This work was supported by the National Institutes of Health (R01 NS110727) to D.T.B.

1. Generation of transgenic flies
<ol>
	<li>To generate transgenic flies for this experiment, collect OK371-Gal425 virgin female flies and males of UAS-TDP-43M337V 26 (Figure 1A) by anesthetizing flies with CO2 on a pad to sort.</li>
	<li>Sort anesthetized flies into vials with standard Drosophila media for the cross. Place labeled vials at 25 &#176;C for the next generation to emerge. 
	NOTE: Clear the adults from the vials before the progeny emerge to ensure the proper genotype.</li>
	<li>Once progeny emerge, collect the transgenic flies into vials and sort by sex to begin aging for experimental conditions.</li>
	<li>Once flies are collected, transfer flies to fresh food every 2 days until the flies are 21 days old.</li>
</ol>
2. Dissection prep
<ol>
	<li>To prepare for dissections, obtain room temperature phosphate buffered saline (1x PBS), a 10 cm dissection dish coated with a silicone elastomer, straight edge dissecting scissors, one set of blunt dissection forceps, a P200 pipette and pipette tips, 2.0 mL microcentrifuge tubes, standard office scissors, 70% ethanol, a 6 cm Petri dish, and 32% formaldehyde diluted to 4% with 1x PBS.</li>
	<li>Label tubes for each genotype or condition and add 900 &#181;L of 1x PBS (room temp) and 150 &#181;L of 32% formaldehyde to each tube. Wear gloves and safety glasses when preparing the 4% formaldehyde fixative.</li>
	<li>Anesthetize 6&#8210;10 flies per group directly from the vial with CO2 and submerge flies into a 6 cm Petri dish with 70% ethanol. Press flies down into the ethanol using a paint brush to ensure that specimens are fully submerged. This will remove the layer of oil on the outer cuticle.</li>
</ol>
3. Thorax isolation and fixation
<ol>
	<li>Before dissecting each specimen, add approximately 7&#8210;10 mL of 1x PBS to the dissection dish coated with silicone elastomer. This volume should ensure that the tissue samples are completely submerged.</li>
	<li>Transfer one fly to the dissection dish from the 70% ethanol using blunt forceps and grasping either the wings or the legs.</li>
	<li>Focus the sample in the dissection dish under a dissecting microscope. Next submerge the sample in 1x PBS, and carefully remove the wings using blunt Dumont #5 fine forceps.</li>
	<li>Using Vannas straight edge spring dissection scissors, remove the legs by creating a small incision in the ventral side of the cuticle. In step 3.8, this incision will allow the formaldehyde to penetrate the tissue.</li>
	<li>Take the scissors in one hand and hold the forceps in the other to position the fly ventral side up. While holding the specimen in place with the blunt forceps, remove the head and abdomen with the dissection scissors.</li>
	<li>Transfer the isolated thorax using the modified pipette tip into the labeled tube, from step 3.2. 
	NOTE: Set the pipette to 40 &#181;L to avoid adding extra 1x PBS to the fixative.</li>
	<li>Repeat steps 3.2&#8210;3.6 above for each specimen.</li>
	<li>Fix samples for 30 min at room temperature.</li>
	<li>Remove the fix using a Pasteur pipette and discard it in the proper waste container under the fume hood. Rinse samples three times with 1.5 mL of 1x PBS each using a Pasteur pipette. Complete a fourth rinse using only 750 &#181;L and leave tissues in 1x PBS. 
	NOTE: At this point, tissue samples can remain at 4 &#176;C for up to 3 days before proceeding to the next steps.</li>
</ol>
4. Flash freezing and thorax bisection
<ol>
	<li>Before beginning the bisections, fill a Dewar flask with liquid nitrogen wearing proper cryo-protective gloves and safety glasses. Obtain a blade breaker, feather blades, one pair of fine forceps, ice, ice-cold 1x PBS, and cryogenic tweezers.</li>
	<li>Prepare an ice bucket to keep 1x PBS ice-cold.</li>
	<li>Use the blade breaker to grab the feather blade at an angle, and bend the blade in order to break off a small piece. The blade breaker can then lock the blade in position for use as a small scalpel. 
	NOTE: One blade should last for all groups. Switch if blade breaks or becomes dull.</li>
	<li>Add a clean pipette tip to the P200 and remove 1/5th of the tip to transport samples.</li>
	<li>Prepare a new microcentrifuge tube for each group and add 200 &#181;L of 1x PBS to each tube. This second tube will be used to collect the final DLM preps.</li>
	<li>Remove all 1x PBS from tubes using a Pasteur pipette.</li>
	<li>Wearing proper protective equipment, submerge the tube into the liquid nitrogen flask for 10 s with the cryogenic tweezers. 
	NOTE: The tubes should be closed tightly to keep the tube from exploding.</li>
	<li>Remove the tube from the liquid nitrogen and add approximately 300 &#181;L of ice-cold 1x PBS to the samples with a Pasteur pipette. Keep the samples on ice.</li>
	<li>Add ice-cold 1x PBS to the 10 cm dissection dish coated with silicone elastomer and dispense the first thorax with the modified 200 &#181;L pipette.</li>
	<li>Place the thorax ventral side up. In one hand use a dull pair of forceps to position the thorax and in the other use a fine pair of forceps to remove some of the thoracic ganglion to expose the midline of the thorax.</li>
	<li>Use the midline of the thorax as a guide to make a shallow cut through 1/3rd of the thorax with the blade.</li>
	<li>Remove the blade from the thorax and position the thorax at a 45&#176; angle with the blunt forceps. Reinsert the blade and cut straight down the midline of the thorax. This will result in two hemithoraces.</li>
	<li>Take one hemithorax at a time and remove the excess tissue under DLM muscle fiber F (Figure 1B), the most ventral fiber. Use the blade to carefully make one or two cuts to remove the excess tissue without damaging the DLMs.</li>
	<li>Once isolated, transfer the hemithorax to the correct tube with 1x PBS.</li>
	<li>Repeat steps 4.6&#8210;4.14 until 10 dissected hemithoraces per group are made.</li>
</ol>
5. Structural staining
<ol>
	<li>After bisecting the thorax samples, place the tissue in blocking buffer (1x PBS with 0.1% normal goat serum, and 0.2% Triton X-100 at pH 7.4) to permeabilize the tissue and prevent non-specific staining. Use a Pasteur pipette to remove excess 1x PBS and add 1.5 mL of blocking buffer to each tube. Block tissues for at least 1 h at 4 &#176;C.</li>
	<li>Prepare the samples for structural staining using a fluorescently conjugated antibody, horseradish peroxidase 488 (anti-HRP-488) at a dilution of 1:200 and Phalloidin-647 at a dilution of 1:1000 in blocking buffer to stain motor neurons and muscle tissue, respectively. Make enough stain to have 150 &#181;L per tube. Store the stain at 4 &#176;C covered in foil or in a dark box until ready for staining.</li>
	<li>After blocking, remove the excess blocking buffer with a glass Pasteur pipette.</li>
	<li>Before dispensing the structural stain, vortex the stain. Add 150 &#181;L of the stain to each tube. Place the samples in a dark box on the rotator at room temperature for 2 h.</li>
	<li>Remove the stain and wash the tissues four times in 1.5 mL of room temp 1x PBS with 0.3% Triton X-100 for 5 min on the rotator in a dark box. The samples are now ready to mount to a slide.</li>
</ol>
6. Mounting tissue
<ol>
	<li>After washing samples in PBST, prepare a microscope slide to mount tissue for staining. Prepare additional supplies including glass cover slips, a P200 pipette, 200 &#181;L pipette tips, scissors, clear reinforcements, straight edge forceps, anti-fade fluorescent mounting media, nail polish, and a dark box to cover the slides.</li>
	<li>Label the slide to identify the samples and clean the slide with kimwipes to ensure there are no smudges.</li>
	<li>To ensure the hemithorax samples are not damaged by the cover slip, build a &#8220;bridge&#8221; using reinforcement labels. Take one reinforcement label, cut it in half, and place each half approximately 15 mm apart. This distance must be smaller than the width of the cover slip. Repeat this step four times to complete a &#8220;bridge&#8221; that is 5 labels high.</li>
	<li>Take the P200 pipette and modify a tip by cutting off 1/5th of the tip to transfer the samples to the slide. Samples should be transferred onto the slide in the center of the bridge.</li>
	<li>Take the edge of a lab wipe and remove any excess PBST. Using forceps, arrange the DLMs such that all samples are facing muscle side up and cuticle side down.</li>
	<li>Using a standard P200 pipette tip, apply 70 &#181;L of mounting media to the slide, avoiding air bubbles. Dispense the media in a circular pattern inside the reinforcements starting from the outside into the center.</li>
	<li>Place a cover slip over the reinforcements.</li>
	<li>Use nail polish to coat the outside edges around the perimeter of the coverslip. Apply generously to form a complete seal of the tissue.</li>
	<li>Place the slide on a flat surface in the dark, allowing at least 10 min to dry and prevent photo-bleaching or loss of fluorescence. Slides may now be used for imaging immediately, or otherwise stored in a slide folder at -20 &#176;C for later viewing.</li>
</ol>
7. Alternative: Staining with primary antibodies
NOTE: This section is optional and should be used directly between sections 4 and 5 if desired.
<ol>
	<li>To stain tissue with primary antibodies, submerge tissue in blocking buffer for at least 1 h.</li>
	<li>Prepare primary antibody with proper dilution in blocking buffer. At minimum, prepare enough antibody mixture to have 150 &#181;L per group. Note that the samples are kept still. Store at 4 &#176;C until ready for use.</li>
	<li>Remove excess blocking buffer with a Pasteur pipette. Briefly vortex the primary antibody and add 150 &#181;L of antibody mixture to each group and place samples at 4 &#176;C overnight.</li>
	<li>On the next day, remove primary antibody and wash tissue 4 times with PBST for 5 min each on a rotator.</li>
	<li>Prepare secondary stain in blocking buffer. Add the secondary stain to the sample and then keep it a room temperature for 2 h in a dark box on the rotator. 
	NOTE: The secondary staining can also include HRP and phalloidin.</li>
	<li>After the 2 h incubation, to remove the secondary stain wash tissue 4 times for 5 min with PBST and proceed to mounting.</li>
</ol>

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The authors have no conflicts of interest to disclose.

Using the methods described in this protocol, we provide a straightforward approach for dissection of the DLM tissue and demonstrate how this can be applied to assess synaptic integrity through structural staining and synaptic markers in adult Drosophila. One critical step in the protocol that makes the DLM tissue easier to dissect is the flash freezing with liquid nitrogen. Without this step, the tissue is less firm and more difficult to cut precisely as observed in Figure 3. This protocol builds upon previous dissection methods to allow the preservation of both motor neurons and muscle tissue18,19,20,21,22,23. One limitation of this protocol is that when making the cut down the midline for the bisection, it can be difficult to get two clean preps per thorax. One way to ensure at least one hemithorax per fly, you can purposely cut off to one side of the thorax to get one clean prep. With this modification, one may also need to remove additional excess tissue from the cut to clean up the sample with the blade breaker. For those new to this technique, with continued practice, accuracy of the bisection will increase.
The method described here allows researchers to easily assess structural integrity of adult DLM NMJs at any time throughout their lifespan. A major advantage of this protocol is the ability to access synaptic integrity in neurodegenerative disease models by using synaptic markers. We demonstrate that this application can help visualize changes in gross morphology with structural staining (Figure 1C&#8210;H). Additionally, synaptic integrity can be assessed with staining of presynaptic markers including but not limited to Synapsin28 (Figure 2A&#8210;F), Syntaxin29 (Figure 2G&#8210;L) and BRP30 (Figure 2M&#8210;R). The postsynaptic muscle tissue can also be assessed using the Glutamate Receptor III subunit antibody31 (Figure 2S&#8210;X), demonstrating the utility of this protocol.
Researchers can also utilize this dissection method to complement functional data to comprehensively examine the structural integrity of synapses associated with a wide variety of diseases. These synapses also allow for functional analysis through electrophysiological recordings32,33,34 and the flight assay10. This protocol can also provide ease of access to the tissue for many applications and assays. Future studies, for example, could use this protocol to quantify synaptic changes through quantification of the density and number of synapses15,16. While the protocol described here specifically examines synaptic integrity of motor neurons, complementary protocols for assessing muscle cell loss can also be performed with this dissection using TUNEL staining35. To examine neuronal loss, dissection of the thoracic ganglion36 could also be used with TUNEL staining. We expect that the dissection described here will have more applications to future studies assessing age-related pathologies as well as neurodegenerative diseases.

Synaptic dysfunction is among the earliest known hallmarks of most major neurodegenerative diseases1,2,3,4,5,6. However, very little is known regarding how these structural and functional impairments relate to later stages of disease progression. Drosophila has proven to be a useful model system for understanding synapse growth and development using larval NMJs7,8,9. However, the third larval instar stage only lasts a few days, limiting their utility in studying progressive, age-dependent neurodegeneration. An alternative to assessing larval NMJs is to examine synaptic structures in adult Drosophila, such as the synapses formed on the Dorsal Longitudinal Muscles (DLMs) that are required for flight10,11,12,13,14,15,16. These tripartite synapses are structurally organized in a similar manner to mammalian synapses17, providing a unique advantage for assessing models of neurodegenerative diseases.
Here we describe a straightforward method for analyzing the structural integrity of adult NMJs in a Drosophila model of neurodegeneration. Previous DLM dissection methods and studies have emphasized the importance of preserving muscle tissue for a variety of applications18,19,20,21,22,23. Our protocol provides a comprehensive method to preserve both neuronal and muscle tissue to investigate neurodegenerative diseases. Another major component of studying these diseases is the ability to understand neuronal loss in an age dependent manner. Previous work provides a critical and in-depth understanding of how the DLM NMJs are formed during metamorphosis into early adulthood11,12,14,15,16,24. Our protocol establishes a method to build upon this work to investigate DLM NMJs in an age-dependent manner in aging and neurodegenerative diseases.

<table><tbody><tr><td>32% Formaldehyde</td><td>Electron Microscopy Sciences</td><td>15714</td><td>Tissue preservation</td></tr><tr><td>Alexa Fluor 568 goat anti mouse</td><td>Fisher Scientific</td><td>A11031</td><td>Labels primary antibodies. Used at 1:200 concentration.</td></tr><tr><td>Alexa Fluor 568 goat anti rabbit</td><td>Fisher Scientific</td><td>A11036</td><td>Labels primary antibodies. Used at 1:200 concentration.</td></tr><tr><td>anti- Bruchpilot (BRP) antibody</td><td>Developmental Studies Hybridoma Bank</td><td>NC82</td><td>Stains the active zones in presynaptic neurons. Used at 1:25 concentration.</td></tr><tr><td>anti-GluRIII antibody</td><td>Gift from Aaron DiAntonio</td><td>N/A</td><td>Labels glutamate receptor subunits. Used at 1:1000 concentration.</td></tr><tr><td>anti-Synapsin antibody</td><td>Developmental Studies Hybridoma Bank</td><td>3C11</td><td>Labels the synaptic protein synapsin. Used at 1:50 concentration.</td></tr><tr><td>anti-Syntaxin antibody</td><td>Developmental Studies Hybridoma Bank</td><td>8C3</td><td>labels the synaptic protein syntaxin. Used at 1:10 concentration.</td></tr><tr><td>BenchRocker</td><td>Genesee Scientific</td><td>31-302</td><td>Rotating samples during staining</td></tr><tr><td>Blade Breaker</td><td>Fine Science Tools</td><td>10053-09</td><td>Used for holding feather blade</td></tr><tr><td>cover slips</td><td>Fisher Scientific</td><td>12548A</td><td>For mounting tissue</td></tr><tr><td>cryogenic gloves</td><td>VWR</td><td>97008-198</td><td>protect hands from liquid nitrogen</td></tr><tr><td>cryogenic tweezers</td><td>VWR</td><td>82027-432</td><td>Hold 2.0 mL tube in liquid nitrogen</td></tr><tr><td>dewar flask-1900 mL</td><td>Thomas Scientific</td><td>5028M54</td><td>Hold liquid nitrogen</td></tr><tr><td>Feather Blades</td><td>Electron Microscopy Sciences</td><td>72002-01</td><td>Scalpel Blades</td></tr><tr><td>Fine Forecps x 2</td><td>Fine Science Tools</td><td>11252-20</td><td>One fine pair for Clearing midline of thorax. The other pair can be dulled using a sharpening stone.</td></tr><tr><td>FITC-conjugated anti HRP</td><td>Jackson Laboratories</td><td>123-545-021</td><td>Stains Motor Neurons. Used at 1:100 concentration</td></tr><tr><td>freezer box (Black)</td><td>Fisher Scientific</td><td>14100F</td><td>Protects samples from light</td></tr><tr><td>glass pasteur pipettes</td><td>VWR</td><td>14637-010</td><td>Used to transfer samples</td></tr><tr><td>glass slides</td><td>Fisher Scientific</td><td>12550143</td><td>For mounting tissue</td></tr><tr><td>mounting media (vectashield) anti-fade</td><td>VWR</td><td>101098-042</td><td>Mounting media retains fluorescent signaling</td></tr><tr><td>nail polish</td><td>Electron Microscopy Sciences</td><td>72180</td><td>Seals microscope slides</td></tr><tr><td>normal goat serum</td><td>Fisher Scientific</td><td>PCN5000</td><td>Prevents non-specific binding of antibodies</td></tr><tr><td>paint brush</td><td>Genesee Scientific</td><td>59-204</td><td>Transferring flies</td></tr><tr><td>PBS</td><td>Fisher Scientific</td><td>10-010-023</td><td>Saline solution for dissecting and staining</td></tr><tr><td>Phalloidin 647</td><td>Abcam</td><td>AB176759</td><td>Stains F-Actin in muscle Tissue. Used at 1:1000 concentration</td></tr><tr><td>plastic petri dish (100 mm)</td><td>VWR</td><td>25373-100</td><td>Dissection dish</td></tr><tr><td>reinforcement labels</td><td>W.B. Mason</td><td>AVE05722</td><td>Provides support for glass coverslip over the mounted tissue</td></tr><tr><td>sharpening block</td><td>Grainger</td><td>1RDF5</td><td>Keeping fine forceps sharp and also dulling separate pair</td></tr><tr><td>slide folder</td><td>VWR</td><td>10126-326</td><td>Sample storage</td></tr><tr><td>standard office scissors</td><td>W.B. Mason</td><td>ACM40618</td><td>Cutting reinforcement labels</td></tr><tr><td>Sylgard 184</td><td>Electron Microscopy Sciences</td><td>24236-10</td><td>Coating for dissection dish</td></tr><tr><td>Triton-X-100</td><td>Electron Microscopy Sciences</td><td>22140</td><td>Helps to permeabilize tissue</td></tr><tr><td>Vannas Disssection Sissors</td><td>Fine Science Tools</td><td>1500-00</td><td>Ued for removing fly legs and making an incision on thorax</td></tr></tbody></table>

visualizing synaptic degeneration in adult drosophila in association with neurodegeneration

Drosophila serves as a useful model for assessing synaptic structure and function associated with neurodegenerative diseases. While much work has focused on neuromuscular junctions (NMJs) in Drosophila larvae, assessing synaptic integrity in adult Drosophila has received much less attention. Here we provide a straightforward method for dissection of the dorsal longitudinal muscles (DLMs), which are required for flight ability. In addition to flight as a behavioral readout, this dissection allows for the both DLM synapses and muscle tissue to be amenable to structural analysis using fluorescently labeled antibodies for synaptic markers or proteins of interest. This protocol allows for the evaluation of the structural integrity of synapses in adult Drosophila during aging to model the progressive, age-dependent nature of most neurodegenerative diseases.

The generation of transgenic flies expressing human Tar-Binding Protein of 43 kDa mutant (TDP-43M337V) is represented by the schematic (Figure 1A). This demonstrates the application of the binary Gal4/UAS system in Drosophila27. The illustration depicts a hemithorax with six muscle fibers, A&#8210;F going from the most dorsal fiber A to the most ventral F (Figure 1B)11,12. To assess synaptic integrity, NMJs were stained with HRP and Phalloidin (Figure 1C&#8210;E). Motor neurons in TDP-43M337V mutants (Figure 1F) have little to no HRP staining by Day 21, while WT (Oregon-R) remains intact (Figure 1C). There are no visible differences in muscle staining (Figure 1D,G). The changes in gross morphology observed in TDP-43M337V mutants demonstrates how synaptic integrity can be implicated in a neurodegenerative disease model of amyotrophic lateral sclerosis (ALS) using the adult DLM model. In addition to structural staining, staining the DLM NMJs can also provide an assessment of synaptic integrity with presynaptic (Figure 2A&#8210;R) and post synaptic (Figure 2S&#8210;X) markers. Together, these results illustrate how this dissection protocol could be applied to studying DLM tissue in neurodegenerative diseases.
One key aspect of this dissection is the application of liquid nitrogen to flash freeze the tissue to make the bisection easier. The utility of the liquid nitrogen is demonstrated in WT flies with liquid nitrogen where muscle tissue has no damage or nicked fibers (Figure 3A&#8210;C). Without liquid nitrogen, the tissue can be more difficult to dissect. For example, following this protocol and skipping the liquid nitrogen flash freezing step allows the tissue to be more susceptible to damage from the dissection tools such as damaged neurons (Figure 3D) or damaged muscle fibers (Figure 3E). The application of liquid nitrogen helps to prevent tissue damage that could occur when working with DLM tissue regardless of the genotype of the specimen (Figure 3C and 3F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61363/61363fig1.jpg" /> 
Figure 1: Progressive denervation of DLM synapses in a Drosophila model of ALS. (A) The generation of ALS transgenic flies expressing a human mutant form of Tar-Binding Protein of 43 kDa (TDP-43) are shown in the schematic. (B) The illustration depicts the shape and orientation of a hemithorax in an adult Drosophila. Using the protocol, we can observe the progressive loss of synaptic integrity of DLM NMJ synapses through structural staining of motor neurons with HRP (green) and muscle tissue with Phalloidin (magenta). Our model depicts the loss of synaptic integrity in an adult model of ALS through the generation of adult flies expressing a mutant from of human TDP-43M337V in motor neurons (Figure 1F&#8210;H) in comparison to WT (Figure 1C&#8210;E) flies in muscle fiber C. Arrows highlight examples of a WT synapse (Figure 1C) and an example of loss of synaptic integrity. Scale bar =20 &#181;m at 63x magnification. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61363/61363fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61363/61363fig2.jpg" /> 
Figure 2: Assessing synaptic integrity using presynaptic markers at adult NMJs. Synaptic integrity can also be assessed using presynaptic and postsynaptic markers in WT flies that are 14 days old in muscle fiber C. The presynaptic markers Synapsin (B), Syntaxin (H), and Bruchpilot (BRP) (N) are co-stained with HRP (A, G, M). The staining depicts the localization of these markers to the presynaptic terminals (C, I, O). At higher magnification, the images illustrate the localization of Synapsin (E), Syntaxin (K), and BRP (Q) with HRP (D, J, and P) in more detail (Figure F, L, and R). We also show a postsynaptic marker Glutamate Receptor III (GluRIII) (T) co-stained with HRP (S). The co-staining demonstrates the utility of these markers (U). At higher magnification the representative images exemplify the localization (X) of GluRIII (W) and HRP (V) to the postsynaptic muscle tissue and the presynaptic terminals, respectively. Scale bar for panels A&#8210;C, G-I, M&#8210;O, S&#8210;U represent 20 &#181;m at 63x magnification. Scale bar for panels D&#8210;F, 2J-2L, 2P-2R, and 2V-2X represent 10 &#181;m at 63x magnification. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61363/61363fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61363/61363fig3.jpg" /> 
Figure 3: Utility of liquid nitrogen for DLM dissections. To demonstrate the utility of liquid nitrogen for the DLM dissections, we show a comparison of day 21 WT flies with and without liquid nitrogen from muscle fiber C. With liquid nitrogen, Phalloidin (B) remains intact and does not compromise the HRP staining (A, C). Without liquid nitrogen, muscle tissue becomes stringy and difficult to bisect (E) and HRP staining (D, F) becomes compromised due to technical error. White arrows show an area of no muscle damage in with liquid nitrogen (B) and damaged muscle tissue (E). Scale bar = 20 &#181;m at 63x magnification. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/61363/61363fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration

The goal of this procedure is to dissect the dorsal longitudinal muscle (DLM) tissue to assess the structural integrity of DLM neuromuscular junctions (NMJs) in neurodegenerative disease models using Drosophila melanogaster.

Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration

Drosophila serves as a useful model for assessing synaptic structure and function associated with neurodegenerative diseases. While much work has focused ...

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6.1K Views.  Lehigh University. The goal of this procedure is to dissect the dorsal longitudinal muscle (DLM) tissue to assess the structural integrity of DLM neuromuscular junctions (NMJs) in neurodegenerative disease models using Drosophila melanogaster.

Video: Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration

Assaying Locomotor, Learning, and Memory Deficits in Drosophila Models of Neurodegeneration

Advances in genetic methods have enabled the study of genes involved in human neurodegenerative diseases using Drosophila as a model system1. Most of these diseases, including Alzheimer's, Parkinson's and Huntington's disease are characterized by age-dependent deterioration in learning and memory functions and movement coordination2. Here we use behavioral assays, including the negative geotaxis assay3 and the aversive phototaxic suppression assay (APS assay)4,5, to show that some of the behavior characteristics associated with human neurodegeneration can be recapitulated in flies. In the negative geotaxis assay, the natural tendency of flies to move against gravity when agitated is utilized to study genes or conditions that may hinder locomotor capacities. In the APS assay, the learning and memory functions are tested in positively-phototactic flies trained to associate light with aversive bitter taste and hence avoid this otherwise natural tendency to move toward light. Testing these trained flies 6 hours post-training is used to assess memory functions. Using these assays, the contribution of any genetic or environmental factors toward developing neurodegeneration can be easily studied in flies.

Assaying Locomotor, Learning, and Memory Deficits in Drosophila Models of Neurodegeneration

Behavior assays for measuring locomotor functions, learning, and memory abilities in Drosophila.

Advances in genetic methods have enabled the study of genes involved in human neurodegenerative diseases using Drosophila as a model system1. Most of ...

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

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Drosophila melanogaster is a valuable model organism to study aging and pathological degenerative processes in the nervous system. The advantages of the fly as an experimental system include its genetic tractability, short life span and the possibility to observe and quantitatively analyze complex behaviors. The expression of disease-linked genes in specific neuronal populations of the Drosophila brain, can be used to model human neurodegenerative diseases such as Parkinson's and Alzheimer's 5.
Dopaminergic (DA) neurons are among the most vulnerable neuronal populations in the aging human brain. In Parkinson's disease (PD), the most common neurodegenerative movement disorder, the accelerated loss of DA neurons leads to a progressive and irreversible decline in locomotor function. In addition to age and exposure to environmental toxins, loss of DA neurons is exacerbated by specific mutations in the coding or promoter regions of several genes. The identification of such PD-associated alleles provides the experimental basis for the use of Drosophila as a model to study neurodegeneration of DA neurons in vivo. For example, the expression of the PD-linked human &alpha;-synuclein gene in Drosophila DA neurons recapitulates some features of the human disease, e.g. progressive loss of DA neurons and declining locomotor function 2. Accordingly, this model has been successfully used to identify potential therapeutic targets in PD 8.
Here we describe two assays that have commonly been used to study age-dependent neurodegeneration of DA neurons in Drosophila: a climbing assay based on the startle-induced negative geotaxis response and tyrosine hydroxylase immunostaining of whole adult brain mounts to monitor the number of DA neurons at different ages. In both cases, in vivo expression of UAS transgenes specifically in DA neurons can be achieved by using a tyrosine hydroxylase (TH) promoter-Gal4 driver line 3, 10.

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Here we describe two assays that have been established to study age-dependent neurodegeneration of dopaminergic (DA) neurons in Drosophila: the climbing/startle-induced negative geotaxis assay which allows to study the functional effects of DA neurons degeneration and the tyrosine hydroxylase immunostaining which is used to identify and count DA neurons in whole brain mounts.

Drosophila melanogaster is a valuable model organism to study aging and pathological degenerative processes in the nervous system. The advantages 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 ...

Quantitative Analysis of Climbing Defects in a Drosophila Model of Neurodegenerative Disorders

Locomotive defects resulting from neurodegenerative disorders can be a late onset symptom of disease, following years of subclinical degeneration, and thus current therapeutic treatment strategies are not curative. Through the use of whole exome sequencing, an increasing number of genes have been identified to play a role in human locomotion. Despite identifying these genes, it is not known how these genes are crucial to normal locomotive functioning. Therefore, a reliable assay, which utilizes model organisms to elucidate the role of these genes in order to identify novel targets of therapeutic interest, is needed more than ever. We have designed a sensitized version of the negative geotaxis assay that allows for the detection of milder defects earlier and has the ability to evaluate these defects over time. The assay is performed in a glass graduated cylinder, which is sealed with a wax barrier film. By increasing the threshold distance to be climbed to 17.5 cm and increasing the experiment duration to 2 min&#160;we have observed a greater sensitivity in detecting mild mobility dysfunctions. The assay is cost effective and does not require extensive training to obtain highly reproducible results. This makes it an excellent technique for screening candidate drugs in Drosophila mutants with locomotion defects.

We present an optimized inexpensive and reliable negative geotaxis assay in Drosophila melanogaster as a model for neurodegenerative disorders. Being more sensitive to mild locomotor defects, this assay will help screen for potential genetic interactions and drug targets.

Locomotive defects resulting from neurodegenerative disorders can be a late onset symptom of disease, following years of subclinical degeneration, and ...

Mass Histology to Quantify Neurodegeneration in Drosophila

Progressive neurodegenerative diseases like Alzheimer&#39;s disease (AD) or Parkinson&#39;s disease (PD) are an increasing threat to human health worldwide. Although mammalian models have provided important insights into the underlying mechanisms of pathogenicity, the complexity of mammalian systems together with their high costs are limiting their use. Therefore, the simple but well-established Drosophila model-system&#160;provides an alternative for investigating the molecular pathways that are affected in these diseases. Besides behavioral deficits, neurodegenerative diseases are characterized by histological phenotypes such as neuronal death and axonopathy. To quantify neuronal degeneration and to determine how it is affected by genetic and environmental factors, we use a histological approach that is based on measuring the vacuoles in adult fly brains. To minimize the effects of systematic error and to directly compare sections from control and experimental flies in one preparation, we use the &#39;collar&#39; method for paraffin sections. Neurodegeneration is then assessed by measuring the size and/or number of vacuoles that have developed in the fly brain. This can either be done by focusing on a specific region of interest or by analyzing the entire brain by obtaining serial sections that span the complete head. Therefore, this method allows one to measure not only severe degeneration but also relatively mild phenotypes that are only detectable in a few sections, as occurs during normal aging.

Mass Histology to Quantify Neurodegeneration in Drosophila

Drosophila is widely used as a model system to study neurodegeneration. This protocol describes a method by which degeneration, as determined by vacuole formation in the brain, can be quantified. It also minimizes effects due to the experimental procedure by processing and sectioning control and experimental flies as one sample.

Progressive neurodegenerative diseases like Alzheimer&#39;s disease (AD) or Parkinson&#39;s disease (PD) are an increasing threat to human health ...

A Simple Neuronal Mechanical Injury Methodology to Study Drosophila Motor Neuron Degeneration

The degeneration of neurons occurs during normal development and in response to injury, stress, and disease. The cellular hallmarks of neuronal degeneration are remarkably similar in humans and invertebrates as are the molecular mechanisms that drive these processes. The fruit fly, Drosophila melanogaster, provides a powerful yet simple genetic model organism to study the cellular complexities of neurodegenerative diseases. In fact, approximately 70% of disease-associated human genes have a Drosophila homolog and a plethora of tools and assays have been described using flies to study human neurodegenerative diseases. More specifically the neuromuscular junction (NMJ) in Drosophila has proven to be an effective system to study neuromuscular diseases because of the ability to analyze the structural connections between the neuron and the muscle. Here, we report on an in vivo motor neuron injury assay in Drosophila, which reproducibly induces neurodegeneration at the NMJ by 24 h. Using this methodology, we have described a temporal sequence of cellular events resulting in motor neuron degeneration. The injury method has diverse applications and has also been utilized to identify specific genes required for neurodegeneration and to dissect transcriptional responses to neuronal injury.

A Simple Neuronal Mechanical Injury Methodology to Study Drosophila Motor Neuron Degeneration

Here we describe a simple and widely accessible method to injure segmental nerves in Drosophila larvae to visualize and quantify neurodegeneration of motor neurons at the neuromuscular junction (NMJ) of third instar larvae.

The degeneration of neurons occurs during normal development and in response to injury, stress, and disease. The cellular hallmarks of neuronal ...

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Axon degeneration is a shared feature in neurodegenerative disease and when nervous systems are challenged by mechanical or chemical forces. However, our understanding of the molecular mechanisms underlying axon degeneration remains limited. Injury-induced axon degeneration serves as a simple model to study how severed axons execute their own disassembly (axon death). Over recent years, an evolutionarily conserved axon death signaling cascade has been identified from flies to mammals, which is required for the separated axon to degenerate after injury. Conversely, attenuated axon death signaling results in morphological and functional preservation of severed axons and their synapses. Here, we present three simple and recently developed protocols that allow for the observation of axonal morphology, or axonal and synaptic function of severed axons that have been cut-off from the neuronal cell body, in the fruit fly Drosophila. Morphology can be observed in the wing, where a partial injury results in axon death side-by-side of uninjured control axons within the same nerve bundle. Alternatively, axonal morphology can also be observed in the brain, where the whole nerve bundle undergoes axon death triggered by antennal ablation. Functional preservation of severed axons and their synapses can be assessed by a simple optogenetic approach coupled with a post-synaptic grooming behavior. We present examples using a highwire loss-of-function mutation and by over-expressing dnmnat, both capable of delaying axon death for weeks to months. Importantly, these protocols can be used beyond injury; they facilitate the characterization of neuronal maintenance factors, axonal transport, and axonal mitochondria.

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Here, we provide protocols to perform three simple injury-induced axon degeneration (axon death) assays in Drosophila melanogaster to evaluate the morphological and functional preservation of severed axons and their synapses.

Axon degeneration is a shared feature in neurodegenerative disease and when nervous systems are challenged by mechanical or chemical forces. However, our ...

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

The molecular and cellular mechanisms underlying neurogenesis in response to disease or injury are not well understood. However, understanding these mechanisms is crucial for developing neural regenerative therapies. Drosophila melanogaster is a leading model for studies of neural development but historically has not been exploited to investigate adult brain regeneration. This is primarily because the adult brain exhibits very low mitotic activity. Nonetheless, penetrating traumatic brain injury (PTBI) to the adult Drosophila central brain triggers the generation of new neurons and new glia. The powerful genetic tools available in Drosophila combined with the simple but rigorous injury protocol described here now make adult Drosophila brain a robust model for neural regeneration research. Provided here are detailed instructions for (1) penetrating injuries to the adult central brain and (2) dissection, immunohistochemistry, and imaging post-injury. These protocols yield highly reproducible results and will facilitate additional studies to dissect mechanisms underlying neural regeneration.

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

This article provides detailed protocols for inflicting Penetrating Traumatic Brain Injury (PTBI) to adult Drosophila and examining the resulting neurogenesis.

The molecular and cellular mechanisms underlying neurogenesis in response to disease or injury are not well understood. However, understanding these ...

Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration

Summary

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Chapters in this video

Assaying Locomotor, Learning, and Memory Deficits in Drosophila Models of Neurodegeneration

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

Assessing Neurodegenerative Phenotypes in Drosophila Dopaminergic Neurons by Climbing Assays and Whole Brain Immunostaining

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Quantitative Analysis of Climbing Defects in a Drosophila Model of Neurodegenerative Disorders

Mass Histology to Quantify Neurodegeneration in Drosophila

A Simple Neuronal Mechanical Injury Methodology to Study Drosophila Motor Neuron Degeneration

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain