We would like to thank the entire Neukomm lab for contributions. This work was supported by a Swiss National Science Foundation (SNSF) Assistant Professor award (grant 176855), the International Foundation for Research in Paraplegia (IRP, grant P180), SNSF Spark (grant 190919) and by support from the University of Lausanne and the Department of Fundamental Neurosciences (&#201;tat de Vaud) to LJN.

1. Observation of Axon Morphology During Axon Death in the PNS
<ol>
	<li>Wing injury: partial injury of axon bundles
	<ol>
		<li>Use 5 virgin females and 5 males from the right genotype (Figure 4A, P0 generation) to perform crosses at room temperature (RT). Pass P0 into new vials every 3&#8211;4 days. Collect freshly eclosed adult progeny (F1 generation) daily and age them for 7&#8211;14 days.</li>
		<li>Anesthetize flies on CO2 pads. Use micro scissors to cut the anterior wing vein roughly in the middle of the wing (Figure 1A). Use one wing for the injury and the other wing as an age-matched uninjured control. Apply one injury per wing, and make sure to get sufficient wings injured (approximately 15 wings). 
		NOTE: The whole wing can be cut through, but it is sufficient to cut only the anterior wing vein. This is the strongest part of the wing.</li>
		<li>Recover the flies in food-containing vials.</li>
	</ol>
	</li>
	<li>Wing dissection and visualization of axons
	<ol>
		<li>Spread 10 &#181;L of halocarbon oil 27 with a pipette along a whole glass slide (Figure 1B).</li>
		<li>Cut off the injured, as well as, the uninjured control wing at desired time points (e.g., 1 or 7 days post injury). Use micro scissors to cut, and tweezers to grab the wing. Mount maximal 4 wings into halocarbon oil 27 (Figure 1B) and cover them with a cover slide.</li>
		<li>Image the wing immediately using a spinning disk microscope. Acquire a series of optical sections along the z-axis with 0.33 &#956;m step-size and compress z-stacks into a single file for subsequent analyses. 
		NOTE: Do not grab the anterior wing vein where cell bodies and axons are housed. Grab the wing at the center. The tissue in wings is not fixed; keep the time from mounting wings to imaging these under 8 min.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60865/60865fig01.jpg" /> 
Figure 1: Observation of axon morphology during axon death in the wing. (A) Schematic fly wing with two sparsely GFP-labeled sensory neurons, which are also separately indicated below. The site of injury and the field of observation are indicated. (B) Schematic setup for wing imaging. Injured and uninjured control wings (grey) are mounted in halocarbon oil 27 (red) on a glass slide (light blue) and covered with a cover slide (black). <a href="https://www.jove.com/files/ftp_upload/60865/60865fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Observation of Axon and Synapse Morphology During Axon Death in the CNS
<ol>
	<li>Antennal ablation: injury of whole axon bundles
	<ol>
		<li>Use 5 virgin females and 5 males from the right genotype (Figure 5A, P0 generation) to perform crosses at RT. Pass P0 into new vials every 3&#8211;4 days. Collect freshly eclosed adult progeny (F1 generation) daily and let them age for 7 up to 14 days.</li>
		<li>Anesthetize flies on CO2 pads. Use tweezers to ablate the right 3rd antennal segment for unilateral ablation; or both left and right 3rd antennal segments for bilateral ablation (Figure 2A-C). This will remove GFP-labeled neuronal cell bodies, while their axonal projections remain in the CNS. 
		NOTE: Antennal ablation severs the whole axon bundle. If unilateral ablation is performed, the axon bundle on the contralateral side (the unablated antenna) serves as internal control. Make sure to perform sufficient antennal ablations (approximately 15 animals).</li>
		<li>Recover the flies in food-containing vials.</li>
	</ol>
	</li>
	<li>Brain dissection and visualization of axons
	<ol>
		<li>Mix silicone elastomer base (9 mL) and curing agent (1 mL) in a volume ratio of 10:1. Transfer each 5 mL mixture into a 35 mm tissue culture plate, and reduce air introduced by mixing with gentle agitation in the fume hood overnight. The mixture solidifies within 24 h. 
		NOTE: Dissection plates must be prepared only once and may be used multiple times.</li>
		<li>Anesthetize flies on CO2 pads and decapitate adult heads using two tweezers at desired time points (e.g., 1 or 7 days after antennal ablation). Use one tweezer to grab the neck, and the other tweezer to fix the thorax. Gently pull the neck and head off the thorax. 
		NOTE: Leave decapitated heads on the CO2 pad until the desired number is achieved, but make sure to proceed to the next step within 30 min.</li>
		<li>Transfer all heads into a 1.5 mL microcentrifuge tube containing 1 mL of fixing solution containing 4% paraformaldehyde (PFA) and 0.1% Triton X-100 in phosphate buffered saline (PBS) using tweezers that have been dipped into the fixing solution. 
		NOTE: Fly heads stick well on wet tweezers. It makes it feasible to transfer all heads readily into the microcentrifuge tube.</li>
		<li>Fix heads for 20 min with gentle agitation at RT. Put the microcentrifuge tube on ice, heads will gravitate to the bottom of the microcentrifuge tube. Remove the supernatant with a pipette and repeat this procedure with five 2 min washes with 1 mL of washing buffer containing 0.1% Triton X-100 in PBS with gentle agitation at RT, to remove residual fixing solution. 
		NOTE: Videos on how to dissect adult Drosophila brains are readily available27.</li>
		<li>Transfer the heads with a glass pipette into a dissection plate filled with washing buffer. Use one tweezer to grab and pull the proboscis off the head, while holding the head with the other tweezer. This will leave a hole were the proboscis was attached to the exoskeleton.</li>
		<li>Use two tweezers to remove the exoskeleton between the hole and each compound eye. This will make it feasible to open the head structure with both tweezers, and to gently scratch out the brain within.</li>
		<li>Clean each brain by removing trachea or fat stuck to it (Figure 2D, top). Once the brain is cleaned, put it in a new microcentrifuge tube containing 1 mL of washing buffer on ice. 
		NOTE: Damaged or lost optic lobes will not affect the olfactory lobe in the center of the brain (Figure 2D, top).</li>
		<li>Replace washing buffer with 1 mL of fixing solution once all brains are collected and accumulated at the bottom of the microcentrifuge tube. Fix brains for 10 min with rocking at RT, followed by five 2 min washes in 1 mL of washing buffer with rocking at RT.</li>
		<li>Apply primary antibodies (1:500) in washing buffer overnight with rocking at 4 &#176;C, followed by 10 washes over 2 h using 1 mL of washing buffer with rocking at RT.</li>
		<li>Apply secondary antibodies (1:500) in washing buffer 2 h with rocking at RT and wrap microcentrifuge tube in aluminum foil to block light. Keep the microcentrifuge tube covered with aluminum foil for the rest of the procedure. Apply ten washes with 1 mL of washing buffer over 2 h with rocking at RT.</li>
		<li>Remove the supernatant and use a single drop of antifade reagent to cover the brains in the microcentrifuge tube. Incubate brains for at least 30 min at 4 &#176;C before preparing them for mounting and imaging.</li>
		<li>Prepare a cover slide, stick lab tape on it, and cut out a &#34;T&#34;-like shape from the tape (Figure 2D, bottom). The resulting space serves as area where the brain-containing antifade reagent28 will be pipetted into, preferably into both chambers. 
		NOTE: Use a 20-200 &#181;L pipette tip where 3 mm of the tip has been cut off to widen the opening of the pipette. This will make it feasible to pipette the brain-containing antifade reagent. Carefully cover the brains with a cover slide.</li>
		<li>Use clay to prepare two small even rolls. Ensure that the clay rolls are not higher than the height of a glass slide. Stick the clay rolls onto the glass slide (Figure 2D, bottom). Place the brain-containing cover slide sandwich onto the clay rolls. 
		NOTE: GFP-labeled axons and their synapses are located in the front of the brain. It is, therefore, easier to image them from the front. However, brains will either face up, or face down on the cover slide sandwich. Clay rolls serve as sandwich holders, and during imaging, the sandwich can be flipped upside down. This will make it feasible to acquire images from the front from every brain.</li>
		<li>Acquire a series of optical sections along the z-axis with 1.0 &#956;m step-size using a confocal microscope, and compress z-stacks into a single file for subsequent analyses, to assess the number of axonal projections remaining intact.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60865/60865fig02.jpg" /> 
Figure 2: Observation of axon and synapse morphology during axon death in the brain. (A) Side view of a schematic fly head with GFP-labeled cell bodies, axons and synapses. (B) High-magnification front view of GPF-labeled olfactory receptor neurons and their axons and synapses. Cell bodies are housed in the 3rd antennal segment, and their axons project into the CNS. Axons form synapses in a glomerulus in the left olfactory lobe, cross the midline and form synapses in the glomerulus on the contralateral olfactory lobe. (C) Examples of fly heads with unilateral antennal ablation. Top: Uninjured control. Middle: Ablation of the 3rd antennal segment. Bottom: Ablation of the 2nd (and thus also 3rd) antennal segment. (D) Brain preparation. Top: Schematic dissected fly brain with indicated olfactory lobes and axonal projections in the field of view. Bottom: Schematic setup for brain imaging. Two clay rolls (green) are mounted onto a glass slide (light blue), they carry a cover slide sandwich, which contains fly brains (grey). Brains are mounted in antifade reagent(purple), surrounded by a lab tape (orange), and covered by two cover slides (black). <a href="https://www.jove.com/files/ftp_upload/60865/60865fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Grooming Induced by Optogenetics as a Readout for Axon and Synapse Function
<ol>
	<li>Optogenetic setup
	<ol>
		<li>Perform the optogenetic experiment in a dark room. Ensure that the setup consists of an 850 nm infrared (IR) LED spotlight to illuminate flies in the dark (Figure 3A), a flashing 660 nm red LED spotlight to activate neurons expressing CsChrimson, and a monochrome camera with a 700 nm longpass filter, which prevents the recording of red light flashes.</li>
		<li>Use a 3D printer to generate a tiny circular behavior chamber with a diameter of 1 cm, cover it with a cover slide, and place an 860 nm emitter coupled to the red LED spotlight next to the chamber (Figure 3B). 
		NOTE: The emitter indicates when the red LED spotlight is on, thus activating the neurons.</li>
		<li>Mount the LED spotlights and the camera on top of the chamber (Figure 3A, C).</li>
		<li>Activate neurons by 10 Hz flashes during 10 s. The duration of activation can be adjusted according to the experimental design.</li>
	</ol>
	</li>
	<li>Preparing flies for optogenetics
	<ol>
		<li>Melt fly food in a microwave. After the food cooled down, before solidification, add 1:100 of 20 mM all trans-retinal in ethanol (EtOH) to a final concentration of 200 &#181;M. Mix well, and pour the food immediately into empty vials. 
		NOTE: Avoid adding all trans-retinal to hot food, this could result in less efficient optogenetics.</li>
		<li>Cover vials containing solidified food with plugs or cotton balls. Wrap vials with aluminum foil. Then, store the food-containing vials in a dark, cold room.</li>
		<li>Use 5 virgin females and 5 males (Figure 6A, generation P0) from the right genotype to perform crosses at RT. Pass P0 into new vials every 3&#8211;4 days. Collect freshly eclosed adult progeny (generation F1) on a daily base and let them age for 7 up to 14 days in aluminum-covered vials containing 200 &#181;M all trans-retinal in fly food.</li>
		<li>Collect flies by tapping them from food-containing vials into an empty vial with no food. Cool the vial down in ice-containing water for approximately 30 s. Flies will fall asleep. Put individual flies rapidly into small chambers covered with a cover slide (Figure 3B). 
		NOTE: As soon as flies warm up, they wake up. It is crucial to quickly spread individual flies into single chambers each. Avoid CO2 pads to anaesthetize flies, this will impact their behavior.</li>
		<li>Perform optogenetics to elicit antennal grooming. Here, the protocol consists of the following intervals: 30 s where the red light is absent, followed by 10 s of red-light exposure at 10 Hz. Repeat this procedure three times in total, followed by an additional 30 s interval where the red light is absent12,29,30. 
		NOTE: This protocol can be adjusted according to the experimental preference.</li>
		<li>Collect individual flies from each chamber on CO2 pads. Subject them to antennal injury. Ablate both the left and the right 2nd antennal segments (Figure 2C). This will remove the cell bodies of Johnston&#39;s organ (JO) neurons, while the axonal projections remain in the CNS. Recover the flies in aluminum-covered vials containing 200 &#181;M all trans-retinal. 
		NOTE: For antennal grooming induced by optogenetics, the sensory neuron cell bodies are housed in the 2nd antennal segment (Figure 2C).</li>
		<li>At corresponding time points (e.g., 7 days post antennal ablation), subject flies to another grooming assay (go back to step 3.2.4).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60865/60865fig03.jpg" /> 
Figure 3: Optogenetic setup to induce grooming as a readout for axon and synapse function. (A) Illustration of assembled components required for optogenetics. Infrared (IR) LED spotlight, camera and red LED spotlight (from left to right, respectively). The components including a detailed description are listed in the Table of Materials. (B) Top view illustration of a behavior chamber including an IR emitter to indicate red LED spotlight activation. (C) Illustration of a single mount setup. A total of three mount setups are required for the two LED spotlights and the camera, respectively. <a href="https://www.jove.com/files/ftp_upload/60865/60865fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+neural+command+circuit+for+grooming+movement+control.">A neural command circuit for grooming movement control.</a> eLife. 4, 08758 (2015).</li><li>Lee, T., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosaic+analysis+with+a+repressible+cell+marker+for+studies+of+gene+function+in+neuronal+morphogenesis.">Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis.</a> Neuron. 22 (3), 451-461 (1999).</li><li>Vosshall, L. B., Wong, A. M., Axel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+olfactory+sensory+map+in+the+fly+brain.">An olfactory sensory map in the fly brain.</a> Cell. 102 (2), 147-159 (2000).</li><li>Hampel, S., McKellar, C. E., Simpson, J. H., Seeds, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+activation+of+parallel+sensory+pathways+promotes+a+grooming+sequence+in+Drosophila.">Simultaneous activation of parallel sensory pathways promotes a grooming sequence in Drosophila.</a> eLife. 6, (2017).</li><li>Farley, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcription+factor+Pebbled/RREB1+regulates+injury-induced+axon+degeneration.">Transcription factor Pebbled/RREB1 regulates injury-induced axon degeneration.</a> Proceedings of the National Academy of Sciences of the United States of America. 23 (6), (2018).</li><li>Wang, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+depletion+of+ESCRT+protein+Vps4+underlies+injury-induced+autophagic+impediment+and+Wallerian+degeneration.">Rapid depletion of ESCRT protein Vps4 underlies injury-induced autophagic impediment and Wallerian degeneration.</a> Science Advances. 5 (2), 4971 (2019).</li><li>Dietzl, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+genome-wide+transgenic+RNAi+library+for+conditional+gene+inactivation+in+Drosophila.">A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila.</a> Nature. 448 (7150), 151-156 (2007).</li><li>Port, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+large-scale+resource+for+tissue-specific+CRISPR+mutagenesis+in+Drosophila.">A large-scale resource for tissue-specific CRISPR mutagenesis in Drosophila.</a> bioRxiv. 102, 636076 (2019).</li><li>Vagnoni, A., Hoffmann, P. C., Bullock, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reducing+Lissencephaly-1+levels+augments+mitochondrial+transport+and+has+a+protective+effect+in+adult+Drosophila+neurons.">Reducing Lissencephaly-1 levels augments mitochondrial transport and has a protective effect in adult Drosophila neurons.</a> Journal of Cell Science. 129 (1), 178-190 (2016).</li><li>Vagnoni, A., Bullock, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+cAMP/PKA/Kinesin-1+Axis+Promotes+the+Axonal+Transport+of+Mitochondria+in+Aging+Drosophila+Neurons.">A cAMP/PKA/Kinesin-1 Axis Promotes the Axonal Transport of Mitochondria in Aging Drosophila Neurons.</a> Current Biology. 28 (8), 1265-1272 (2018).</li><li>Cao, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+imaging+reveals+mitophagy+independence+in+the+maintenance+of+axonal+mitochondria+during+normal+aging.">In vivo imaging reveals mitophagy independence in the maintenance of axonal mitochondria during normal aging.</a> Aging Cell. 16 (5), 1180-1190 (2017).</li><li>Smith, G. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutathione+S-Transferase+Regulates+Mitochondrial+Populations+in+Axons+through+Increased+Glutathione+Oxidation.">Glutathione S-Transferase Regulates Mitochondrial Populations in Axons through Increased Glutathione Oxidation.</a> Neuron. 103 (1), 52-65 (2019).</li></ol>

The authors declare that they have nothing to disclose.

The protocols described here allow for the robust and reproducible observation of morphology as well as function of axons and their synapses separated from their cell bodies in Drosophila. The wing assay facilitates the observation of axon death side-by-side of uninjured control axons in the PNS14, while the antennal assay facilitates the observation of whole nerve bundles of GFP-labeled axons and their synapses, to assess both morphology and function in the brain (CNS)12. There are critical steps and certain advantages for each approach to study morphology that have to be taken into consideration when designing experiments.
To observe axon morphology in the PNS in the wing, experiments can be readily performed, because of the transparency of the wing: it allows to bypass dissection and immunohistochemistry. However, due to the lack of fixation, the wings have to be imaged immediately after mounting14. Currently, two distinct Gal4 drivers are frequently used, either ok371Gal4 or dpr1Gal4, and both references offer distinct approaches to quantify degeneration14,26. Sparse labeling of a few neurons is recommended, by using &#34;Mosaic Analysis with a Repressible Cell Marker (MARCM)&#34;14,31, as the resolution of axonal morphology is unprecedented. In contrast, the observation of synapses is not possible in wings, they are located in the ventral nerve cord inside the thorax of the flies. Furthermore, additional axonal markers cannot be visualized by immunohistochemistry: the waxy cuticle makes it impossible for the diffusion of fixatives and antibodies into the underlying tissue.
To observe axon and synapse morphology in the CNS, brain dissections have to be performed. They offer the advantage of visualizing additional axonal and synaptic markers by the use of immunohistochemistry, and synapses can be observed alongside axons in the same field of view10,13. A large collection of characterized olfactory receptor neuron (ORN) Gal4 drivers is readily available32, and frequently, OR22aGal4 is the driver of choice. For antennal ablation, cell bodies of OR22a neurons are housed in the 3rd segment (Figure 2B). A fluorescence intensity-based quantification is used to quantify the degeneration of either axons or synapses13. Conversely, experiments are time consuming due to brain dissection and antibody staining.
To visualize axonal and synaptic function after axotomy, optogenetics is used to trigger antennal grooming: it serves as a readout for functional preservation of severed axons and their synapses12. The grooming circuit and corresponding sensory, inter- and motorneuron Gal4 drivers have been thoroughly described29,30. GMR60E02Gal4 labels a subset of Johnston&#39;s organ (JO) sensory neurons, which are required and sufficient for grooming29,30. For antennal ablation, cell bodies of JO neurons are housed in the 2nd antennal segment (Figure 2B). An optogenetic setup can readily be built from scratch, or an existing setup adjusted. Importantly, experiments have to be performed in a dark room, and flies thus visualized with an infrared (IR) LED spotlight. When using CsChrimson as a channel, it is crucial to supply the food with all trans-retinal and a red LED spotlight to activate JO neurons29. Alternatively, blue light-sensitive channels and a blue LED spotlight, or the TrpA1 channel and temperature can be used for neuronal activation29,33. The quantification of grooming behavior has already been described12,29.
When these assays are used to specifically study axon death, it is important to note that the phenotype of morphological or functional preservation should be robust over time. There are cases where axon death leads to a consistent yet less pronounced phenotype in morphological preservation34,35, and whether such a phenotype translates into functional preservation remains to be determined.
Axon death phenotypes have also been observed in neurons during development of Drosophila larvae, where nerves were crushed rather than injured11,23. Here, we specifically focused on adult Drosophila neurons which completed development. In this context, the use of RNA interference36, or tissue-specific CRISPR/Cas937 can readily be implemented. Importantly, the above techniques can be used in an axon death independent context: they facilitate the characterization of neuronal maintenance factors38, axonal transport39, age-dependent axonal mitochondria changes40, and morphology of axonal mitochondria41.

The morphological integrity of neurons is essential for sustained nervous system function throughout life. The vast majority of the neuronal volume is taken by axons1,2; thus life-long maintenance of particularly long axons is a major biological and bioenergetic challenge for the nervous system. Multiple axonal-intrinsic and glial-extrinsic support mechanisms have been identified, ensuring life-long axonal survival. Their impairment results in axon degeneration3, which is a common feature of nervous systems being challenged in disease, and by mechanical or chemical forces4,5. However, the underlying molecular mechanisms of axon degeneration remain poorly understood in any context, making the development of efficacious treatments to block axon loss challenging. The development of effective therapies against these neurological conditions is important, as they create an enormous burden in our society6.
Injury-induced axon degeneration serves as a simple model to study how severed axons execute their own disassembly. Discovered by and named after Augustus Waller in 1850, Wallerian degeneration (WD) is an umbrella term that comprises two distinct, molecularly separable processes7. First, after axonal injury, axons separated from their cell bodies actively execute their own self-destruction (axon death) through an evolutionarily conserved axon death signaling cascade within one day after injury8. Second, surrounding glia and specialized phagocytes engage and clear the resulting axonal debris within three to five days. The attenuation of axon death signaling results in severed axons that remain preserved for weeks9,10,11,12, while the attenuation of glial engulfment culminates in axonal debris which persists for weeks in vivo13,14,15.
Research in flies, mice, rats and zebrafish revealed several evolutionarily conserved and essential mediators of axon death signaling8. Axon death mutants contain severed axons and synapses that fail to undergo axon death; they remain morphologically and functionally preserved for weeks, in the absence of cell body support9,10,12,13,16,17,18,19,20,21,22,23. The discovery and characterization of these mediators led to the definition of a molecular pathway executing axon death. Importantly, axon death signaling is activated not only when the axon is cut, crushed or stretched24,25; it also seems to be a contributor in distinct animal models of neurological conditions (e.g., where axons degenerate in an injury-independent manner4, yet with a range of beneficial outcomes4,8). Therefore, understanding how axon death executes axon degeneration after injury might offer insights beyond a simple injury model; it could also provide targets for therapeutic intervention.
The fruit fly Drosophila melanogaster (Drosophila) has proven to be an invaluable system for axon death signaling. Research in the fly revealed four essential evolutionarily conserved axon death genes: highwire (hiw)11,14, dnmnat12,26, dsarm10 and axundead (axed)12. The modification of these mediators &#8212; loss-of-function mutations of hiw, dsarm and axed, and over-expression of dnmnat &#8212; potently blocks axon death for the life span of the fly. While severed wild type axons undergo axon death within 1 day, severed axons and their synapses lacking hiw, dsarm or axed remain not only morphologically, but also functionally preserved for weeks. Whether functional preservation can also be achieved through high levels of dnmnat remains to be determined.
Here, we will present three simple and recently developed protocols to study axon death (e.g., the morphology and function of severed axons and their synapses over time) in the absence of cell body support. We demonstrate how attenuated axon death results in severed axons which are morphologically preserved with a hiw loss-of-function mutation (hiw&#8710;N) and how attenuated axon death results in severed axons and synapses that remain functionally preserved for at least 7 days with over-expression of dnmnat (dnmnatOE). These protocols allow for the observation of individual axonal and synaptic morphology either in the central, or peripheral nervous system (CNS and PNS, respectively)13,14, while the functional preservation of severed axons and their synapses in the CNS can be visualized by the use of a simple optogenetic setup combined with grooming as a behavioral readout12.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Tweezers (high precision, ultra fine)</td>
			<td>EMS</td>
			<td>78520-5</td>
			<td>Antennal ablation</td>
		</tr>
		<tr>
			<td>MicroPoint Scissors (5-mm cutting edge)</td>
			<td>EMS</td>
			<td>72933-04</td>
			<td>Wing injury</td>
		</tr>
		<tr>
			<td>1.5 mL microcentrifuge tube</td>
			<td>Eppendorf</td>
			<td>30120086.0000</td>
			<td></td>
		</tr>
		<tr>
			<td>35mm tissue culture dish</td>
			<td>Sarstedt</td>
			<td>83.3900</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover Slips, Thickness 1</td>
			<td>Thermo Scientific&#8482;</td>
			<td>BB02400600A113MNT0</td>
			<td></td>
		</tr>
		<tr>
			<td>Superfrost Microscope Slides</td>
			<td>Thermo Scientific&#8482;</td>
			<td>AA00008032E00MNT10</td>
			<td></td>
		</tr>
		<tr>
			<td>High-Sensitivity USB 2.0 CMOS Camera, 1280 x 1024, Global Shutter</td>
			<td>Thorlabs</td>
			<td>DCC1240M</td>
			<td>Camera setup</td>
		</tr>
		<tr>
			<td>SM1 Retaining Ring for &#216;1&#34; Lens Tubes and Mounts</td>
			<td>Thorlabs</td>
			<td>SM1RR</td>
			<td></td>
		</tr>
		<tr>
			<td>25mm 1/1.2&#34; C mount Lens</td>
			<td>Tamron</td>
			<td>M112FM25</td>
			<td></td>
		</tr>
		<tr>
			<td>Adapter with External M27 x 0.5 Threads and Internal SM1 Threads</td>
			<td>Thorlabs</td>
			<td>SM1A36</td>
			<td></td>
		</tr>
		<tr>
			<td>Aspheric Condenser Lens, &#216;25 mm, f=20.1 mm, NA=0.60, ARC: 650-1050 nm</td>
			<td>Thorlabs</td>
			<td>ACL2520U-B</td>
			<td></td>
		</tr>
		<tr>
			<td>&#216;25.0 mm Premium Longpass Filter, Cut-On Wavelength: 700 nm</td>
			<td>Thorlabs</td>
			<td>FELH0700</td>
			<td></td>
		</tr>
		<tr>
			<td>SM1 (1.035&#34;-40) Coupler, External Threads, 0.5&#34; Long</td>
			<td>Thorlabs</td>
			<td>SM1T2</td>
			<td></td>
		</tr>
		<tr>
			<td>SM1 Lens Tube Without External Threads, 1&#34; Long, Two Retaining Rings Included</td>
			<td>Thorlabs</td>
			<td>SM1M10</td>
			<td></td>
		</tr>
		<tr>
			<td>850 nm, 900 mW (Min) Mounted LED, 1200 mA</td>
			<td>Thorlabs</td>
			<td>M850L3</td>
			<td>IR LED spotlight</td>
		</tr>
		<tr>
			<td>SM1 (1.035&#34;-40) Coupler, External Threads, 0.5&#34; Long</td>
			<td>Thorlabs</td>
			<td>SM1T2</td>
			<td></td>
		</tr>
		<tr>
			<td>SM1 Lens Tube Without External Threads, 2&#34; Long, Two Retaining Rings Included</td>
			<td>Thorlabs</td>
			<td>SM1M20</td>
			<td></td>
		</tr>
		<tr>
			<td>Aspheric Condenser Lens, &#216;25 mm, f=20.1 mm, NA=0.60, ARC: 650-1050 nm</td>
			<td>Thorlabs</td>
			<td>ACL2520U-B</td>
			<td></td>
		</tr>
		<tr>
			<td>&#216;25.0 mm Premium Longpass Filter, Cut-On Wavelength: 850 nm</td>
			<td>Thorlabs</td>
			<td>FELH0850</td>
			<td></td>
		</tr>
		<tr>
			<td>SM1 Retaining Ring for &#216;1&#34; Lens Tubes and Mounts</td>
			<td>Thorlabs</td>
			<td>SM1RR</td>
			<td></td>
		</tr>
		<tr>
			<td>660 nm, 940 mW (Min) Mounted LED, 1200 mA</td>
			<td>Thorlabs</td>
			<td>M660L4</td>
			<td>Red LED spotlight</td>
		</tr>
		<tr>
			<td>Aspheric Condenser Lens, &#216;25 mm, f=20.1 mm, NA=0.60, ARC: 650-1050 nm</td>
			<td>Thorlabs</td>
			<td>ACL2520U-B</td>
			<td></td>
		</tr>
		<tr>
			<td>SM1 (1.035&#34;-40) Coupler, External Threads, 0.5&#34; Long</td>
			<td>Thorlabs</td>
			<td>SM1T2</td>
			<td></td>
		</tr>
		<tr>
			<td>SM1 Lens Tube Without External Threads, 2&#34; Long, Two Retaining Rings Included</td>
			<td>Thorlabs</td>
			<td>SM1M20</td>
			<td></td>
		</tr>
		<tr>
			<td>15 V, 2.4 A Power Supply Unit with 3.5 mm Jack Connector for One K- or T-Cube</td>
			<td>Thorlabs</td>
			<td>KPS101</td>
			<td>LED control</td>
		</tr>
		<tr>
			<td>T-Cube LED Driver, 1200 mA Max Drive Current</td>
			<td>Thorlabs</td>
			<td>LEDD1B</td>
			<td></td>
		</tr>
		<tr>
			<td>150 mm x 300 mm x 12.7 mm Aluminum Breadboard, M6 Double-Density Taps</td>
			<td>Thorlabs</td>
			<td>MB1530/M</td>
			<td>Mount base</td>
		</tr>
		<tr>
			<td>&#216;12.7 mm Universal Post Holder, Spring-Loaded Locking Thumbscrew, L = 75 mm</td>
			<td>Thorlabs</td>
			<td>UPH75/M</td>
			<td>Mount, 3x (IR LED, red LED, cam)</td>
		</tr>
		<tr>
			<td>&#216;1.20&#34; Slip Ring for SM1 Lens Tubes and C-Mount Extension Tubes, M4 Tap</td>
			<td>Thorlabs</td>
			<td>SM1RC/M</td>
			<td></td>
		</tr>
		<tr>
			<td>&#216;12.7 mm Optical Post, SS, M4 Setscrew, M6 Tap, L = 150 mm</td>
			<td>Thorlabs</td>
			<td>TR150/M</td>
			<td></td>
		</tr>
		<tr>
			<td>&#216;12.7 mm Optical Post, SS, M4 Setscrew, M6 Tap, L = 40 mm</td>
			<td>Thorlabs</td>
			<td>TR40/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Right-Angle Clamp for &#216;1/2&#34; Posts, 5 mm Hex</td>
			<td>Thorlabs</td>
			<td>RA90/M</td>
			<td></td>
		</tr>
		<tr>
			<td>M6 x 1.0 Stainless Steel Cap Screw, 16 mm Long, Pack of 25</td>
			<td>Thorlabs</td>
			<td>SH6MS16</td>
			<td>screws for mount onto base</td>
		</tr>
		<tr>
			<td>USB-6001 14-Bit 20 kS/s Multifunction I/O and NI-DAQmx</td>
			<td>National Instruments</td>
			<td>782604-01</td>
			<td>Red LED spotlight controller</td>
		</tr>
		<tr>
			<td>20k Ohm 1 Gang Linear Panel Mount Potentiometer</td>
			<td>TT Electronics/BI</td>
			<td>P230-2EC22BR20K</td>
			<td>fintuner for indicator</td>
		</tr>
		<tr>
			<td>IR (860nm) emitter, 100 mA radial</td>
			<td>Osram</td>
			<td>475-1365-ND</td>
			<td>Red light indicator</td>
		</tr>
		<tr>
			<td>cable</td>
			<td>-</td>
			<td>-</td>
			<td>Misc</td>
		</tr>
		<tr>
			<td>All-trans retinal</td>
			<td>Sigma</td>
			<td>R2625</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol absolute</td>
			<td>Vwr</td>
			<td>20821.296</td>
			<td></td>
		</tr>
		<tr>
			<td>Halocarbon Oil 27</td>
			<td>Sigma</td>
			<td>H8773</td>
			<td></td>
		</tr>
		<tr>
			<td>Mowiol</td>
			<td>Merk</td>
			<td>81381</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma</td>
			<td>F8775</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS)</td>
			<td>Sigma</td>
			<td>P5493</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184 silicone elastomer base</td>
			<td>Dow Corning Corp</td>
			<td>4019862</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184 silicone elastomer curing agent</td>
			<td>Dow Corning Corp</td>
			<td>4019862</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
			<td></td>
		</tr>
		<tr>
			<td>Chicken anti-GFP antibodies</td>
			<td>Rockland</td>
			<td>600-901-215</td>
			<td>Antibodies</td>
		</tr>
		<tr>
			<td>Goat Dylight anti-Chicken</td>
			<td>Abcam</td>
			<td>ab96947</td>
			<td></td>
		</tr>
		<tr>
			<td>FM7a, B</td>
			<td>BDSC</td>
			<td>RRID:BDSC_785</td>
			<td>X chromosome</td>
		</tr>
		<tr>
			<td>FRT19A[hs-neo]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_1709</td>
			<td></td>
		</tr>
		<tr>
			<td>hiw[&#916;N]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_51637</td>
			<td></td>
		</tr>
		<tr>
			<td>hs-FLP[12]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_1929</td>
			<td></td>
		</tr>
		<tr>
			<td>tub-Gal80[LL1]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_5132</td>
			<td></td>
		</tr>
		<tr>
			<td>w[1118]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_3605</td>
			<td></td>
		</tr>
		<tr>
			<td>20xUAS-IVS-CsChrimson::mVenus</td>
			<td>BDSC</td>
			<td>RRID:BDSC_55135</td>
			<td>2nd chromosome</td>
		</tr>
		<tr>
			<td>5xUAS-Gal4[12B]</td>
			<td>Kyoto</td>
			<td>RRID:Kyoto_108492</td>
			<td></td>
		</tr>
		<tr>
			<td>5xUAS-HA::dnmnat</td>
			<td>BDSC</td>
			<td>RRID:BDSC_39702</td>
			<td></td>
		</tr>
		<tr>
			<td>5xUAS-mCD8::GFP[LL5]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_5134</td>
			<td></td>
		</tr>
		<tr>
			<td>ase-FLP[2d]</td>
			<td>Freeman laboratory</td>
			<td>Neukomm et al., 2014 (PNAS)</td>
			<td></td>
		</tr>
		<tr>
			<td>CyO</td>
			<td>BDSC</td>
			<td>RRID:BDSC_2555</td>
			<td></td>
		</tr>
		<tr>
			<td>dpr1-Gal4</td>
			<td>BDSC</td>
			<td>RRID:BDSC_25083</td>
			<td></td>
		</tr>
		<tr>
			<td>OR22a-Gal4</td>
			<td>BDSC</td>
			<td>RRID:BDSC_9952</td>
			<td></td>
		</tr>
		<tr>
			<td>ey-FLP[6]</td>
			<td>BDSC</td>
			<td>RRID:BDSC_5577</td>
			<td>3rd chromosome</td>
		</tr>
		<tr>
			<td>GMR60E02-Gal4</td>
			<td>BDSC</td>
			<td>RRID:BDSC_39250</td>
			<td></td>
		</tr>
		<tr>
			<td>TM3,Sb,e</td>
			<td>BDSC</td>
			<td>RRID:BDSC_3644</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Above, we described three methods to study the morphology and function of severed axons and their synapses. The first method allows for high-resolution observation of individual axons in the PNS. It requires clones generated by the MARCM technique14,31. Here, we performed crosses to generate wild type and highwire mutant MARCM clones (Figure 4A). A simple cut in the middle of the wing induces axon injury of neurons housed distal (e.g., at the outer side of the wing), while proximal neurons (e.g., between the cut site and the thorax) remain uninjured. This approach makes it feasible to observe axon death side-by-side of uninjured control axons in the same nerve bundle (Figure 1A, Figure 4B). Here, we used a genetic background resulting in low numbers of GFP-labeled clones (e.g., two in each experiment14). We present examples of 1 and 7 days after injury of wild type axons, to provide examples of control axons, axons undergoing axon death, and axonal fragments being cleared by surrounding glia, respectively. In addition, we repeated axonal injury in highwire mutants where we analyzed the outcome 7 days after injury.
Uninjured control wings harbor two wild-type clones, thus two GFP-labeled wild-type axons (Figure 4B, wild type, uninjured control). One day after cutting the middle of the wing by the use of micro scissors, axon death is induced in GFP-labeled axons where cell bodies are distal to the cut site, while axons from proximally housed cell bodies serve as an internal control within the same nerve bundle (Figure 4B, wild type, 1 day post injury). Note the axonal debris trace in the upper part indicated by the arrow. 7 days after axonal injury, GFP-labeled axonal debris is cleared by surrounding glia, while GFP labeled uninjured control axons remain in the nerve bundle (Figure 4B, wild type, 7 days post injury, arrow). In contrast, highwire mutant axons that have been severed for 7 days remain morphologically preserved, consistent with previous findings11,14 (Figure 4B, highwire, 7 days post injury, arrow). These results demonstrate the powerful visual resolution of the Drosophila wing. Axon death can be observed side-by-side of uninjured controls in the same nerve bundle. While wild-type axons undergo axon death within 1 day after injury and the resulting debris is cleared within 7 days, axon death deficient highwire mutants remain morphologically preserved for 7 days.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60865/60865fig04.jpg" /> 
Figure 4: Approach to study axon death of GFP-labeled sensory neuron axons in the wing. (A) Schematic crosses to generate wild type and highwire clones in the wing (P0 and F1 generation, respectively). Virgin females are on the left, males on the right. See Table of Materials for genotype details. (B) Examples of control and injured GFP-labeled axons. The field of view is indicated in (Figure 1A). From left to right: uninjured wild type control axons, wild type axons 1-day post injury, wild type axons 7 days post injury, highwire mutant axons 7 days post injury, respectively. Arrows indicate severed axons, Scale bar = 5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/60865/60865fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The second method describes how to visualize whole axon bundles projecting into the CNS where they form synapses, which belong to neurons housed in both left and right antennae (Figure 2A-C). Here, we performed crosses to generate wild type and highwire mutant MARCM clones (Figure 5A). Uninjured, GFP-labeled axons and their synapses can be visualized over the course of days to weeks, in the absence of injury (Figure 5B, Wild type, uninjured control). Alternatively, animals can be subjected to 3rd antennal segment ablation, and severed GFP-labeled axons and their synapses can be observed during a time course over hours to days. We focused on 7 days after antennal ablation, because at this time point, axons and their synapses have undergone axon death, and resulting debris is cleared by surrounding glia. If unilateral ablation of the right antenna is performed, then the right axon bundle is severed and will disassemble and the resulting debris is fully cleared 7 days after injury (Figure 5B, wild type, unilateral ablation, 7 days post injury, arrows), consistent with previous findings13. Alternatively, both the right and the left antennae can be ablated, which will sever both axon bundles, and 7 days after injury, axons and their synapses disappeared (Figure 5B, wild type, bilateral ablation, 7 days post injury, arrow). In contrast, unilateral ablation of the right antennae in highwire mutants results in severed axons that remain preserved 7 days post injury, consistent with previous findings11,14 (Figure 5B, highwire, unilateral ablation, 7 days post injury, arrow). These results demonstrate that severed wild-type axons undergo axon death and the resulting debris is cleared within 7 days, while axon death deficient highwire mutants fail to undergo axon death and remain morphologically preserved for 7 days.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60865/60865fig05.jpg" /> 
Figure 5: Approach to study axon death of GFP-labeled sensory neuron axons in the brain. (A) Schematic crosses to generate wild type and highwire clones in the brain (P0 and F1 generation, respectively). Virgin females are on the left, males on the right. See Table of Materials for genotype details. (B) Examples of control and injured GFP-labeled axons. From left to right: uninjured wild type controls, wild type 7 days post unilateral antennal ablation, wild type 7 days post bilateral antennal ablation, and highwire mutants 7 days post unilateral antennal ablation, respectively. Arrows indicate severed axon bundles, Scale bar = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/60865/60865fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The third method allows for the observation of functional preservation of severed axons and their synapses in the CNS. It relies on the manipulation of a subset of JO neurons housed in the 2nd antennal segment which are sufficient to induce antennal grooming. Expression of a red-shifted channelrhodopsin (CsChrimson) in JO neurons, combined with dietary supplementation of all trans-retinal, is sufficient to elicit a simple post-synaptic grooming behavior upon red light exposure12,30. Here, we performed crosses to generate wild type JO neurons, and JO neurons over-expressing dnmnat (dnmnatOE) (Figure 6A). Wild type flies or flies containing JO neurons with attenuated axon death (dnmnatOE), both harbor a potent grooming behavior before injury. However, 7 days post injury (e.g., bilateral ablation of the 2nd antennal segment), grooming fails to be elicited by optogenetics in wild type flies due to injury-induced axon and synapse degeneration, while animals with attenuated axon death continue to groom (Figure 6B, Movie 1,2). Attenuated axon death is therefore capable of functionally preserving severed axons and their synapses for 7 days.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60865/60865fig06.jpg" /> 
Figure 6: Approach to visualize axonal and synaptic function after axotomy. (A) Schematic crosses to generate wild type and dnmnat over-expressing JO sensory neurons (P0 and F1 generation, respectively). Virgin females are on the left, males on the right. See Table of Materials for genotype details. (B) Individual ethograms of grooming behavior induced by optogenetics. Top: individual ethograms of wild type flies before and 7 days after injury (blue). Bottom: individual ethograms of flies over-expressing dnmnat (dnmnatOE) in JO neurons before and 7 days after injury (red). Each bin indicates at least 1 grooming behavior within 1 s. The black line indicates the sum of all bins. (C) Quantification of grooming behavior. Data is shown as average &#177; standard deviation, p &#62; 0.001 (one-way ANOVA, multiple comparison with Tukey&#39;s post hoc test). <a href="https://www.jove.com/files/ftp_upload/60865/60865fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Movie 1: Representative wild type grooming behavior elicited by optogenetics before and 7 days after antennal ablation. <a href="https://cloudflare.jove.com/files/ftp_upload/60865/Movie_1_wild_type.mp4" target="_blank">Please click here to download this video.</a>
Movie 2: Representative grooming behavior elicited by optogenetics in flies over-expressing dnmnat in JO neurons before and 7 days after antennal ablation. <a href="https://cloudflare.jove.com/files/ftp_upload/60865/Movie_2_nmnat.mp4" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about Morphological and Functional Evaluation of Axons and their Synapses during Axon Death in Drosophila melanogaster at JoVE.com

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.

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 ...

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7.7K Views.  University of Lausanne. 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.

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

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

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

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

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

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

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

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such behavior is spontaneous locomotor activity. Here we describe our protocol that utilizes Drosophila population monitors and a tracking system that allows continuous monitoring of the spontaneous locomotor activity of flies for several days at a time. This method is simple, reliable, and objective and can be used to examine the effects of aging, sex, changes in caloric content of food, addition of drugs, or genetic manipulations that mimic human diseases.

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster are useful in studying genetic or environmental manipulations that affect behaviors such as spontaneous locomotor activity.&#160; Here we describe a protocol that utilizes monitors with infrared beams and data analysis software to quantify spontaneous locomotor activity.&#160;

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such ...

Functional and Morphological Assessment of Diaphragm Innervation by Phrenic Motor Neurons

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and morphological levels. Compound muscle action potential (CMAP) recording following phrenic nerve stimulation can be used to quantitatively assess functional diaphragm innervation by PhMNs of the cervical spinal cord in vivo in anesthetized rats and mice. Because CMAPs represent simultaneous recording of all myofibers of the whole hemi-diaphragm, it is useful to also examine the phenotypes of individual motor axons and myofibers at the diaphragm NMJ in order to track disease- and therapy-relevant morphological changes such as partial and complete denervation, regenerative sprouting and reinnervation. This can be accomplished via whole-mount immunohistochemistry (IHC) of the diaphragm, followed by detailed morphological assessment of individual NMJs throughout the muscle. Combining CMAPs and NMJ analysis provides a powerful approach for quantitatively studying diaphragmatic innervation in rodent models of CNS and PNS disease.

Compound muscle action potential recording quantitatively assesses functional diaphragm innervation by phrenic motor neurons. Whole-mount diaphragm immunohistochemistry assesses morphological innervation at individual neuromuscular junctions. The goal of this protocol is to demonstrate how these two powerful methodologies can be used in various rodent models of spinal cord disease.

This protocol specifically focuses on tools for assessing phrenic motor neuron (PhMN) innervation of the diaphragm at both the electrophysiological and ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

Measuring and Altering Mating Drive in Male Drosophila melanogaster

Despite decades of investigation, the neuronal and molecular bases of motivational states remain mysterious. We have recently developed a novel, reductionist, and scalable system for in-depth investigation of motivation using the mating drive of male Drosophila&#160;melanogaster (Drosophila), the methods for which we detail here. The behavioral paradigm centers on the finding that male mating drive decreases alongside fertility over the course of repeated copulations and recovers over ~3 d. In this system, the powerful neurogenetic tools available in the fly converge with the genetic accessibility and putative wiring diagram available for sexual behavior. This convergence allows rapid isolation and interrogation of small neuronal populations with specific motivational functions. Here we detail the design and execution of the satiety assay that is used to measure and alter courtship motivation in the male fly. Using this assay, we also demonstrate that low male mating drive can be overcome by stimulating dopaminergic neurons. The satiety assay is simple, affordable, and robust to influences of genetic background. We expect the satiety assay to generate many new insights into the neurobiology of motivational states.

Measuring and Altering Mating Drive in Male Drosophila melanogaster

This article describes a behavioral assay that uses male mating drive in Drosophila melanogaster to study motivation. Using this method, researchers can utilize advanced fly neurogenetic techniques to uncover the genetic, molecular, and cellular mechanisms that underlie this motivation.

Despite decades of investigation, the neuronal and molecular bases of motivational states remain mysterious. We have recently developed a novel, ...

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system. They provide multiple experimental advantages, such as accessibility to numerous imaging approaches, electrophysiological techniques and behavioral assays. The Xenopus tadpole olfactory system is particularly well suited to investigate the function of synapses established during normal development or reformed after injury. Here, we describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae. We outline a combination of in vivo measurements of presynaptic calcium responses in glomeruli of the olfactory bulb with olfactory-guided behavior assays. Methods can be combined with the transection of olfactory nerves to study the rewiring of synaptic connectivity. Experiments are presented using both wild-type and genetically modified animals expressing GFP reporters in central nervous system cells. Application of the approaches described to genetically modified tadpoles can be useful for unraveling the molecular bases that define vertebrate behavior.

Functional Evaluation of Olfactory Pathways in Living Xenopus Tadpoles

Xenopus tadpoles offer a unique platform to investigate the function of the nervous system in vivo. We describe methodologies to evaluate the processing of olfactory information in living Xenopus larvae in normal rearing conditions or after injury.