Experiments on the 2-photon microscope were performed in the FAU Stiles-Nicholson Brain Institute Advanced Cell Imaging Core. We would like to thank the Jupiter Life Science Initiative for financial support.

All animals used for the protocol were of the species Drosophila melanogaster. There are no ethical issues surrounding the use of this species. Ethical clearance was not necessary to carry out this work. The details of the Drosophila species, reagents, and equipment used in the study are listed in the Table of Materials.
1. Breeding Drosophila and selecting the correct larval stage
<ol>
	<li>Choose a Gal4 driver line that drives expression in the cells that are to be ablated and recombine it with or cross it to a UAS-GFP reporter line. Raise flies on standard fly food at 25 &#176;C. 
	NOTE: For giant fiber (GF) ablation, R91H05-Gal4 or A307-Gal4 was recombined with UAS-GFP. ShakB(lethal)-GAL4 recombined with UAS-GFP was used for TTMn ablations (see Table of Materials).</li>
	<li>Select larvae that have started to emerge from the food and crawl up the side of the food vial. Those larvae are in the wandering stage, which is the preferred stage for performing GF cell ablation. 
	NOTE: Ablation can be performed at any larval stage. The last larval stage was important for the manipulation of the development of the GFS because&#160;the GF cell bodies can be easily identified in the brain, but the GF axons have not made connections to their targets at this stage.</li>
</ol>
2. Preparing the 3rd instar larva
NOTE: Larvae were anesthetized using a method similar to Burra et al.9. To keep procedure times short, only prepare one larva at a time. Short exposure to anesthetic will enhance the survival of experimental animals10.
<ol>
	<li>Prepare a glass dish with a tightly fitting lid. Place a cotton ball in the dish, and under a fume hood, add about 3-5 ml of ethyl ether (dangerous, flammable liquid, lab coat, gloves, goggles). Cover the dish with the lid.</li>
	<li>Collect individual 3rd instar larvae from fly vials. At the 3rd larval stage, larvae leave the food medium and crawl up along the side of the food vial11.
	<ol>
		<li>Ensure that larvae are still moving and haven&#39;t started puparium formation. Larvae that are stationary and show a shortening of the body are starting pupation and are unsuitable for ablation. Pick larvae up with a small paintbrush. 
		NOTE: The 3rd larval stage starts after the second molt at 72 h of development (after egg laying), and lasts until puparium formation at 120 h for flies raised at 25 &#176;C.</li>
	</ol>
	</li>
	<li>Transfer one larva into a small, open container. Place the container in the glass dish with ethyl ether and close the dish tightly. When opening the glass dish, place it under a snorkel or in a fume hood to prevent fumes from escaping.
	<ol>
		<li>In intervals of 30 s, remove the lid with the larva from the dish and check the larva for mobility under a dissection microscope. The larva will generally be immobilized within 1 min. Once the mouth hooks stop twitching, the larva is ready for mounting. Discard any larvae that are not fully immobilized after 3 min. 
		NOTE: The top of an upside-down lid of a microcentrifuge tube (lid removed from the tube) can be used as a container.</li>
	</ol>
	</li>
</ol>
3. Mounting larvae on slides for ablation
<ol>
	<li>Place the anesthetized larva on a glass microscope slide. Submerge the larva in a drop of insect saline; this will wash away some of the food debris that might stick to the larva.</li>
	<li>Under the dissection microscope, remove most of the saline with a paper tissue. Position the larva dorsal side up for GF ablation or the ventral side up for TTMn ablation. Slowly lower a glass coverslip onto the larva. Add saline to the side of the coverslip to fill in the space between the glass slide and the cover slip.</li>
	<li>For GF ablation, check the positioning of the brain under high magnification on the dissection microscope. Ensure that the brain is lying level and is visible through the cuticle. Often, the brain will be covered with fat tissue, making visualization and cell ablation impossible.
	<ol>
		<li>To displace any fat tissue covering the brain, apply slight pressure to the coverslip with forceps and move the coverslip from side to side. If the fat tissue can&#39;t be moved away from the brain this way, use a different larva instead.</li>
	</ol>
	</li>
</ol>
4. Locating the target cells
NOTE: The multi-photon system used for this study was mounted on an upright microscope. System-specific software was used to control acquisition and laser stimulation settings. The system was equipped with an epifluorescence light source to locate the samples. The objective lens was a water immersion lens with 25x magnification, a long 2 mm working distance, and an NA of 1.10.
<ol>
	<li>Place the sample on the multi-photon microscope&#39;s stage and use the GFP filter to locate the sample in epifluorescence mode. For GF ablation, the cell bodies can be identified in the brain, as seen in Figure 1A,C,D. For TTMn ablation, a cluster of four cells can be identified from the ventral side in the ventral nerve cord (VNC), as shown in Figure 1A,B. When done focusing, center the cells in the field of view and switch to 2-photon mode.</li>
</ol>
5. Setting up the ablation parameters
<ol>
	<li>Adjust laser settings and detector gain to view the GFP-expressing cells with the Galvano scanner. A wavelength of 870 nm works best for visualizing the GFP signal. Center the cell in the field of view.</li>
	<li>Use a circular region of interest (ROI) to define the area for ablation. Ensure that the ROI covers most of the surface of the cell (Figure 1E,G).</li>
	<li>Set up the ablation protocol in the software. Set one frame of acquisition, then stimulation, followed by another acquisition frame.</li>
	<li>Start the stimulation laser power setting at a lower value (10%-20 %) and run the stimulation protocol. A successful ablation can be recognized by a clear circle in the center of the cell soma at the location of the ROI and an increase in fluorescence surrounding the ROI (Figure 1F).
	<ol>
		<li>If the ablation was not successful and merely bleached the cell (Figure 1H), increase the laser power by increments of 5% or the number of loops one at a time and run the protocol again. The applied laser power was between 10% and 40% for the system used in the protocol. 
		&#8203;NOTE: Laser power settings for ablation depend greatly on how deep the cells are located in the tissue and other factors such as cuticle folding. If cells are bright and clearly visible, laser power will be on the lower end of the range. Laser power will be different for each system depending on the laser model and the age of the laser.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67053/67053fig01.jpg" /> 
Figure 1: Identification of neurons for laser ablation in intact Drosophila L3 larvae. (A) Schematic of the location of giant fiber (GF) soma in the brain and cluster of motor neurons containing the tergotrochanteral motorneuron (TTMn) in the ventral nerve cord (VNC). (B) Maximum intensity projection of shakB(lethal)-Gal4 driving expression of GFP in the larval VNC. The midline is indicated by the dotted line. The circle indicates the cluster of neurons containing the TTMn that are targeted for laser ablation. Scale bar: 50 &#181;m. (C) Partial projection view of the larval brain expressing GFP under the control of A307-Gal4. The circle indicates the GF soma targeted for laser ablation. Scale bar: 50 &#181;m. (D) Maximum intensity projection of the brain with R91H05-Gal4 driving UAS-GFP. Both GFs (circles) are identifiable by location, shape, and size. Scale bar: 50 &#181;m. (E) GF soma enlarged view before laser ablation. The magenta circle indicates the target region for laser ablation. Scale bar: 10 &#181;m. (F) GF soma enlarged view after delivery of localized laser power and successful ablation. Scale bar: 10 &#181;m. (G) GF soma enlarged view before laser ablation. The magenta circle indicates the target region for laser ablation. Scale bar: 10 &#181;m. (H) GF soma enlarged view after delivery of localized laser power and unsuccessful ablation (bleaching). Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/67053/67053fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Recovering the larvae
<ol>
	<li>After successfully ablating the cell, remove the coverslip and gently pick up the larvae from the slide with a paintbrush. Place the larvae in a food vial. Larvae will start crawling around again within 30 min. 
	NOTE: Keeping procedure time short improves the survival rates of animals. With experience, the experiment can be performed in just under 10 min.</li>
	<li>Continue raising larvae at standard conditions until they eclose from the pupal casing.</li>
</ol>
7. Testing the functionality of the GFS in adult flies
NOTE: The following steps are explained in detail in Allan and Godenschwege12, and Augustin et al.13.
<ol>
	<li>Test adult flies at age 2-5 days after eclosion. Anesthetize flies with CO2 and mount them on dental wax by pushing the legs into the wax. Spread the wings at a 90&#176; angle and fix them in place with the wax. Secure the head by pushing the proboscis into the wax.</li>
	<li>Place stimulation wires into both eyes, a ground wire into the abdomen, and glass recording electrodes into the jump muscles (TTMs) on each side.</li>
	<li>Stimulate the GF circuit with single stimuli to measure response latency for both muscles. Stimulate the circuit with 100 Hz and 200 Hz trains of stimuli, respectively, to measure the following frequency for both muscles.</li>
</ol>
8. Dissection and labeling of the giant fiber system for confocal imaging
NOTE: Fly CNS dissection and dye injection are detailed in Boerner and Godenschwege14.
<ol>
	<li>Gently remove the fly from the dental wax with forceps and transfer it to a silicone elastomer-lined Petri dish under a dissection scope.</li>
	<li>Remove the legs, wings, and proboscis with scissors and use insect pins to secure the fly dorsal side up in the elastomer coating.</li>
	<li>Make a lateral incision along the midline from the middle abdomen, through the entire thorax, and up the neck, connecting the thorax to the head. Open the fly with insect pins. Remove organs and glands from the fly without damaging the nervous system. 
	NOTE: At this point, GF axons can be injected with dyes to label the GFs and electrically coupled neurons13.</li>
	<li>Remove the head capsule from the head to expose the brain.</li>
</ol>
9. Immunohistochemistry of the nervous system
<ol>
	<li>After CNS dissection, fix the fly in 4% PFA in PBS for 45 min at room temperature (RT). Wash for 20 min in PBS at RT (3 times).</li>
	<li>Block the sample for 1 h at RT in 3% BSA in PBS with 0.5% detergent solution.</li>
	<li>Incubate for 2-3 nights at 4 &#176;C in 3% BSA in PBS with 0.3% detergent solution and rabbit anti-GFP (1:500). Wash for 20 min in PBS at RT (6 times).</li>
	<li>Incubate overnight at 4 &#176;C in PBS with anti-rabbit Alexa 488. Wash for 20 min in PBS at RT (6 times).</li>
	<li>Dehydrate samples with an ascending ethanol series (50%, 70%, 90%, 100% ethanol) for 10 min each step. Remove the pins and trim off the abdomen and the flight muscles. Transfer the rest of the thorax containing the nervous system onto a slide and cover it with a mounting medium like methyl salicylate. Place a coverslip on the sample and seal it with nail polish.</li>
	<li>Image the nervous system on a confocal microscope to analyze the anatomy of the GF terminal.</li>
</ol>

Single-Neuron Ablation in Drosophila for Studying Nervous System Adaptation

<ol>
	<li>Chung, S. H., Mazur, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femtosecond+laser+ablation+of+neurons+in+C.+elegans+for+behavioral+studies.">Femtosecond laser ablation of neurons in C. elegans for behavioral studies.</a> Appl Phys A Mater Sci Process. 96 (2), 335-341 (2009).</li><li>Bower, D. V., 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=Airway+branching+has+conserved+needs+for+local+parasympathetic+innervation+but+not+neurotransmission.">Airway branching has conserved needs for local parasympathetic innervation but not neurotransmission.</a> BMC Biol. 12, 92 (2014).</li><li>Angelo, J. R., Tremblay, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laser-mediated+cell+ablation+during+post-implantation+mouse+development.">Laser-mediated cell ablation during post-implantation mouse development.</a> Dev Dyn. 242 (10), 1202-1209 (2013).</li><li>Allen, M. J., Godenschwege, T. A., Tanouye, M. A., Phelan, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Making+an+escape:+Development+and+function+of+the+Drosophila+giant+fibre+system.">Making an escape: Development and function of the Drosophila giant fibre system.</a> Semin Cell Dev Biol. 17 (1), 31-41 (2006).</li><li>Hubel, D. H., Wiesel, T. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Binocular+interaction+in+striate+cortex+of+kittens+reared+with+artificial+squint.">Binocular interaction in striate cortex of kittens reared with artificial squint.</a> J Neurophysiol. 28 (6), 1041-1059 (1965).</li><li>Wiesel, T. N., Hubel, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+the+effects+of+unilateral+and+bilateral+eye+closure+on+cortical+unit+responses+in+kittens.">Comparison of the effects of unilateral and bilateral eye closure on cortical unit responses in kittens.</a> J Neurophysiol. 28 (6), 1029-1040 (1965).</li><li>Brand, A. H., Perrimon, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+gene+expression+as+a+means+of+altering+cell+fates+and+generating+dominant+phenotypes.">Targeted gene expression as a means of altering cell fates and generating dominant phenotypes.</a> Development. 118 (2), 401-415 (1993).</li><li>Allen, M. J., Drummond, J. A., Moffat, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+giant+fiber+neuron+of+Drosophila+melanogaster.">Development of the giant fiber neuron of Drosophila melanogaster.</a> J Comp Neurol. 397 (4), 519-531 (1998).</li><li>Burra, S., Wang, Y., Brock, A. R., Galko, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+Drosophila+larvae+to+study+epidermal+wound+closure+and+inflammation.">Using Drosophila larvae to study epidermal wound closure and inflammation.</a> Methods Mol Biol. 1037, 449-461 (2013).</li><li>Kakanj, P., Eming, S. A., Partridge, L., Leptin, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+in+vivo+imaging+of+Drosophila+larvae.">Long-term in vivo imaging of Drosophila larvae.</a> Nat Protoc. 15 (3), 1158-1187 (2020).</li><li>Bainbridge, S. P., Bownes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Staging+the+metamorphosis+of+Drosophila+melanogaster.">Staging the metamorphosis of Drosophila melanogaster.</a> J Embryol Exp Morphol. 66, 57-80 (1981).</li><li>Allen, M. J., Godenschwege, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+recordings+from+the+Drosophila+giant+fiber+system+(GFs).">Electrophysiological recordings from the Drosophila giant fiber system (GFs).</a> Cold Spring Harb Protoc. 2010 (7), 5453 (2010).</li><li>Augustin, H., Allen, M. J., Partridge, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+recordings+from+the+giant+fiber+pathway+of+d.+Melanogaster.">Electrophysiological recordings from the giant fiber pathway of d. Melanogaster.</a> J Vis Exp. (47), e2412 (2011).</li><li>Boerner, J., Godenschwege, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole+mount+preparation+of+the+adult+Drosophila+ventral+nerve+cord+for+giant+fiber+dye+injection.">Whole mount preparation of the adult Drosophila ventral nerve cord for giant fiber dye injection.</a> J Vis Exp. (52), e3080 (2011).</li><li>Blagburn, J. M., Alexopoulos, H., Davies, J. A., Bacon, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Null+mutation+in+shaking-b+eliminates+electrical,+but+not+chemical,+synapses+in+the+Drosophila+giant+fiber+system:+A+structural+study.">Null mutation in shaking-b eliminates electrical, but not chemical, synapses in the Drosophila giant fiber system: A structural study.</a> J Comp Neurol. 404 (4), 449-458 (1999).</li><li>Kennedy, T., Broadie, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Newly+identified+electrically+coupled+neurons+support+development+of+the+Drosophila+giant+fiber+model+circuit.">Newly identified electrically coupled neurons support development of the Drosophila giant fiber model circuit.</a> eNeuro. 5 (6), 0346 (2018).</li><li>Mcfarland, B. 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The authors have nothing to disclose.

Cell ablation with a 2-photon microscope proved to be a highly successful method to manipulate neuronal circuit development in Drosophila. Since this method is non-invasive, it causes minimal damage to the animal. The data support the usefulness of this cell-specific manipulation of known circuits.
Crucial for the success of the ablation was selecting the most appropriate Gal4 driver. Since the GFS is well studied, many specific Gal4 driver lines have been described7...

Laser ablation is a preferred tool for dissecting neural circuits in a wide variety of organisms. Developed in model genetic systems like worms and flies, it has been applied across the animal kingdom to study the structure, function, and development of the nervous system1,2,3. Here, single-neuron ablation was employed to investigate how neurons interact during circuit assembly in Drosophila. The escape system of the fly is a favorite circuit for analysis because it contains the largest neurons and the largest synapses in the adult fly, and the circuit has been well-characterized in the past decades4. The role neuron-neuron interactions play in the assembly of the Giant Fiber circuit is a focal point of this research.
One type of interaction that has been a focal point in neuroscience since the work of Hubel and Wiesel in the 1960s is &#34;synaptic competition&#34;5,6. In this protocol, laser ablation was used to revisit the role of competition through single-cell ablation in the giant fiber system (GFS) of Drosophila, where the molecular underpinnings of the phenomena might be discovered.
Ablation of neurons in the developing fly has been difficult for a variety of reasons, including visualizing the target neurons, the precision of the ablation method, and the survival of the specimen. To overcome these problems in the GFS, the UAS/Gal4 system7 was used to label neurons of interest, and a two-photon microscope was used to remove the presynaptic giant fiber or the postsynaptic jump motor neuron (TTMn).
In this study, to determine the role that neighboring bilateral neurons play in adjusting synaptic connectivity and synaptic strength in the GFS, one of the bilateral pairs of neurons (either presynaptic GF or postsynaptic motor neuron) was deleted just before pupal development. At this developmental stage, GF axonogenesis has not been completed8. The GF structure and function of the synaptic circuit in the adult were then examined, with particular attention given to the output of the remaining GF.

<table><tbody><tr><td>Alexa Fluor 488 AffiniPure Goat Anti-Rabbit IgG (H+L)</td><td>Jaxkson ImmunoResearch</td><td>111-545-003</td><td></td></tr><tr><td>Anti-green fluorescent protein, rabbit</td><td>Fisher Scientific</td><td>A11122</td><td>1:500 concentration</td></tr><tr><td>Apo LWD 25x/1.10W Objective</td><td>Nikon</td><td>MRD77220</td><td>water immersion long working distance</td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma</td><td>B4287-25G</td><td></td></tr><tr><td>Chameleon Ti:Sapphire Vision II Laser</td><td>Coherent</td><td></td><td></td></tr><tr><td>Cotton Ball</td><td>Genesee Scientific</td><td>51-101</td><td></td></tr><tr><td>Dextra, Tetramethylrhodamine, 10,000 MW, Lysine Fixable (fluoro-Ruby)</td><td>Fisher Scientific</td><td>D1817</td><td></td></tr><tr><td>Drosophila saline</td><td></td><td></td><td>recipe from Gu and O&#039;Dowd, 2006</td></tr><tr><td>Ethyl Ether</td><td>Fisher Scientific</td><td>E134-1</td><td>Danger, Flammable liquid</td></tr><tr><td>Fly food B (Bloomington recipe)</td><td>LabExpress</td><td>7001-NV</td><td></td></tr><tr><td>Methyl salicylate</td><td>Fisher Scientific</td><td>O3695-500</td><td></td></tr><tr><td>Microcentrifuge tube 1.5 mL</td><td>Eppendorf</td><td>22363204</td><td></td></tr><tr><td>Microscope cover-slip 18x18 #1.5</td><td>Fisher Scientific</td><td>12-541A</td><td></td></tr><tr><td>Neurobiotin Tracer</td><td>Vector Laboratories</td><td>SP-1120</td><td></td></tr><tr><td>Nikon A1R multi-photon microscope</td><td>Nikon</td><td></td><td>on an upright FN1 microsope stand</td></tr><tr><td>NIS Elements Advanced Research</td><td>Nikon</td><td></td><td>Acquisition and data analysis software</td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Fisher Scientific</td><td>T353-500</td><td></td></tr><tr><td>PBS (Phosphate Buffered Salin)</td><td>Fisher BioReagents</td><td>BP2944-100</td><td>Tablets</td></tr><tr><td>R91H05-Gal4</td><td>Bloomington Drosophila Stock Center</td><td>40594</td><td></td></tr><tr><td>shakB(lethal)-GAl4</td><td>Bloomington Drosophila Stock Center</td><td>51633</td><td></td></tr><tr><td>Superfrost microscope glass slide</td><td>Fisher Scientific</td><td>12-550-143</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher Scientific</td><td>422355000</td><td>detergent solution</td></tr><tr><td>UAS-10xGFP</td><td>Bloomington Drosophila Stock Center</td><td>32185</td><td></td></tr></tbody></table>

laser cell ablation in intact drosophila larvae reveals synaptic competition

The protocol describes single-neuron ablation with a 2-photon laser system in the central nervous system (CNS) of intact Drosophila melanogaster larvae. Using this non-invasive method, the developing nervous system can be manipulated in a cell-specific manner. Disrupting the development of individual neurons in a network can be used to study how the nervous system can compensate for the loss of synaptic input. Individual neurons were specifically ablated in the giant fiber system of Drosophila, with a focus on two neurons: the presynaptic giant fiber (GF) and the postsynaptic tergotrochanteral motor neuron (TTMn). The GF synapses with the ipsilateral TTMn, which is crucial to the escape response. Ablating one of the GFs in the 3rd instar brain, just after the GF starts axonal growth, permanently removes the cell during the development of the CNS. The remaining GF reacts to the absent neighbor and forms an ectopic synaptic terminal to the contralateral TTMn. This atypical, bilaterally symmetric terminal innervates both TTMns, as demonstrated by dye coupling, and drives both motor neurons, as demonstrated by electrophysiological assays. In summary, the ablation of a single interneuron demonstrates synaptic competition between a bilateral pair of neurons that can compensate for the loss of one neuron and restore normal responses to the escape circuit.

This method can be used to manipulate the development of specific neuronal networks in the nervous system of Drosophila. The primary research question here was the formation of synaptic connections. Removing either the presynaptic GF or the postsynaptic TTMn enabled the investigation of reactive synaptogenesis at this central synapse and the molecular mechanisms crucial for synaptic function and development. As described in the protocol, laser cell ablation of one of the GFs or one of the TTMns was performed, an...

Author Spotlight: Investigating the Mechanisms of Neural Circuit Assembly and Synapse Formation in Drosophila

This protocol demonstrates the laser cell ablation of individual neurons in intact Drosophila larvae. The method enables the study of the effect of reducing competition between neurons in the developing nervous system.

Laser Cell Ablation in Intact Drosophila Larvae Reveals Synaptic Competition

The protocol describes single-neuron ablation with a 2-photon laser system in the central nervous system (CNS) of intact Drosophila melanogaster larvae. ...

laser-cell-ablation-intact-drosophila-larvae-reveals-synaptic

Research

JoVE Journal

Neuroscience

376 Views.  Florida Atlantic University. This protocol demonstrates the laser cell ablation of individual neurons in intact Drosophila larvae. The method enables the study of the effect of reducing competition between neurons in the developing nervous system.

Video: Author Spotlight: Investigating the Mechanisms of Neural Circuit Assembly and Synapse Formation in Drosophila 

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The Drosophila larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila is well known for the ease of powerful genetic manipulations and the larval nervous system has proven particularly useful in studying not only normal function but also perturbations that accompany some neurological disease (Lloyd and Taylor, 2010). Many key synaptic molecules found in Drosophila are also found in mammals and like most CNS excitatory synapses in mammals, the Drosophila NMJ is glutamatergic and demonstrates activity-dependent remodeling (Kohet al. , 2000). Additionally, Drosophila neurons can be individually identified because their innervation patterns are stereotyped and repetitive making it possible to study identified synaptic terminals, such as those between motor neurons and the body-wall muscle fibers that they innervate (Keshishian and Kim, 2004). The existence of evolutionarily conserved synapse components along with the ease of genetic and physical manipulation make the Drosophila model ideal for investigating the mechanisms underlying synaptic function (Budnik, 1996).
The active zones at synaptic terminals are of particular interest because these are the sites of neurotransmitter release. NC82 is a monoclonal antibody that recognizes the Drosophila protein Bruchpilot (Brp), a CAST1/ERC family member that is an important component of the active zone (Waghet al. , 2006). Brp was shown to directly shape the active zone T-bar and is responsible for effectively clustering Ca2+ channels beneath the T-bar density (Fouquetet al. , 2009). Mutants of Brp have reduced Ca2+ channel density, depressed evoked vesicle release, and altered short-term plasticity (Kittelet al. , 2006). Alterations to active zones have been observed in Drosophila disease models. For example, immunofluorescence using the NC82 antibody showed that the active zone density was decreased in models of amyotrophic lateral sclerosis and Pitt-Hopkins syndrome (Ratnaparkhiet al. , 2008; Zweieret al. , 2009). Thus, evaluation of active zones, or other synaptic proteins, in Drosophila larvae models of disease may provide a valuable initial clue to the presence of a synaptic defect.
Preparing whole-mount dissected Drosophila larvae for immunofluorescence analysis of the NMJ requires some skill, but can be accomplished by most scientists with a little practice. Presented is a method that provides for multiple larvae to be dissected and immunostained in the same dissection dish, limiting environmental differences between each genotype and providing sufficient animals for confidence in reproducibility and statistical analysis.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The neuromuscular junction (NMJ) of Drosophila melanogaster is an important model system for studying normal synaptic function as well as perturbations to synaptic function found in certain neurological diseases. We present a protocol for dissection of the Drosophila larval motor system and immunostaining for active zone proteins within the NMJ.

The Drosophila  larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila  is well known for ...

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Bioengineering

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the underlying neuronal components in the circuits1-6. Application of optogenetics7,8 in the larval neural circuit allows us to manipulate neuronal activity in spatially and temporally patterned ways9-13. Typically, specimens are broadly illuminated with a mercury lamp or LED, so specificity of the target neurons is controlled by binary gene expression systems such as the Gal4-UAS system14,15. In this work, to improve the spatial resolution to "sub-genetic resolution", we locally illuminated a subset of neurons in the ventral nerve cord using lasers implemented in a conventional confocal microscope. While monitoring the motion of the body wall of the semi-intact larvae, we interactively activated or inhibited neural activity with channelrhodopsin16,17 or halorhodopsin18-20, respectively. By spatially and temporally restricted illumination of the neural tissue, we can manipulate the activity of specific neurons in the circuit at a specific phase of behavior. This method is useful for studying the relationship between the activities of a local neural assembly in the ventral nerve cord and the spatiotemporal pattern of motor output.

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Here we describe a protocol for optogenetic manipulation of motoneuronal activity while monitoring changes in motor output (muscle contraction) in semi-intact Drosophila larvae using lasers within a conventional confocal microscope. This technique enables researchers to achieve local perturbation of neural activity in a few neuromeres to elucidate the dynamics of motor circuits.

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the ...

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

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

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

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

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

Biology

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Medicine

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because the larval body wall and most internal organs are transparent. However, continuous live imaging of intact Drosophila larvae for longer than 30 min has been challenging because it is difficult to noninvasively immobilizeimmobilizing larvae for a long time. Here we present a larval mounting method called LarvaSPA that allows for continuous imaging of live Drosophila larvae with high temporal and spatial resolution for longer than 10 hours. This method involves partially attaching larvae to the coverslip using a UV-reactive glue and additionally restraining larval movement using a polydimethylsiloxane (PDMS) block. This method is compatible with larvae at developmental stages from second instar to wandering third instar. We demonstrate applications of this method in studying dynamic processes of Drosophila somatosensory neurons, including dendrite growth and injury-induced dendrite degeneration. This method can also be applied to study many other cellular processes that happen near the larval body wall.

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

This protocol describes a method for mounting Drosophila larvae to achieve longer than 10 h of uninterrupted time-lapse imaging in intact live animals. This method can be used to image many biological processes close to the larval body wall.

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because ...

Two-Photon Laser-Induced Neural Injury: A Method to Observe Axon Degeneration and Regeneration in Drosophila Larvae

Source: Li, D., et al. <a href="https://www.jove.com/video/57557/" target="_blank">A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System</a>. J. Vis. Exp. (2018).
This video describes two-photon laser ablation of axons in a Drosophila melanogaster larva and an example protocol to induce injury and image the injury site.

Source: Li, D., et al. A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System. J. Vis. Exp. 
...

Laser Cell Ablation in Intact Drosophila Larvae Reveals Synaptic Competition

Summary

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Chapters in this video

Laser Capture Microdissection of Drosophila Peripheral Neurons

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

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

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Two-Photon Laser-Induced Neural Injury: A Method to Observe Axon Degeneration and Regeneration in Drosophila Larvae