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 border="0" cellpadding="0" cellspacing="0" style="width:725px;" width="724">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="92">
			<td height="92" style="height:92px;width:216px;">32% Formaldehyde</td>
			<td style="width:121px;">Electron Microscopy Sciences</td>
			<td style="width:161px;">15714</td>
			<td style="width:227px;">Tissue preservation</td>
		</tr>
		<tr height="86">
			<td height="86" style="height:87px;width:216px;">Alexa Fluor 568 goat anti mouse</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">A11031</td>
			<td style="width:227px;">Labels primary antibodies. Used at 1:200 concentration.</td>
		</tr>
		<tr height="66">
			<td height="66" style="height:67px;width:216px;">Alexa Fluor 568 goat anti rabbit</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">A11036</td>
			<td style="width:227px;">Labels primary antibodies. Used at 1:200 concentration.</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:216px;">anti- Bruchpilot (BRP) antibody</td>
			<td style="width:121px;">Developmental Studies Hybridoma Bank</td>
			<td>NC82</td>
			<td style="width:227px;">Stains the active zones in presynaptic neurons. Used at 1:25 concentration.</td>
		</tr>
		<tr height="73">
			<td height="73" style="height:73px;width:216px;">anti-GluRIII antibody</td>
			<td style="width:121px;">Gift from Aaron DiAntonio</td>
			<td style="width:161px;">N/A</td>
			<td style="width:227px;">Labels glutamate receptor subunits. Used at 1:1000 concentration.</td>
		</tr>
		<tr height="76">
			<td height="76" style="height:76px;width:216px;">anti-Synapsin antibody</td>
			<td style="width:121px;">Developmental Studies Hybridoma Bank</td>
			<td style="width:161px;">3C11</td>
			<td style="width:227px;">Labels the synaptic protein synapsin. Used at 1:50 concentration.</td>
		</tr>
		<tr height="46">
			<td height="46" style="height:47px;width:216px;">anti-Syntaxin antibody</td>
			<td style="width:121px;">Developmental Studies Hybridoma Bank</td>
			<td style="width:161px;">8C3</td>
			<td style="width:227px;">labels the synaptic protein syntaxin. Used at 1:10 concentration.</td>
		</tr>
		<tr height="44">
			<td height="44" style="height:44px;width:216px;">BenchRocker</td>
			<td style="width:121px;">Genesee Scientific</td>
			<td style="width:161px;">31-302</td>
			<td style="width:227px;">Rotating samples during staining</td>
		</tr>
		<tr height="93">
			<td height="93" style="height:94px;width:216px;">Blade Breaker</td>
			<td style="width:121px;">Fine Science Tools</td>
			<td style="width:161px;">10053-09</td>
			<td style="width:227px;">Used for holding feather blade</td>
		</tr>
		<tr height="76">
			<td height="76" style="height:76px;width:216px;">cover slips</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">12548A</td>
			<td style="width:227px;">For mounting tissue</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:216px;">cryogenic gloves</td>
			<td style="width:121px;">VWR</td>
			<td>97008-198</td>
			<td style="width:227px;">protect hands from liquid nitrogen</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:216px;">cryogenic tweezers</td>
			<td style="width:121px;">VWR</td>
			<td>82027-432</td>
			<td style="width:227px;">Hold 2.0 mL tube in liquid nitrogen</td>
		</tr>
		<tr height="56">
			<td height="56" style="height:56px;width:216px;">dewar flask-1900 mL</td>
			<td style="width:121px;">Thomas Scientific</td>
			<td>5028M54</td>
			<td style="width:227px;">Hold liquid nitrogen</td>
		</tr>
		<tr height="62">
			<td height="62" style="height:63px;width:216px;">Feather Blades</td>
			<td style="width:121px;">Electron Microscopy Sciences</td>
			<td style="width:161px;">72002-01</td>
			<td style="width:227px;">Scalpel Blades</td>
		</tr>
		<tr height="53">
			<td height="53" style="height:54px;width:216px;">Fine Forecps x 2</td>
			<td style="width:121px;">Fine Science Tools</td>
			<td style="width:161px;">11252-20</td>
			<td style="width:227px;">One fine pair for Clearing midline of thorax. The other pair can be dulled using a sharpening stone.</td>
		</tr>
		<tr height="58">
			<td height="58" style="height:59px;width:216px;">FITC-conjugated anti HRP</td>
			<td style="width:121px;">Jackson Laboratories</td>
			<td style="width:161px;">123-545-021</td>
			<td style="width:227px;">Stains Motor Neurons. Used at 1:100 concentration</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:216px;">freezer box (Black)</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">14100F</td>
			<td style="width:227px;">Protects samples from light</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:216px;">glass pasteur pipettes</td>
			<td style="width:121px;">VWR</td>
			<td style="width:161px;">14637-010</td>
			<td style="width:227px;">Used to transfer samples</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">glass slides</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">12550143</td>
			<td style="width:227px;">For mounting tissue</td>
		</tr>
		<tr height="86">
			<td height="86" style="height:87px;width:216px;">mounting media (vectashield) anti-fade</td>
			<td style="width:121px;">VWR</td>
			<td style="width:161px;">101098-042</td>
			<td style="width:227px;">Mounting media retains fluorescent signaling</td>
		</tr>
		<tr height="53">
			<td height="53" style="height:54px;width:216px;">nail polish</td>
			<td style="width:121px;">Electron Microscopy Sciences</td>
			<td style="width:161px;">72180</td>
			<td style="width:227px;">Seals microscope slides</td>
		</tr>
		<tr height="49">
			<td height="49" style="height:49px;width:216px;">normal goat serum</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">PCN5000</td>
			<td style="width:227px;">Prevents non-specific binding of antibodies</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:216px;">paint brush</td>
			<td style="width:121px;">Genesee Scientific</td>
			<td style="width:161px;">59-204</td>
			<td style="width:227px;">Transferring flies</td>
		</tr>
		<tr height="53">
			<td height="53" style="height:54px;width:216px;">PBS</td>
			<td style="width:121px;">Fisher Scientific</td>
			<td style="width:161px;">10-010-023</td>
			<td style="width:227px;">Saline solution for dissecting and staining</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">Phalloidin 647</td>
			<td style="width:121px;">Abcam</td>
			<td>AB176759</td>
			<td style="width:227px;">Stains F-Actin in muscle Tissue. Used at 1:1000 concentration</td>
		</tr>
		<tr height="66">
			<td height="66" style="height:67px;width:216px;">plastic petri dish (100 mm)</td>
			<td style="width:121px;">VWR</td>
			<td style="width:161px;">25373-100</td>
			<td style="width:227px;">Dissection dish</td>
		</tr>
		<tr height="57">
			<td height="57" style="height:58px;width:216px;">reinforcement labels</td>
			<td style="width:121px;">W.B. Mason</td>
			<td>AVE05722</td>
			<td style="width:227px;">Provides support for glass coverslip over the mounted tissue</td>
		</tr>
		<tr height="92">
			<td height="92" style="height:92px;width:216px;">sharpening block</td>
			<td style="width:121px;">Grainger</td>
			<td style="width:161px;">1RDF5</td>
			<td style="width:227px;">Keeping fine forceps sharp and also dulling separate pair</td>
		</tr>
		<tr height="68">
			<td height="68" style="height:68px;width:216px;">slide folder</td>
			<td style="width:121px;">VWR</td>
			<td style="width:161px;">10126-326</td>
			<td style="width:227px;">Sample storage</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">standard office scissors</td>
			<td style="width:121px;">W.B. Mason</td>
			<td>ACM40618</td>
			<td style="width:227px;">Cutting reinforcement labels</td>
		</tr>
		<tr height="56">
			<td height="56" style="height:56px;width:216px;">Sylgard 184</td>
			<td style="width:121px;">Electron Microscopy Sciences</td>
			<td style="width:161px;">24236-10</td>
			<td style="width:227px;">Coating for dissection dish</td>
		</tr>
		<tr height="76">
			<td height="76" style="height:76px;width:216px;">Triton-X-100</td>
			<td style="width:121px;">Electron Microscopy Sciences</td>
			<td style="width:161px;">22140</td>
			<td style="width:227px;">Helps to permeabilize tissue</td>
		</tr>
		<tr height="56">
			<td height="56" style="height:56px;width:216px;">Vannas Disssection Sissors</td>
			<td style="width:121px;">Fine Science Tools</td>
			<td style="width:161px;">1500-00</td>
			<td style="width:227px;">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://www.jove.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://www.jove.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://www.jove.com/files/ftp_upload/61363/61363fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration at JoVE.com

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 ...

visualizing-synaptic-degeneration-adult-drosophila-association-with

Universidad San Sebastian

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Neuroscience

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

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 ...

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 ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

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 One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

Modeling Amyloid-&#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to synaptic degeneration, inflammation, neuronal death, and learning deficits. Humans cannot regenerate lost neurons in the case of AD in part due to impaired proliferative capacity of the neural stem/progenitor cells (NSPCs) and reduced neurogenesis. Therefore, efficient regenerative therapies should also enhance the proliferation and neurogenic capacity of NSPCs. Zebrafish (Danio rerio) is a regenerative organism, and we can learn the basic molecular programs with which we could design therapeutic approaches to tackle AD. For this reason, the generation of an AD-like model in zebrafish was necessary. Using our methodology, we can introduce synthetic derivatives of A&#946;42 peptide with tissue penetrating capability into the adult zebrafish brain, and analyze the disease pathology and the regenerative response. The advantage over the existing methods or animal models is that zebrafish can teach us how a vertebrate brain can naturally regenerate, and thus help us to treat human neurodegenerative diseases better by targeting endogenous NSPCs. Therefore, the amyloid-toxicity model established in the adult zebrafish brain may open new avenues for research in the field of neuroscience and clinical medicine. Additionally, the simple execution of this method allows for cost-effective and efficient experimental assessment. This manuscript describes the synthesis and injection of A&#946;42 peptides into zebrafish brain.

This protocol describes the synthesis, characterization, and injection of monomeric amyloid-&#946;42 peptides for generating amyloid toxicity in adult zebrafish to establish an Alzheimer&#39;s disease model, followed by histological analyses and detection of aggregations.

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration

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