending on the confocal microscope and imaging software being used.
	<ol>
		<li>Make sure to obtain a brighter cuticle signal from the 566&#8211;652 detector rather than from the 498&#8211;535 detector. To do this, use the same laser power of the 488-laser for both detectors. Using the saturation markers, first adjust the gain of detector 1 to obtain a bright enough GFP signal (Figure 3B, D). Then adjust the gain of detector 2 to ensure that some areas with high cuticle signal (edges of legs, joints) produce a saturated signal in this detector (Figure 3C, E).</li>
		<li>If the leg is extended and too large to be imaged in a single frame, use the Tile or Position option from the microscope imaging software.</li>
	</ol>
	</li>
	<li>Reconstruct the final images either using the proprietary microscope imaging software or using ImageJ/FIJI freeware (see below).</li>
</ol>
4. Post Imaging Processing
<ol>
	<li>Open the confocal stack in ImageJ/FIJI.
	<ol>
		<li>Use the Bioformats plugin (Plugins| Bio-Formats| Bio-formats Importer) in FIJI to open images that are not in .TIF format from proprietary imaging software packages9.</li>
	</ol>
	</li>
	<li>Split the channels by selecting Image| Color| Split Channels. 
	NOTE: If images from several positions were made and must be recombined, use the pairwise stitching plugin in FIJI from (Plugins &#62; Stitching &#62; Pairwise Stitching)10,11.</li>
	<li>To subtract the cuticle signal from the GFP signal, open the Image Calculator (Process| Image Calculator): select the stack from detector 1 as Image 1 (GFP + cuticle) (Figure 4A), on the operation window select subtract, and select the stack from detector 2 as Image 2 (cuticle) (Figure 4B). As a result, only the endogenous GFP signal (GFP) is obtained (Figure 4C).
	<ol>
		<li>Merge back this stack (GFP) with the &#8216;cuticle-only&#8217; stack obtained from detector 2 (Figure 4B Image| Color| Merge Channels) to obtain an RGB image comprising of the tissue-specific GFP signal as well as the cuticle signal, which helps identifying the axon arbors within the leg segments (Figure 4D).</li>
		<li>Adjust contrast and brightness of each image (Image| Adjust| Brightness/Contrast) and then generate a single RGB image (Image| Type| RGB Color). 
		NOTE: To generate a maximum projection of the Z-stack (Image| Stacks| Z Project&#8230;), use maximum intensity projection for the GFP signal and average intensity for the cuticle, which can then be adjusted for brightness prior to merging the two channels.</li>
	</ol>
	</li>
	<li>Alternatively, use a macro for automatic subtraction and processing of images in ImageJ/FIJI. Copy the text in Supplemental File 1 in the macro editor (Plugins| New| Macro), save the macro in the Plugins folder, and restart ImageJ/FIJI once to be able to use the macro.
	<ol>
		<li>To use the macro, open the confocal stack, and open the Brightness/Contrast window (Image| Adjust| Brightness/Contrast). Then run the macro (Plugin| Macro| Run) and follow instructions.</li>
		<li>When asked to specify the operation (Image calculator window), choose Image 1 as stack 1 (GFP), Image 2 as stack 2 (cuticle), and Subtract. 
		Note: The macro generates a maximal projection image of the processed GFP signal (MAX_Result of stack1), an average projection of the cuticle (AVG_stack2), the result of the substraction of the cuticle stack from the GFP stack (Result of stack1), and the unprocessed cuticle stack (stack2).</li>
		<li>When asked to adjust contrast, use the controls in the brightness/control window to adjust the brightness/contrast of the maximal projection image of GFP, and of the average projection of the cuticle, to generate an RGB merged image of both signals showing the GFP in green and the cuticle in gray. Merge Result of stack1, and stack2, to similarly generate a combined GFP and cuticle stack that can be used in the next stage.</li>
	</ol>
	</li>
	<li>To visualize the legs in three dimensions, use specialized 3D software with the newly generated stacks (Figure 4E).
	<ol>
		<li>Open the RGB stack with 3D software (see Table of Materials). In the pop-up dialog window, select all channels in the mode conversion section and enter the size of a voxel (volume of a pixel). 
		Note: All necessary information is contained in the image files from whatever proprietary microscope software being used.</li>
		<li>On channel 2 module (GFP signal), right-click and select Display | volren. Then, left-click on the volren module. In the properties section (bottom left of the screen) click left on edit and select volrengreen.col. On channel 1 module (cuticle signal) right-click and select Display | volren. Then, left-click on the volren module. In the Properties section left-click on advanced and select ddr (a transparent radiograph-like display), then adjust the gamma value to see the cuticle background (Figure 4E).</li>
		<li>To generate a 3D rotation, right-click on the project stack module. Then select animate| rotate. Left-click on the rotate module.</li>
		<li>In the properties section click on use bbox center. On the rotate module right-click and select compute| movie maker. On the movie maker module click the left mouse button. In the properties section left-click left on Advanced (top right of the properties section).</li>
		<li>In the file name field fill the name of the movie rotation. In the quality field write 1 (max quality). Leave frame at 200 if an 8.3 s rotation is desired (this number can be decreased or increased to modify the speed of the rotation). Then press apply (green button, bottom left of the properties section (See Video 1)).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The cuticle of adult Drosophila and of other arthropods, which contains many dark pigments, is a major obstacle for viewing structures inside their body. In addition, it is strongly auto fluorescent which is made worse by fixation. These two features are very problematic for observations of fluorescent dyes or molecules inside the body of animals with an exoskeleton.
The procedure that we have described and that we routinely use in the lab yields clean and detailed images of axon trajectories ...

Like the vertebrate limb, the Drosophila adult leg is organized into segments. Each fly leg contains 14 muscles, each of which comprises multiple muscle fibers1,2. The cell bodies of the adult leg MNs are located in the T1 (prothoracic), T2 (mesothoracic), and T3 (metathoracic) ganglia on each side of the ventral nerve cord (VNC), a structural analogous to the vertebrate spinal cord (Figure 1). There are approximately 50 MNs in each ganglia, which target muscles in four segments of the ipsilateral leg (coxa, trochanter, femur, and tibia) (Figure 1)3. Importantly, each individual adult leg MN has a unique morphological identity that is highly stereotyped between animals3,4. All these unique MNs are derived from 11 stem cells, called neuroblasts (NBs) producing leg MNs during the larval stages3,4. At the end of the larval stages all the immature postmitotic MNs differentiate during metamorphosis to acquire their specific dendritic arbors and axonal terminal targets that define their unique morphology3,4. Previously we tested the hypothesis that a combinatorial code of transcription factors (TFs) specifies the unique morphology of each Drosophila adult leg MN5. As a model, we used lineage B, one of the 11 NB lineages which produces seven out of the MNs and demonstrated that a combinatorial code of TFs expressed in postmitotic adult leg MNs dictates their individual morphologies. By reprograming the TF code of MNs we have been able to switch MN morphologies in a predictable manner. We call these TFs: mTFs (morphological TFs)5.
One of the most challenging parts of the morphological analyses of adult MNs is to visualize the axons through a thick and auto-fluorescent cuticle with high resolution. We usually label axons with a membrane-tagged GFP that is expressed in MNs with a binary expression system, such as DVglut-Gal4/UAS-mCD8::GFP or DVglut-QF/ QUAS mCD8::GFP, where DVglut is a strong driver expressed in motoneurons6. By combining these tools with other clonal techniques such as mosaic analysis with a repressible marker (MARCM)7, cis-MARCM8, or MARCMbow5, we can restrict the GFP expression to subpopulations of MNs making the phenotypic analysis of axons easier. We have generated a protocol in order to keep leg MN axonal morphology intact for imaging and subsequent 3D reconstruction by addressing specific issues intrinsic to the adult Drosophila leg such as (1) fixation of the internal structures of the adult leg without affecting axon morphology, endogenous fluorescent expression, and leg musculature, (2) mounting of the leg to preserve the overall structure under a coverslip and in the appropriate orientation for imaging, and (3) image processing to obtain the cuticle background as well as axonal fluorescent signal. While this protocol has been detailed for the detection of fluorescent expression in MN axons, it can be applied to visualize other components of leg neuromusculature in arthropods.

<table><tbody><tr><td>Ethanol absolute</td><td>Fisher</td><td>E/6550DF/17</td><td>Absolute analytical reagent grade</td></tr><tr><td>nonionic surfactant detergent</td><td>Sigma-Aldrich</td><td>T8787</td><td>Triton X-100, for molecular biology</td></tr><tr><td>Fine forceps</td><td>Sigma-Aldrich</td><td>F6521</td><td>Jewelers forceps, Dumont No. 5</td></tr><tr><td>Glass multi-well plate</td><td>Electron Microscopy Sciences</td><td>71563-01</td><td>9 cavity Pyrex, 100 mm x 85 mm</td></tr><tr><td>PFA</td><td>Thermofisher</td><td>28908</td><td>Pierc 16% Formaldehyde (w/v), Methanol-free</td></tr><tr><td>Glycerol</td><td>Fisher BioReagents</td><td>BP 229-1</td><td>Glycerol (Molecular Biology)</td></tr><tr><td>Spacers</td><td>Sun Jin Lab Co</td><td>IS006</td><td>iSpacer, four wells, around 12 &amp;mu;L working volume per well, 7 mm diameter, 0.18 mm deep</td></tr><tr><td>Square 22 mm x 22 mm coverslips</td><td>Fisher Scientific</td><td>FIS#12-541-B</td><td>No.1.5-0.16 to 0.19 mm thick</td></tr><tr><td>Mounting Medium</td><td>Vector Laboratories</td><td>H-1000</td><td>Vectashield Antifade Mounting Medium</td></tr><tr><td>Confocal microscope</td><td>Carl Zeiss</td><td></td><td>LSM780; objective used LD LCI Plan-Apochromat&lt;br/&gt; 25X/0.8 Imm Korr DIC M27 (oil/&lt;br/&gt; silicon/glycerol/water&lt;br/&gt; immersion) (420852-9871-000)</td></tr><tr><td>imaging software</td><td>Carl Zeiss</td><td></td><td>ZEN 2011</td></tr><tr><td>3D-Image software</td><td>ThermoFisher Scientific</td><td></td><td>Amira 6.4</td></tr><tr><td>ImageJ</td><td>National Institutes of Health</td><td>https://imagej.nih.gov/ij/</td><td>ImageJ/FIJI</td></tr></tbody></table>

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.

As shown in Figure 4, this procedure allows excellent imaging of GFP-labeled axons in adult Drosophila legs, together with their terminal arbors. Importantly a clean GFP signal is obtained without any contamination from the fluorescence emitted by the leg cuticle. The signal from the cuticle can then be combined with the GFP signal to identify the positioning of axons in the legs (Figure 4E, Figure 1...

Watch this Scientific Journal Video about Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle at JoVE.com

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.

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 ...

visualize-drosophila-leg-motor-neuron-axons-through-the-adult-cuticle

Research

JoVE Journal

Developmental Biology

9.3K Views.  Institut de Génomique Fonctionnelle de Lyon, ENS de Lyon, CNRS. 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.

Video: Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

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

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

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

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

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

Neuroscience

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 ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

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

Preparation of Drosophila Central Neurons for in situ Patch Clamping

Short generation times and facile genetic techniques make the fruit fly Drosophila melanogaster an excellent genetic model in fundamental neuroscience research. Ion channels are the basis of all behavior since they mediate neuronal excitability. The first voltage gated ion channel cloned was the Drosophila voltage gated potassium channel Shaker1,2. Toward understanding the role of ion channels and membrane excitability for nervous system function it is useful to combine powerful genetic tools available in Drosophila with in situ patch clamp recordings. For many years such recordings have been hampered by the small size of the Drosophila CNS. Furthermore, a robust sheath made of glia and collagen constituted obstacles for patch pipette access to central neurons. Removal of this sheath is a necessary precondition for patch clamp recordings from any neuron in the adult Drosophila CNS. In recent years scientists have been able to conduct in situ patch clamp recordings from neurons in the adult brain3,4 and ventral nerve cord of embryonic5,6, larval7,8,9,10, and adult Drosophila11,12,13,14. A stable giga-seal is the main precondition for a good patch and depends on clean contact of the patch pipette with the cell membrane to avoid leak currents. Therefore, for whole cell in situ patch clamp recordings from adult Drosophila neurons must be cleaned thoroughly. In the first step, the ganglionic sheath has to be treated enzymatically and mechanically removed to make the target cells accessible. In the second step, the cell membrane has to be polished so that no layer of glia, collagen or other material may disturb giga-seal formation. This article describes how to prepare an identified central neuron in the Drosophila ventral nerve cord, the flight motoneuron 5 (MN515), for somatic whole cell patch clamp recordings. Identification and visibility of the neuron is achieved by targeted expression of GFP in MN5. We do not aim to explain the patch clamp technique itself.

Preparation of Drosophila Central Neurons for in situ Patch Clamping

In situ patch clamp recordings are used for electrophysiological characterization of neurons in intact circuitry. In the Drosophila genetic model patch clamping is difficult because the CNS is small and surrounded by a robust sheath. This article describes the procedure to remove the sheath and clean neurons for subsequent patch clamp recordings.

Short generation times and facile genetic techniques make the fruit fly Drosophila melanogaster an excellent genetic model in fundamental neuroscience ...

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.

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.

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 ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

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 ...

Dissection and Immunohistochemistry of the Drosophila Adult Leg to Detect Changes at the Neuromuscular Junction for an Identified Motor Neuron

Drosophila melanogaster represents a genetically tractable model to study neuronal structure and function, and subsequent changes in disease states. The well characterized larval neuromuscular junction is often used for such studies. However, rapid larval development followed by muscle histolysis and nervous system remodeling during metamorphosis makes this model problematic for the study of slow age-dependent degenerative changes like those occurring in amyotrophic lateral sclerosis. Alternatively, adult flies live for 90 days and the adult leg can be used to study motor neuron changes over the course of adult lifespan using in vivo fluorescent imaging through the cuticle. Here, we describe a leg dissection technique coupled with immunocytochemistry, which allows for the study of molecular changes at the neuromuscular junction of identified adult leg motor neurons. These techniques can be coupled with a myriad of antibodies labeling both pre- and post-synaptic structures. Together these procedures allow a more complete characterization of slow age-dependent changes in adult flies and can be applied across multiple motor neuron disease models.

Dissection and Immunohistochemistry of the Drosophila Adult Leg to Detect Changes at the Neuromuscular Junction for an Identified Motor Neuron

We describe a dissection technique that preserves the architecture of the neuromuscular junction and enables a detailed immunocytochemical study of motor neurons in the adult Drosophila leg.

Drosophila melanogaster represents a genetically tractable model to study neuronal structure and function, and subsequent changes in disease states. The ...

Processing Insect Legs for Fluorescence Microscopy: A Method to Preserve Neuromuscular Structures for Imaging

Source: Guan, W., et al. <a href="https://www.jove.com/video/58365/" target="_blank">Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle</a>. J. Vis. Exp. (2018).
This video describes a method to dissect, fix, and mount the Drosophila adult leg while preserving its neuromusculature intact for imaging analysis.

Source: Guan, W., et al. Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle. J. Vis. Exp. (2018).
This video describes a method to ...

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

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Chapters in this video

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

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Preparation of Drosophila Central Neurons for in situ Patch Clamping

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

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

Dissection and Immunohistochemistry of the Drosophila Adult Leg to Detect Changes at the Neuromuscular Junction for an Identified Motor Neuron

Processing Insect Legs for Fluorescence Microscopy: A Method to Preserve Neuromuscular Structures for Imaging