We thank Jessica Goldshteyn for technical support. Work in the Song lab is funded by the NIH grant R00NS088211, and the Intellectual and Developmental Disabilities Research Center (IDDRC) New Program Development Award.

1. Preparation of Culture Plates and Bottles
<ol>
	<li>Preparation of grape juice agar plates
	<ol>
		<li>Add 10 g of agar powder, 200 mL grape juice, and 192 mL ddH2O into a beaker and microwave for about 4-5 min, stirring intermittently until the agar is completely dissolved.</li>
		<li>In a fume hood, cool down the solution to approx. 60 &#176;C. Add 4.2 mL 95% ethanol and 4.0 mL glacial acetic acid. Adjust the total volume of the solution to 400 mL with ddH2O. Mix well.</li>
		<li>For each 35-mm plate, add about 2-3 mL of the solution. Make approximately 120-150 plates for a total of 400 mL grape juice agar solution.</li>
		<li>Wait for 10 min to cool down the plates and solidify the agar solution. Pack the grape juice agar plates in self-sealed ziplock bags and store at 4&#176;C for future use.</li>
	</ol>
	</li>
	<li>Preparation of Drosophila culture bottles
	<ol>
		<li>Use a blade to punch a 1.5 cm side length triangular hole on one wall of the Drosophila culture bottle and fill the hole with a 2-2.5 cm diameter ball of cotton for ventilation.</li>
		<li>Plug the bottle with a grape juice agar plate supplemented with about 0.5 cm3 of yeast paste.</li>
	</ol>
	</li>
</ol>
2. Collection of Drosophila Larvae
<ol>
	<li>Set up crosses of adult flies
	<ol>
		<li>Place 10 virgin females and 5 male adult flies together in a culture bottle plugged with a grape juice agar plate.</li>
		<li>Place the bottle bottoms up at 25&#176;C, so that flies will lay eggs on the grape juice agar plate. Change the plate with yeast paste at least once a day. For a 2-h collection period, which permits the harvesting of larvae of a homogenous developmental stage, start with more than 20 virgin females and 10 males in Step 2.1.1.</li>
		<li>Culture the plate at 25&#176;C in a 60-mm Petri dish with a wet tissue, for example, soaked in 0.5% propionic acid solution. Arrange the tissue so that it does not block oxygen flow. 
		NOTE: The propionic acid solution is used to maintain humidity in the dish and avoid the growth of mold.</li>
	</ol>
	</li>
	<li>Harvest larvae of a specific age
	<ol>
		<li>Use a pair of forceps to transfer larvae of the desired stage, for example, 2nd to 3rd instar larvae at 48-72 h after egg laying (AEL), to a new grape juice agar plate without yeast paste.</li>
		<li>Remove the yeast from the larvae&#39;s skin by letting them crawl around on the new plate, to prevent from potential interfering of yeast with laser axotomy and imaging. Alternatively, thoroughly clean the larvae by washing them in a dish of PBS and drying briefly on a piece of tissue paper.</li>
	</ol>
	</li>
</ol>
3. Two-photon Injury and Confocal Imaging
<ol>
	<li>Microscope setup 
	NOTE: A confocal laser scanning microscope with a two-photon laser was used for this experiment, but other systems with an equivalent setup will also suffice. The two-photon laser (930 nm) was used for delivering injury and an Argon laser (488 nm) was used for confocal imaging of GFP.
	<ol>
		<li>At the beginning of each session, turn on the two-photon laser and/or the confocal laser(s), and the microscope. Open the imaging software.</li>
		<li>For two-photon injury, set up the following parameters for imaging GFP with the two-photon laser at 930 nm (1,950 mW).
		<ol>
			<li>Select line scan mode. Open up the pinhole all the way. Increase laser intensity to ~20% (390 mW).</li>
			<li>Select 512 x 512 as the frame scan. Use maximal scanning speed (typically with the pixel dwell time at 0.77 &#181;s). Ensure that the average number is 1 and bit depth is 8 bits.</li>
			<li>Set gain to ~750, and the offset to 0.</li>
			<li>Save this pre-set experimental protocol as 2P GFP 930 Ablation, allowing for easy reuse in future experiments.</li>
		</ol>
		</li>
		<li>For confocal imaging, set up the following parameters for imaging GFP with the Argon laser at 488 nm:
		<ol>
			<li>Select the Acquisition tab and then Z-stack.</li>
			<li>Under &#34;Laser&#34;, turn on the power for the 488 nm Argon laser.</li>
			<li>Go to Channels, select the 488 mm laser, and increase the laser power to 5-10%. For the pinhole, use the 1-2 airy unit (AU). Adjust the gain to 650.</li>
			<li>In Acquisition Mode, select 1024 x 1024 as the frame scan, use the maximum scan speed, an average number of 2, and bit depth of 8 Bit.</li>
			<li>Save this pre-set experimental protocol as GFP Imaging.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Larvae anesthesia with diethyl ether and mounting
	<ol>
		<li>In a fume hood, place a 60-mm glass dish in a 15-cm plastic Petri dish. Fold and lay a piece of tissue paper at the bottom of the glass dish, then place a grape juice agar plate on the tissue. Add diethyl ether into the glass dish, to the point where the tissue paper is soaked and there is a layer of liquid ether remaining in the dish. Keep the lid on at all times.</li>
		<li>Prepare a glass slide with one drop of halocarbon 27 oil in the center. Add 4 spots of vacuum grease onto the four corners of the slide, to later support the coverslip.</li>
		<li>Use forceps to pick up a cleaned larva and place it on the agar plate in the 60-mm glass dish. Cover the glass dish with its lid and wait until the larva stops moving. For PNS injury/imaging, take out the larva as soon as its tail stops twitching. For the CNS, wait until the entire larva becomes motionless, especially the head segments. 
		NOTE: The timing of ether exposure is critical. See Discussion.</li>
		<li>Carefully pick up the anesthetized larva and place it head-upright into the drop of halocarbon oil on the slide. Add a coverslip on top of the slide. Use gentle pressure to push down on the coverslip, until it touches the larva (Figure 1A).</li>
		<li>Adjust the larva&#39;s position by gently pushing the coverslip towards the left or right to roll the larva, so that the neuron/axon/dendrite of interest is on the top and closest to the microscope lens.</li>
		<li>For PNS injury, mount the larva dorsal side up, so that both the tracheas are visible. Then roll the larva ~30 degrees to the left for injuring class III da neuron axons (Figure 1B and 1C), 90 degrees for injuring class IV da neuron axons (Figure 1B and 1E), or ~30 degrees for injuring class IV da neuron dendrites (Figure 2A).</li>
		<li>For CNS injury, position the larva to be perfectly ventral side up (Figure 3A), so that the region of interest is closest to the microscope lens in the z-plane.</li>
	</ol>
	</li>
	<li>Injury by two-photon laser
	<ol>
		<li>Place the slide with the larva under the microscope and secure it in place with the slide holder on the stage. Use the 10X (0.3 NA) objective to find the larva.</li>
		<li>Add 1 drop of objective oil onto the coverslip, switch to the 40X (1.3 NA) objective and adjust the focus.</li>
		<li>Switch to the scanning mode and reuse the experimental protocol 2P GFP 930 Ablation. Make sure the pinhole is opened all the way. 
		NOTE: The configuration needs to be optimized based on individual systems.</li>
		<li>Start the Live mode to locate the region of interest (ROI), and fine-tune the settings to achieve good image quality with the appropriate zoom. 
		NOTE: The purpose of this step is to find the neuron/axon/dendrite to injure, rather than taking the best quality image. Therefore, use the minimal settings sufficient to visualize the target area, in order to avoid overexposure or photobleaching.</li>
		<li>Stop Live scan, so that the Crop button will become available. Let the still image serve as the roadmap. Select the Crop function and adjust the scan window to focus on the target to be injured.</li>
		<li>Reduce the ROI to be the size of the prospective site of injury. For example, just cover the width of an axon or a dendrite, to ensure the precision of the injury and reduce damage to neighboring tissues. If desired, zoom in on the ROI before cropping, allowing for more precise injury.</li>
		<li>Open a new imaging window. Reduce the scan speed and increase laser intensity. Determine the increase in laser intensity based on the tissue fluorescence signal scanned in Live mode.</li>
		<li>Typically, set the two-photon laser intensity starting from 25% for PNS injury and 50-100% for VNC injury. For PNS axon injury, ensure that the laser intensity is ~480 mW and pixel dwell time is 8.19 &#956;s. For the VNC axon injury, ensure that the laser intensity and pixel dwell time are usually 965-1930 mW and 8.19-32.77 &#956;s, respectively.</li>
		<li>Start Continuous scan. Leave the cursor hovering over the Continuous button. Keep a close eye on the image and stop the scan as soon as a drastic increase in fluorescence is observed. 
		NOTE: The appearance of the fluorescence spike is due to auto-fluorescence at the injury site.</li>
		<li>Switch back to the Live mode by reusing the settings. Find the region of interest that was just targeted by adjusting the focus. 
		NOTE: A good indication of successful injury is the appearance of a small crater, ring-like structure, or localized debris right at the injury site.</li>
		<li>Move to the next neuron and repeat from Step 3.3.5, to injure multiple neurons in a single animal. Or repeat Step 3.3.5 while gradually increasing the power and/or reducing scan speed if the initial injury was insufficient. 
		NOTE: In the case that the laser power is too high, a large damaged area will be visible in the post-injury live scan image. Too much injury may cause the death of the larva.</li>
		<li>Recover the larva by carefully removing the coverslip and transferring the injured larva onto a new plate with yeast paste. Ditch several caves on the agar plate with forceps; alternatively, make an island of agar in the plate instead of using the whole plate, to reduce the possibility of the larva crawling out of the plate.</li>
		<li>Put the plate in a 60-mm Petri dish with wet tissue (soaked with 0.5% propionic acid solution) and culture at room temperature or 25&#176;C. 
		NOTE: The larva will remain in the larval stage for approximately an extra day at room temperature (22&#176;C) compared to at 25&#176;C.</li>
	</ol>
	</li>
	<li>Post-injury confocal imaging
	<ol>
		<li>Image the injured larva at desired time points by preparing the larva using the same procedure of anesthesia and mounting as in Step 3.2, then imaging with the confocal laser. 
		NOTE: Image the larva at 24 h after injury (AI) to confirm axonal injury and at 48 h AI (class IV da neurons) or 72 h AI (class III da neurons) to assess regeneration.</li>
		<li>Locate the larva using the 10X objective, then switch to a 25X (0.8 NA) objective. Reuse the experimental protocol GFP Imaging.</li>
		<li>Click the Live button and find the same neuron injured previously.</li>
		<li>Set the first and last Z positions in the live scan window. Press stop and click Start experiment to acquire a Z-stack image. 
		NOTE: Make sure that a normalization point (the axon converging point) is included when capturing images so that quantification of regeneration is possible (Figure 1D, 1F) &#8211; this is discussed further in the data analysis section.</li>
		<li>Switch to Image Processing, select the image just taken, and generate a maximum intensity projection. Save both the z-stack and maximum intensity projection images.</li>
	</ol>
	</li>
</ol>
4. Data Analysis
<ol>
	<li>Process and quantify images using either the imaging software or ImageJ.</li>
	<li>Quantification of axon regeneration in the PNS
	<ol>
		<li>Calculate Regeneration percentage, which refers to the percent of regenerating axons among all the axons that were lesioned. Score an axon as regenerating as long as it regrows beyond the injury site.</li>
		<li>Measure Regeneration length, which is the increase in axon length. If quantifying with ImageJ, use the Segmented Line tool to trace the regenerated axon, and use Measure in the Analyze drop-down menu to obtain the length of the line representing the regenerated axon.</li>
		<li>Calculate Regeneration Index, which is the increase in normalized axon length. 
		NOTE: Axon length is normalized by the distance between the cell body and the axon converging point (DCAC) &#8211; axon length/DCAC (Figure 1D and 1F). This value helps account for any axonal growth that is due to larval scaling. A positive value represents regeneration, a value of 0 means no regeneration, and a negative value means retraction.</li>
	</ol>
	</li>
	<li>Quantification of dendrite regeneration
	<ol>
		<li>Calculate Regeneration percentage, which is the percent of da neurons among all those severed that show obvious dendrite regrowth. 
		NOTE: Dendrite regrowth is scored as positive if new dendrites regrow out from the retracted dendritic stem and beyond the injury site. The injury site is determined by landmarks, and in some cases, is readily visible due to the residual injury-induced autofluorescence.</li>
		<li>Calculate Increase of branch points, which counts the addition of new dendritic branch points after injury.</li>
		<li>Calculate Increase of total dendrite length, which is the cumulative length of all the new dendrites added after injury.</li>
	</ol>
	</li>
	<li>Quantification of axon regeneration in the CNS 
	NOTE: If a lesion site shows degeneration at the first time point imaged (Figure 3B), it will be included in the analysis of regeneration length and rate.
	<ol>
		<li>Measure Regeneration length, which is the length of the regrowing axon. 
		NOTE: The regrowing axon of a single lesion site is identified as it originates off the original axon route before the injury.</li>
		<li>Calculate Normalized regeneration length, which normalizes the regeneration length to the length of the commissure segment - the longitudinal distance between commissures (&#34;Y&#34; in Figure 3A and 3B). 
		NOTE: This value helps correct for the effect of larval size differences.</li>
		<li>Calculate Regeneration rate, which is the percent of regenerating segments among all the segments that were lesioned, to reflect the regeneration capacity of a particular genotype.</li>
	</ol>
	</li>
</ol>

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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=Neurosurgery:+functional+regeneration+after+laser+axotomy.">Neurosurgery: functional regeneration after laser axotomy.</a> Nature. 432, 822 (2004).</li><li>Wu, Z., 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=Caenorhabditis+elegans+neuronal+regeneration+is+influenced+by+life+stage,+ephrin+signaling,+and+synaptic+branching.">Caenorhabditis elegans neuronal regeneration is influenced by life stage, ephrin signaling, and synaptic branching.</a> Proc Natl Acad Sci U S A. 104, 15132-15137 (2007).</li><li>Stone, M. C., Nguyen, M. M., Tao, J., Allender, D. L., Rolls, M. 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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+axon+regeneration+and+muscle+reinnervation+in+young+adult+and+aged+animals.">Motor axon regeneration and muscle reinnervation in young adult and aged animals.</a> J Neurosci. 33, 19480-19491 (2013).</li><li>Duan, 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=Subtype-specific+regeneration+of+retinal+ganglion+cells+following+axotomy:+effects+of+osteopontin+and+mTOR+signaling.">Subtype-specific regeneration of retinal ganglion cells following axotomy: effects of osteopontin and mTOR signaling.</a> Neuron. 85, 1244-1256 (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, 451-461 (1999).</li><li>Grueber, W. B., 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=Projections+of+Drosophila+multidendritic+neurons+in+the+central+nervous+system:+links+with+peripheral+dendrite+morphology.">Projections of Drosophila multidendritic neurons in the central nervous system: links with peripheral dendrite morphology.</a> Development. 134, 55-64 (2007).</li><li>Buss, 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=NG2+and+phosphacan+are+present+in+the+astroglial+scar+after+human+traumatic+spinal+cord+injury.">NG2 and phosphacan are present in the astroglial scar after human traumatic spinal cord injury.</a> BMC Neurol. 9, 32 (2009).</li><li>McKeon, R. J., Jurynec, M. J., Buck, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chondroitin+sulfate+proteoglycans+neurocan+and+phosphacan+are+expressed+by+reactive+astrocytes+in+the+chronic+CNS+glial+scar.">The chondroitin sulfate proteoglycans neurocan and phosphacan are expressed by reactive astrocytes in the chronic CNS glial scar.</a> J Neurosci. 19, 10778-10788 (1999).</li><li>Raabe, I., Vogel, S. K., Peychl, J., Tolic-Norrelykke, I. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+nanosurgery+and+cell+enucleation+using+a+picosecond+laser.">Intracellular nanosurgery and cell enucleation using a picosecond laser.</a> J Microsc. 234, 1-8 (2009).</li><li>Hutson, M. S., Ma, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma+and+cavitation+dynamics+during+pulsed+laser+microsurgery+in+vivo.">Plasma and cavitation dynamics during pulsed laser microsurgery in vivo.</a> Phys Rev Lett. 99, 158104 (2007).</li><li>Venugopalan, V., Guerra, A., Nahen, K., Vogel, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+laser-induced+plasma+formation+in+pulsed+cellular+microsurgery+and+micromanipulation.">Role of laser-induced plasma formation in pulsed cellular microsurgery and micromanipulation.</a> Phys Rev Lett. 88, 078103 (2002).</li><li>Bourgeois, F., Ben-Yakar, A. <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+nanoaxotomy+properties+and+their+effect+on+axonal+recovery+in+C.+elegans.">Femtosecond laser nanoaxotomy properties and their effect on axonal recovery in C. elegans.</a> Opt Express. 16, 5963 (2008).</li><li>O'Brien, G. S., 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=Two-photon+axotomy+and+time-lapse+confocal+imaging+in+live+zebrafish+embryos.">Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos.</a> J Vis Exp. , (2009).</li><li>Tsai, P. S., 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=Plasma-mediated+ablation:+an+optical+tool+for+submicrometer+surgery+on+neuronal+and+vascular+systems.">Plasma-mediated ablation: an optical tool for submicrometer surgery on neuronal and vascular systems.</a> Curr Opin Biotechnol. 20, 90-99 (2009).</li><li>Chung, S. H., Clark, D. A., Gabel, C. V., Mazur, E., Samuel, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+the+AFD+neuron+in+C.+elegans+thermotaxis+analyzed+using+femtosecond+laser+ablation.">The role of the AFD neuron in C. elegans thermotaxis analyzed using femtosecond laser ablation.</a> BMC Neurosci. 7, 30 (2006).</li><li>Williams, W., Nix, P., Bastiani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Constructing+a+low-budget+laser+axotomy+system+to+study+axon+regeneration+in+C.+elegans.">Constructing a low-budget laser axotomy system to study axon regeneration in C. elegans.</a> J Vis Exp. , (2011).</li></ol>

The authors have nothing to disclose.

When setting up fly crosses, the number of females and males used can vary depending on the genotypes and the number of larvae needed for specific experiments. For WT flies, typical cross uses 10 females and 5 males. The collection window may be narrowed, depending on the accuracy of the larvae age required. For example, a 2-h collection period will yield larvae of a more homogenous population. In this case, using 20 or more virgin females to set up the crosses will help yield sufficient eggs. The yield from the first da...

The inability of axons to regenerate after an injury to the central nervous system (CNS), may lead to permanent disabilities in patients, and also plays a role in the irreversible neurological deficits in neurodegenerative diseases1,2,3,4,5. The CNS environment, as well as the intrinsic growth ability of neurons, determines whether axons are able to regenerate after trauma. Extracellular factors from oligodendrocyte, astroglial, and fibroblastic sources have been shown to impede neuronal growth4,6,7,8, but the elimination of these molecules only allows for limited sprouting5. Intrinsic regeneration signals can influence regenerative success5,9 and represent potential therapeutic targets, but these processes are still not well-defined at the molecular level. Increases in trophic factor signaling or elimination of endogenous brakes, such as the Pten phosphatase10, can result in axonal regeneration in certain circumstances. Combinations of different methods found to be individually effective also only provide limited overall recovery to date11,12,13,14. Therefore, there is a desperate need to identify additional pathways for targeted therapy. In addition to the initiation of axon regrowth, whether and how axons re-wire to the correct target, reform synapse specificity, and achieve functional recovery are important unanswered questions.
In summary, current understanding of the machinery dictating axon regeneration is still very fragmentary. Part of the problem is the technical difficulty of studying axon regeneration in mammals in real-time, an approach that is costly, time-consuming, and challenging for conducting large-scale genetic screens. Drosophila melanogaster, on the other hand, has proven to be an exceptionally powerful system for the study of complex biological questions. The fruit fly has been instrumental in defining genes and signaling pathways that are strikingly conserved in humans and has been a successful model for the study of human conditions, such as neurodegenerative diseases, through the vast molecular genetics tools available to manipulate gene function15. In particular, fruit flies are considered to be an ideal tool for the discovery of genes involved in neural injury and regrowth15,16. Several fly neural injury models have been developed, including adult-head or larval ventral nerve cord (VNC) stabbing with needles, larval VNC or nerve crush with forceps, larval neuron laser axotomy, olfactory receptor neuron removal, brain explants injury, and peripheral nerve lesion by wing severance15,17,18,19,20,21,22,23. Excitingly, recent work utilizing Drosophila injury models have&#160;advanced our understanding of the cellular and genetic pathways used by the nervous system to respond to neural injuries, some of which have been shown to be conserved in mammals24,25. Again, this emphasizes the utility of this model organism for identifying novel mechanisms of neural repair.
Described here is a two-photon laser-based Drosophila larval sensory neuron injury model. A two-photon laser was first used to cut axons in zebrafish in vivo in 200326. In the same year, the first laser dendriotomy was performed in Drosophila using a pulsed nitrogen laser27. Shortly afterward, several C. elegans labs used femtosecond lasers to establish models of axon regeneration28. In 2007, Wu and colleagues compared and reported the differences between laser injuries in C. elegans induced by various types of lasers29. In 2010, axon regeneration after laser axotomy was first shown to occur in Drosophila30. Building on this extensive laser injury literature, we have developed a fly neural injury model using the two-photon laser, which allows precise induction of injury to targeted sites with minimal perturbation of neighboring tissues, providing a relatively clean system to study both the intrinsic and extrinsic properties of neuroregeneration with single-cell resolution. Specifically, we have established a set of injury methods for dendritic arborization (da) sensory neurons in both the peripheral nervous system (PNS) and CNS. Da neurons can be grouped into four distinct classes distinguished primarily by their dendrite branching complexities: class I to IV31. Our published work shows that da neuron regeneration resembles mammalian injury models at the phenotypic and molecular level: da neurons display class specific regeneration properties, with class IV but not class I or III da neurons exhibiting regeneration in the PNS; class IV da neuron axons regenerate robustly in the periphery, but their regenerative potential is dramatically reduced in the CNS, thus resembling dorsal root ganglion (DRG) neurons in mammals; enhancing mTOR activity via Pten deletion or Akt overexpression enhances axon regeneration in the fly CNS19. Utilizing this injury model, we have been performing genetic screens and have identified the RNA processing enzyme Rtca as an evolutionarily conserved inhibitory factor for axon regeneration, linking axon injury to cellular stress and RNA modification20.
In the presented paradigm, the injury is induced via laser axotomy/dendriotomy of larval class IV or III da neurons, labeled by ppk-CD4-tdGFP or 19-12-Gal4, UAS-CD4-tdGFP, repo-Gal80, respectively. The injury is performed on 2nd to 3rd instar larvae at around 48 - 72 h after egg laying (h AEL). For PNS axotomy the lesion is targeted to the section of axon ~20 - 50 &#181;m away from the cell body, for CNS axotomy to an area of ~20 &#181;m in diameter at the commissure junction in the VNC, and for dendriotomy to the primary dendritic branch points. The same neuron is imaged at 8 - 24 h after injury (AI) to confirm complete transection, and at 48 - 72 h AI to assess regeneration. Through time-lapse confocal imaging, the degeneration and regeneration of individual axons/dendrites that have been injured in vivo can be monitored over time.

<table><tbody><tr><td>Diethyl ether, ACS reagent, anhydrous</td><td>Acros Organics</td><td>AC615080010</td><td></td></tr><tr><td>Halocarbon 27 Oil</td><td>Genesee Scientific</td><td>59-133</td><td></td></tr><tr><td>Phosphate buffered saline (PBS), 20x Concentrate, pH 7.5, supplier # E703-1L</td><td>VWR</td><td>97062-948&amp;nbsp;</td><td></td></tr><tr><td>Agar powder, Alfa Aesar, 500GM</td><td>VWR</td><td>AAA10752-36</td><td></td></tr><tr><td>Grape juice</td><td>Welch&amp;rsquo;s</td><td></td><td></td></tr><tr><td>Ethanol 95% (Reagent Alcohol 95%)</td><td>VWR</td><td>64-17-5</td><td></td></tr><tr><td>Acetic acid</td><td>Sigma-Aldrich</td><td>A6283</td><td></td></tr><tr><td>Propionic Acid</td><td>J.T.Baker</td><td>U33007</td><td></td></tr><tr><td>Cover Glasses: Rectangles</td><td>Fisher Scientific</td><td>12-544-D</td><td>50 mm X 22 mm</td></tr><tr><td>Zeiss LSM 880 laser scanning microscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Zen software</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Chameleon Ultra II</td><td>Coherent</td><td></td><td></td></tr></tbody></table>

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.

Da neurons show differential regeneration potential between the peripheral and central nervous system, as well as class specificity. This provides unique opportunities to screen for novel factors that are required for axon regeneration (using class IV PNS injury), as well as those that are inhibitory for regeneration (with class IV CNS injury and class III PNS injury).
Axon regeneration in the PNS
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A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System | Medicine | Video

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.

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 ...

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9.6K Views.  The Children's Hospital of Philadelphia. 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.

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

Single Drosophila Ommatidium Dissection and Imaging

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

Single Drosophila Ommatidium Dissection and Imaging

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

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

Neuroscience

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

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

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

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

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

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

Developmental Biology

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

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

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

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

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

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

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

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

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

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

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

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

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

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

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

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

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

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. 
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JoVE Encyclopedia of Experiments

Drosophila melanogaster (fruit fly)

Larva

Inducing Targeted Neuronal Injury Using a High-Power Laser in a Drosophila Larva

Source: Li, D., Li, F., et al. <a href="https://app.jove.com/v/57557">A Drosophila in vivo injury model for studying neuroregeneration in the peripheral and central nervous system.</a> J. Vis. Exp. (2018)
This video demonstrates a method for inducing targeted neuronal injury in Drosophila larvae using a high-power laser. After positioning the larva under a microscope, a low-power laser scans for fluorescent neurons, and then the laser power is increased to induce localized damage. The injury is confirmed by increased fluorescence and a crater at the injury site.

Source: Li, D., Li, F., et al. A Drosophila in vivo injury model for studying neuroregeneration in the peripheral and central nervous system. J. Vis. Exp. ...

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

Summary

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

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Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

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

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

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

Inducing Targeted Neuronal Injury Using a High-Power Laser in a Drosophila Larva