The authors thank Arnim Jenett, Aljoscha Nern, and other members of the Rubin laboratory for advice and sharing of unpublished reagents and the Janelia Fly Light Project Team for generating confocal images. The authors also thank the members of the Gaul laboratory for comments on the manuscript.

1. Preparing Flies for MultiColor FlpOut (MCFO) Experiments
NOTE: The MCFO technique refers to a modified version of the so called Flp-mediated stop cassette excision (FlpOut). Transgenic MCFO flies carry a heat shock promoter (hsp)-Flp recombinase and different reporters under UAS control. Each reporter consists of a common backbone of a myristoylated (myr) super folder green fluorescent protein (sfGFP), in which 10 copies of an epitope tag (e.g., HA, FLAG or V5) have been inserted. The resulting non-fluorescent proteins are named &#34;spaghetti monster GFPs&#34; (smGFPs)14 and can be detected using specific antibodies against the different epitope tags.
<ol>
	<li>Cross flies with the MCFO cassette and the hsp-Flp (hsp-Flp; 10XUAS-FRT-stop-FRT-myr-smGFP-HA,10XUAS-FRT-stop-FRT-myr-smGFP-FLAG, 10XUAS-FRT-stop-FRT-myr-smGFP-V5) to the appropriate glial GAL4 driver lines (see Table of Materials).
	<ol>
		<li>As the hsp is already active at 25 &#176;C and few cells may undergo recombination leading to an unspecific labeling, set up crosses at 18 &#176;C in order to avoid leakiness of the system. Wait around 20 days at 18 &#176;C to obtain the F1 generation. After the heat shock (see step 2), maintain flies at 25 &#176;C (Figure 3).</li>
	</ol>
	</li>
</ol>
2. Heat Shock
NOTE: Depending on the insertion site, the efficiency of the hsp-Flp may differ; therefore, the optimal heat shock time has to be optimized. For this protocol, a MCFO fly stock in which the hsp-Flp was inserted on the X chromosome has been used (see Table of Materials).
<ol>
	<li>Transfer the progeny of the cross in step 1.1 (F1 generation flies, 3-4 days old) into new vials containing food and heat shock them at 37 &#176;C in a water bath. 
	NOTE: Young flies (1-2 days old) are very sensitive to heat shock. Wait 1 or 2 more days before performing the heat shock in order to reduce the mortality rate caused by the treatment. Use some of the F1 generation flies as a control, without heat shock.</li>
	<li>During the heat shock, maintain flies in normal vials with food in order to decrease the rate of stress-induced mortality. Put females and males in separate vials.</li>
	<li>Make sure to immerse the entire vial in the water to ensure a homogeneous heat shock. Use a 5-8 min shock as a suitable starting point for sparse labeling of glia cells with many GAL4 driver lines; the shock duration must be optimized for each driver and experiment (shorter or longer heat shock times may be necessary).</li>
	<li>After the heat shock, lay the vials horizontally on the bench for a few minutes in order to allow the flies to recover from the heat shock. 
	NOTE: It is not necessary to change the vials.</li>
	<li>Maintain the flies at 25 &#176;C and dissect (see step 3 and 4) them 2 or more days after the heat shock for optimal reporter expression.</li>
</ol>
3. Dissection Preparation and Solutions
<ol>
	<li>The day of the dissection, prepare fresh fixative solution in 200 &#181;L PCR tubes by mixing 180 &#181;L of S2 cell culture medium with 20 &#181;L of 20% paraformaldehyde (PFA) to obtain a 2% PFA solution. Maintain the fixative solution on ice before and during the dissection. 
	Caution: PFA is toxic and must be handled with care. 
	NOTE: Mix 160 &#181;L of S2 cell culture medium with 40 &#181;L of 20% paraformaldehyde (PFA) in a 200 &#181;L PCR tube to prepare a 4% PFA solution instead of a 2% solution.</li>
	<li>Use one PCR tube per genotype with a maximum of 8-10 brains per tube for optimal staining results.</li>
	<li>Prepare the adult brain washing solution: 0.5% bovine serum albumin, 0.5% Triton X-100 in PBS. 
	NOTE: Depending on the staining, a solution containing 1% Triton X-100 may be needed. A higher concentration of Triton X-100 increases the penetration of the antibody into the tissue and it is necessary to stain regions that lie deep inside the tissue (e.g., synaptic regions in the adult Drosophila brain).</li>
	<li>Prepare the blocking solution: 3% normal goat serum, 3% normal donkey serum, and 0.5% Triton X-100 in PBS.</li>
	<li>The adult brain washing solution and the blocking solution can be stored at 4 &#176;C for a few weeks.</li>
	<li>Put deep depression wells glass plate on ice and fill each well with the following solutions: one with 70% ethanol, one with PBS, and one with S2 cell culture medium.</li>
</ol>
4. Adult Brain Dissection
<ol>
	<li>Position a 3 cm dissecting dish on the stage of a stereomicroscope and fill it with cold S2 cell culture medium. Conduct all the dissections in this medium to ensure that the tissue remains healthy during the dissection procedure. 
	NOTE: Use a dissecting dish lined with several millimeters of black silicone which provides a good background contrast (in the images), and preserve the forceps during the dissection.</li>
	<li>Anesthetize the flies of the desired genotype with CO2.</li>
	<li>With forceps, grab the fly by the wings and wash it in cold 70 % ethanol for 30 s, then with cold PBS for additional 30 s.</li>
	<li>Transfer the fly into the deep depression well that is filled with S2 cell culture medium.</li>
	<li>Repeat the same procedure for all flies of the same genotype and maintain them in cold S2 cell culture medium until dissection, which will preserve the tissue. 
	NOTE: Anesthetize only the number of flies that can be dissected in a time period of about 30 min. Long incubation times in S2 cell culture medium may damage the tissue.</li>
	<li>Grab the fly with forceps and, under a stereomicroscope, submerge the fly in S2 cell culture medium.</li>
	<li>Detach the head from the rest of the body. Discard the body and keep the head submerged in S2 cell culture medium. It is extremely important to hold the head with forceps for the entire duration of the dissection. If the head starts to float, it will be difficult to retrieve it without damaging the tissue.</li>
	<li>Begin the dissection by pulling out one eye, while holding the head with at least one forceps at all times. Make sure that the tip of the forceps is just below the retina to avoid damaging the tissue underneath. Pull out the other eye and begin removing the cuticle until the brain appears.</li>
	<li>Remove all tracheal tissue and cuticle around the brain until the tissue appears clean. Make sure to remove all tracheal tissue around the brain for optimal staining results; the tissues are now ready to be fixed and stained.</li>
	<li>Maintain all the dissected brains in cold S2 cell culture medium before transferring them into the PFA solution in order to time the fixation step.</li>
</ol>
5. Adult Brain Staining
<ol>
	<li>Transfer the isolated brain with a P10 pipette into the PFA solution at room temperature (RT). To avoid tissue damage, never use forceps for transferring the brains after the dissection. Perform all steps in 200 &#181;L PCR tubes on a nutator.</li>
	<li>For all steps, before discarding the old solution with a P200 pipette, allow the brains to settle on the bottom of the tube. Try to remove the supernatant without aspirating the tissue. Protect the tissue from light exposure.</li>
	<li>Fix the brains in 200 &#181;L of 2% PFA in S2 cell culture medium for 1 h or in 4% PFA for 30 minutes. Keep fixation times identical between samples in order to increase reproducibility.</li>
	<li>After 3 (or more) washes of 15 minutes each in 200 &#181;L of adult brain washing solution, block the tissues with 200 &#181;L of blocking solution for 30 minutes. Longer incubation times in blocking solution are possible.</li>
	<li>Incubate the brains overnight at 4 &#176;C with primary antibodies diluted in adult brain washing solution in a total volume of 200 &#181;L. 
	NOTE: The incubation times may vary depending on the type of antibody used. Longer incubations may be necessary.
	<ol>
		<li>For labeling of the three MCFO markers, incubate the brains with anti-HA (1:500), anti-FLAG (DYKDDDDK epitope) (1:100), and anti-V5 primary antibodies.</li>
		<li>Primary directly-conjugated antibodies could be used instead of the normal primary + secondary combination (e.g., anti-V5:DyLight 549 (1:200)). In this case, treat the directly-conjugated antibodies as secondary antibodies. 
		NOTE: For each genotype, keep some brains as controls to verify the specificity of the staining. Skip the primary antibody incubation and go directly to the next step.</li>
	</ol>
	</li>
	<li>Wash the tissues 3 times for 1 h in 200 &#181;L of adult brain washing solution (step 3.3) and incubate them with secondary fluorophore-conjugates/primary directly-conjugated antibodies diluted in adult brain washing solution overnight at 4 &#176;C or for 4 h at RT, in a total volume of 200 &#181;L 
	NOTE: Examples of appropriate antibodies: AlexaFluor 488 (1:250), anti-V5:DyLight 549 (1:200) and DyLight 647 (1:100)-conjugated antibodies.</li>
	<li>Wash the brains 3 times for 1 h in 200 &#181;L of adult brain washing solution, followed by a final wash in 200 &#181;L of PBS overnight at 4 &#176;C or for 1-2 h at RT; the final washing step in PBS removes any traces of Triton X-100 and is necessary for optimal staining results. Mount the brains in mounting medium with an anti-fade agent on glass coverslips.</li>
</ol>
6. Mounting of Brains for Imaging
<ol>
	<li>Trim an imaging spacer to the appropriate size using scissors. For high-resolution microscopy, sandwich the specimen and imaging spacer between two glass coverslips. 
	NOTE: Imaging spacers are thin (0.12 mm thick) adhesive spacers that peel and stick to glass coverslips or microscope slides, allowing the mounting of specimens without compression.</li>
	<li>Using forceps, remove the adhesive liner from one surface and apply the spacer, adhesive side down, on the surface of a glass coverslip (22 mm x 60 mm). Apply 10 &#181;L of mounting medium to a new glass coverslip.</li>
	<li>With a P10 pipette, transfer the brains from the 200 &#181;L tube to the coverslip, next to the drop of mounting medium. Together with the tissues, transfer some PBS in order to maintain the brains in solution.</li>
	<li>With a P10 pipette, move the brains from the PBS to the drop of mounting medium trying to limit the amount of PBS. Transfer the specimens in mounting medium on the coverslip with the imaging spacer. Use enough mounting media to cover the specimen.</li>
	<li>Remove the other adhesive liner from the upper side of the spacer and add a glass coverslip on top of the specimens. With the tip of a forceps, apply a gentle pressure over the adhesive area to seal. Image the mounted tissues immediately or store the slides at -20 &#176;C for later analysis.</li>
</ol>
7. Image Acquisition
<ol>
	<li>Acquire stacks of confocal fluorescence images using a confocal microscope with a 40X (NA = 1.2, water immersion) objective or 63X (NA =1.4, oil immersion) objective (pixel sizes = 1,024 x 1,024 in the xy plane; Distance between two confocal sections = 0.5 &#181;m). Adjust detector gain and offset in each channel in such a way that no saturated pixels and zero pixel values are present in the images; this avoids information loss and ensures use of the full dynamic range of the detector. 
	NOTE: Since 3D imaging is time consuming, the measurements can be automated by scanning multiple samples in parallel at different locations. To avoid evaporation with the water immersion objective, use special oil immersion medium with refractive index n = 1.33.</li>
	<li>Process the confocal stacks for maximum density projections, changes in channel hue, and adjustment of brightness and contrast using any standard image analysis software (see Table of Materials).</li>
</ol>

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None of the authors have competing or conflicting interests.

This protocol describes an easy and efficient method to study the morphology of different cell types within a tissue of interest at high resolution. With the MCFO technique, multiple reporters with different epitope tags are used in combination for multicolor stochastic labeling (Figure 2). Similar to other methods such as Brainbow/Flybow15,16,17, MCFO increases the label diversity through marker co...

Glia, the non-neuronal cell population of the nervous system (NS), were long believed to provide a static framework for neurons and therefore were not studied in detail. However, in humans, glia constitute the vast majority of cells in the NS (~90%) and fall into several different categories, including astrocytes, oligodendrocytes, microglia and Schwann cells. In Drosophila, glia constitute about 10% of the cells in the NS. Intriguingly, their morphologies and functions are remarkably similar to those found in vertebrates1,2. Their morphologies include blood-brain barrier (BBB) forming epithelia, ensheathing, and astrocyte-like cells.
The Drosophila central nervous system (CNS) consists of the following principal structures: cortex regions that contain the neuronal cell bodies; neuropils that harbor synaptic connections; small and large axon tracts that connect the different neuropiles; peripheral nerves that connect sensory organs and muscles with the CNS (Figure 1). Glia are found associated with all these anatomical structures: Cortex glia (CG) in the cortical regions, astrocyte-like glia (ALG) and ensheathing glia (EG) in the neuropile regions, ensheathing glia are also associated with central axon tracts and peripheral nerves (EGN), and finally, two sheet-like glia, perineurial glia (PG) and subperineurial (SPG), which together form a contiguous layer that covers the entire NS (Figure 2).
Previous studies have shown that glia play important roles in the development of the NS; they monitor neuronal cell numbers by reacting to systemically circulating insulin-like peptides, provide trophic support to neurons, such as the astrocyte-neuron lactate shuttle, and eliminate dying neurons by phagocytosis3,4,5,6. In the mature NS, glia maintain the BBB, take up neurotransmitters and maintain ionic homeostasis, act as the major immune cells in the NS, since macrophages cannot breach the BBB, and modulate synaptic activity as well as animal behavior6,7,8,9,10,11.
Whether the different glial subtypes perform specialized functions remains an important open question. However, a systematic genome-wide analysis of glia, especially in the adult, has been hampered by a lack of appropriate genetic tools for their manipulation. Here, a method that allows the efficient and easy characterization of cell shapes to study complex cell-cell interactions is presented. This technique has been applied to characterize the morphology of the different glial subtypes in the adult Drosophila brain, but, depending on the specific GAL4 driver used, it could be adapted to study neurons12,13, any kind of intermingling cells, and in principle any tissue in any developmental stages.

<table><tbody><tr><td>Water bath</td><td>Grant</td><td>GD100</td><td></td></tr><tr><td>PCR tubes</td><td>Sarstedt</td><td>72.737.002</td><td></td></tr><tr><td>Forceps</td><td>Dumont</td><td>11251-20</td><td></td></tr><tr><td>Dissecting dish 30 mm x 12 mmm</td><td>Electron Microscopy Sciences</td><td>70543-30</td><td>Glass dissection dish</td></tr><tr><td>Pyrex 3 Depression Glass Spot Plate</td><td>Corning</td><td>7223-34</td><td>Glass dissection plates</td></tr><tr><td>Sylgard Black</td><td>SYLGARD, Sigma-Aldrich</td><td>805998</td><td>home made with charcoal</td></tr><tr><td>ExpressFive S2 cell culture medium</td><td>Invitrogen</td><td>10486-025</td><td></td></tr><tr><td>20% PFA</td><td>Electron Microscopy Sciences</td><td>15713</td><td></td></tr><tr><td>Triton X-100</td><td>Roth</td><td>3051.3</td><td></td></tr><tr><td>Normal goat serum</td><td>Jackson Laboratories</td><td>005-000-121</td><td></td></tr><tr><td>Normal donkey serum</td><td>Jackson Laboratories</td><td>017-000-121</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma</td><td>A9647</td><td></td></tr><tr><td>Rabbit HA-tag</td><td>Cell Signaling</td><td>C29F4</td><td>Primary antibody, dilution 1:500</td></tr><tr><td>Rat FLAG-tag</td><td>Novus Biologicals</td><td>NBP1-06712</td><td>Primary antibody, dilution 1:100</td></tr><tr><td>Mouse V5-tag:DyLight 549</td><td>AdSerotec</td><td>0411</td><td>Conjugated antibody, dilution 1:200</td></tr><tr><td>anti-rabbit AlexaFluor 488</td><td>Invitrogen</td><td>A11034</td><td>Secondary antibody, dilution 1:250</td></tr><tr><td>anti-rat DyLight 647</td><td>Jackson Laboratories</td><td>712-605-153</td><td>Secondary antibody, dilution 1:100</td></tr><tr><td>Vecta Shield</td><td>Vector Laboratories</td><td>H-1000</td><td></td></tr><tr><td>SlowFate Gold</td><td>Invitrogen</td><td>S36937</td><td></td></tr><tr><td>Secure Seal Spacer</td><td>Grace Biolabs</td><td></td><td>Contact company for ordering</td></tr><tr><td>Microscope cover glass 22 X 60 mm</td><td>Marienfeld</td><td>101152</td><td></td></tr><tr><td>Microscope cover glass 22 x 22 mm</td><td>Roth</td><td>H874</td><td></td></tr><tr><td>Stereo Microscope, Leica MZ6</td><td>Leica</td><td></td><td></td></tr><tr><td>Confocal laser scanning microscope LSM710</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Immersol</td><td>Zeiss</td><td>518 F</td><td>Immersion oil for fluorescence-microscopy, halogen free</td></tr><tr><td>Immersol</td><td>Zeiss</td><td>W 2010</td><td>Immersion fluid for water-immersion objectives, halogen free</td></tr><tr><td>R56F03-GAL4 (EG)</td><td>Bloomington Stock Center</td><td>39157</td><td>GAL4 driver</td></tr><tr><td>R86E01-GAL4 (ALG)</td><td>Bloomington Stock Center</td><td>45914</td><td>GAL4 driver</td></tr><tr><td>hspFlpPestOpt; UAS-FRT-stop-FRT-myr-smGFP-HA, UAS-FRT-stop-FRT-myr-smGFP-FLAG, UAS-FRT-stop-FRT-myr-smGFP-V5</td><td>Bloomington Stock Center</td><td>64085</td><td>UAS reporter (https://bdscweb.webtest.iu.edu/stock/misc/mcfo.php)</td></tr><tr><td>Fiji (Image J)</td><td></td><td></td><td>Image analysis software</td></tr><tr><td>Multi Time Macro</td><td>Zeiss</td><td></td><td>Software for automated scanning</td></tr></tbody></table>

application of multicolor flpout technique to study high resolution single cell morphologies and cell interactions of glia in drosophila

Cells display different morphologies and complex anatomical relationships. How do cells interact with their neighbors? Do the interactions differ between cell types or even within a given type? What kinds of spatial rules do they follow? The answers to such fundamental questions in vivo have been hampered so far by a lack of tools for high resolution single cell labeling. Here, a detailed protocol to target single cells with a MultiColor FlpOut (MCFO) technique is provided. This method relies on three differently tagged reporters (HA, FLAG and V5) under UAS control that are kept silent by a transcriptional terminator flanked by two FRT sites (FRT-stop-FRT). A heat shock pulse induces the expression of a heat shock-induced Flp recombinase, which randomly removes the FRT-stop-FRT cassettes in individual cells: expression occurs only in cells that also express a GAL4 driver. This leads to an array of differently colored cells of a given cell type that allows the visualization of individual cell morphologies at high resolution. As an example, the MCFO technique can be combined with specific glial GAL4 drivers to visualize the morphologies of the different glial subtypes in the adult Drosophila brain.

This section illustrates examples of results that can be obtained by using the MCFO technique in the adult Drosophila brain. Figure 3 shows a schematic of the method. Three differently membrane-tagged reporters (myr-smGFP-HA, myr-smGFP-FLAG and myr-smGFP-V5) under UAS control are kept silent by a transcriptional terminator flanked by two FRT sites (FRT-stop-FRT). A heat shock pulse induces the expression of Flp recombinase which rand...

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Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

Cells display different morphologies and establish a variety of interactions with their neighbors. This protocol describes how to reveal the morphology of single cells and to investigate cell-cell interaction by using the well-established Gal4/UAS expression system.

Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

Cells display different morphologies and complex anatomical relationships. How do cells interact with their neighbors? Do the interactions differ between ...

application-multicolor-flpout-technique-to-study-high-resolution

Research

JoVE Journal

Behavior

8.2K Views.  Ludwig-Maximilians-University Munich. Cells display different morphologies and establish a variety of interactions with their neighbors. This protocol describes how to reveal the morphology of single cells and to investigate cell-cell interaction by using the well-established Gal4/UAS expression system.

Video: Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Neuroscience

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Developmental Biology

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

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

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 ...

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of the fly such as imaginal discs, the larval brain or egg chambers of adult females or the adult intestine can be extracted and kept in culture for imaging with time-lapse microscopy, providing valuable insights into cell and developmental biology. Here, we describe in detail our current protocol for the dissection of Drosophila larval brains, and then present our current approach for immobilizing and orienting larval brains and other tissues on a glass coverslip using Fibrin clots. This immobilization method only requires the addition of Fibrinogen and Thrombin to the culture medium. It is suitable for high-resolution time lapse imaging on inverted microscopes of multiple samples in the same culture dish, minimizes the lateral drifting frequently caused by movements of the microscope stage in multi-point visiting microscopy and allows for the addition and removal of reagents during the course of imaging. We also present custom-made macros that we routinely use to correct for drifting and to extract and process specific quantitative information from time-lapse analysis.

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

This protocol describes applications of sample immobilization using Fibrin clots, limiting drifting, and allowing the addition and washout of reagents during live imaging. Samples are transferred to a drop of Fibrinogen containing culture medium on the surface of a coverslip, after which polymerization is induced by adding thrombin.

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of ...

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 ...

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.

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

Multiplex Detection of Gene Expression in the Intact Drosophila Brain Using Expansion-Assisted Iterative Fluorescence In Situ Hybridization

Understanding gene expression is essential for deciphering cellular functions. However, methods for analyzing the expression of numerous genes in situ within a given tissue remain limited. The EASI-FISH protocol described here has been adapted to detect the expression of dozens of genes in the intact adult Drosophila central nervous system (CNS) using commercially available reagents. This protocol includes a new gel formulation that enhances gel robustness, enabling multiple rounds of hybridization and allowing the embedding of multiple brains per gel. This improvement increases throughput, facilitates optimal comparison of experimental conditions, and reduces reagent costs. Additionally, by employing the GAL4-UAS system for co-detection of green fluorescent protein (GFP), gene expression can be visualized in specific neuronal or glial cell types. Notably, the high resolution achieved through expansion microscopy, combined with the sensitivity of the method, allows for the detection of single RNA transcripts. This approach effectively integrates high image quality with high throughput, making it a powerful tool for studying gene expression throughout the intact fly brain.

Expansion-Assisted Iterative Fluorescence In Situ Hybridization (EASI-FISH) is a robust technique that combines expansion microscopy with fluorescence in situ hybridization (FISH) to detect the expression of multiple genes in thick tissue. This protocol outlines recent advancements in the method and its adaptation for labeling the intact Drosophila central nervous system..

Understanding gene expression is essential for deciphering cellular functions. However, methods for analyzing the expression of numerous genes in situ ...

Application of MultiColor FlpOut Technique to Study High Resolution Single Cell Morphologies and Cell Interactions of Glia in Drosophila

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Przeglądaj więcej filmów

Single Drosophila Ommatidium Dissection and Imaging

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

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

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

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

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

Multiplex Detection of Gene Expression in the Intact Drosophila Brain Using Expansion-Assisted Iterative Fluorescence In Situ Hybridization