Study of Axonal Injury and Degeneration in Drosophila
- 1Department of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann Arbor, Michigan 48109, USA
- 2Department of Neurosciences, Case Western Reserve University, Cleveland, Ohio 44106, USA
- ↵4Correspondence: cxc1215{at}case.edu
Abstract
A fundamental feature of nervous systems is a highly specified synaptic connectivity between cells and the ability to adaptively change this connectivity through plasticity mechanisms. Plasticity mechanisms are highly relevant for responding to nervous system damage, and studies using nervous system injury paradigms in Drosophila (as well as other model organisms) have revealed conserved molecular pathways that are triggered by axon damage. Simple assays that introduce injuries to axons in either adult flies or larvae have proven to be particularly powerful for uncovering mechanisms of axonal degeneration and clearance. They have also been used to reveal requirements for regrowth of axons and dendrites, as well as signaling pathways that regulate cellular responses to nerve injury. Here we review commonly used and simple to carry out techniques that enable experimenters to study responses to axonal damage in either adult flies (following antennal transection) or larvae (following nerve crush to segmental nerves). Because axons and dendrites in the larval peripheral nervous system can be readily visualized through the translucent cuticle, another versatile method to probe injury responses is to focus high-energy laser light to a small and specific location in the animal. We therefore discuss a method for immobilizing intact larvae for imaging through the cuticle to carry out injury by pulse dye laser, which can be used to generate many different kinds of injuries and directed ablations within intact larvae. These techniques, combined with powerful genetic tools in Drosophila, make the fruit fly an excellent model system for studying the effects of injury and the mechanisms of axon degeneration, synapse plasticity, and immune response.
INTRODUCTION
Injury is a natural hazard of life. For the nervous system, injury is a strong trigger of innate plasticity mechanisms, many of which are highly conserved across animal life. The Drosophila nervous system has been probed for its responses to many different kinds of injuries (Fang and Bonini 2012; Rooney and Freeman 2014; Hao and Collins 2017; Brace and DiAntonio 2020), including models of head-impact brain injury in adult flies (Katzenberger et al. 2013; Saikumar et al. 2020), removal of body structures that contain neurons (antennae, legs, or part of the wings) (MacDonald et al. 2006; Fang et al. 2013; Neukomm et al. 2014; Purice et al. 2017), peripheral nerve crush (Xiong et al. 2010), and focused laser microsurgery to axons and/or dendrites in defined locations (Stone et al. 2010; Ghannad-Rezaie et al. 2012; Song et al. 2012; Soares et al. 2014; DeVault et al. 2018). Many of these studies have revealed cellular pathways triggered by injury shared in vertebrates and mammals.
Most compellingly, key components of a molecular pathway that drives the degeneration of injured axons and dendrites removed from their cell bodies were discovered through forward genetic screens in flies (Osterloh et al. 2012; Neukomm et al. 2017). Termed Wallerian degeneration, after its first description by Augustus Waller in 1850, this pathway occurs when axons that become disconnected from their cell body following nerve injury degenerate through a “self-destruction” program akin to apoptosis but with different molecular components (Coleman and Höke 2020). It was first demonstrated more than 15 years ago that the degeneration of injured axons from both flies and mice can be delayed through similar genetic manipulations that increase the levels of nicotinamide mononucleotide adenylyltransferase (NMNAT) (Mack et al. 2001; Hoopfer et al. 2006; Llobet Rosell and Neukomm 2019). Because axons are supplied with this essential enzyme via transport from the cell body, injury to axons causes a drop in NMNAT levels, which leads to local degeneration of the distal stump (Gilley and Coleman 2010; Coleman and Höke 2020).
A key effector of axonal degeneration was discovered through a forward genetic screen in flies. Osterloh et al. (2012) used the mosaic analysis with a repressible cell marker (MARCM) approach to introduce homozygous EMS-generated mutations into GFP-labeled olfactory receptor neurons (ORNs) and screened for mutations that could alter the degeneration or clearance of distal axons following the removal of their cell bodies using a simple antenna removal surgery described in Protocol: Wallerian Degeneration and Clearance of Olfactory Receptor Neuron Axons Following Drosophila Antennal Transection (Waller et al. 2024a). This screen revealed that the degeneration of injured axons requires the function of the sterile-α and Toll/interleukin-1 receptor (TIR) domain-containing protein SARM1. Follow-up structure–function and biochemical studies in mammalian neurons established that SARM1 is a highly regulated enzyme that degrades NAD+ (Coleman and Höke 2020). Since NAD+ is a key coenzyme for metabolism and cellular respiration, its rapid loss following SARM1 activation is thought to drive metabolic catastrophe, where ATP drops below the level required for cellular function (Loring and Thompson 2020). The mechanisms of SARM1 and NMNAT in driving degeneration remain an active area of research (Sambashivan and Freeman 2021; Waller and Collins 2022; Smith et al. 2023). An indication of additional checkpoints downstream from SARM1 activation and NAD+ loss comes from the discovery that SARM1-induced axon loss can be generated by mutations in Axundead (Axed) (Neukomm et al. 2017). This raises fascinating questions of how Wallerian degeneration remains so carefully regulated even after critical loss of such a vital metabolite as ATP.
Coupled with this cell-autonomous and local mechanism for destruction is the conversion of damaged axons and dendrites into debris that can be effectively cleared by surrounding glial cells or epithelial cells, depending on the location (Doherty et al. 2009; Han et al. 2014). Drosophila have proven to be a powerful model for unraveling the cellular pathways that are engaged by cells that recognize and phagocytose neuronal debris (MacDonald et al. 2006; Sapar and Han 2019) in addition to Wallerian degeneration.
Because of the conservation and broad interest in degeneration and clearance mechanisms, we describe in our associated protocols two technically simple and widely used methods that will enable experimenters to probe the effect of their own manipulations on the process of degeneration and clearance of damaged axons in the adult fly brain (see Protocol: Wallerian Degeneration and Clearance of Olfactory Receptor Neuron Axons Following Drosophila Antennal Transection [Waller et al. 2024a]) and larval peripheral nerves (see Protocol: Peripheral Nerve Crush in Drosophila Larvae [Waller et al. 2024b]).
The ability of injured axons or dendrites to grow again (“regenerate”) has also been studied using various injury paradigms in Drosophila (Soares et al. 2014; Hao and Collins 2017; Li et al. 2018; Brace and DiAntonio 2020; Wang et al. 2020). Study of regeneration requires an injury method that preserves the cell body and enables visualization of the new growth from the cell body (or proximal axon stump) following the lesion. Such an assay has been developed for sensory neuron axons in the fly wing (Fang and Bonini 2012; Fang et al. 2013; Soares et al. 2014) and in the larval peripheral nervous system (PNS), where GFP-expressing cells can be visualized through the semitranslucent cuticle. These assays include laser-induced transection of axons and/or dendrites in the dendritic arborization (da) sensory neurons, whose dendrites and cell bodies can be readily visualized in intact animals and subjected to focal microsurgery using a two-photon confocal microscope (Song et al. 2012; Li et al. 2018) or MicroPoint nitrogen pump pulse dye UV (Stone et al. 2010) laser. Time-lapse imaging in intact animals has enabled studies of the degeneration and clearance of injured dendrites (Tao and Rolls 2011; Chen et al. 2012, 2016; Han et al. 2014), as well as regeneration of the dendritic arbor (Stone et al. 2014; Thompson-Peer et al. 2016), which exhibits molecular requirements distinct from those of axon regeneration (Song et al. 2012; Stone et al. 2014; Rao et al. 2016; Nye et al. 2020). Sensory neuron axons injured close to their cell bodies in the periphery show robust ability to regrow along their initial path (Song et al. 2012; Rao et al. 2016; Rao and Rolls 2017). In contrast, growth following injury to sensory neuron axon terminals within the nerve cord is not robust but can be induced by the addition of genetic (or optogenetic) manipulations that have been previously observed to stimulate axonal regeneration in the mammalian central nervous system (CNS) (Song et al. 2012; Wang et al. 2020). Therefore, both mechanisms that enable and restrict the ability of injured axons to initiate new growth can be studied in Drosophila. Excitingly, recent studies have reported manipulations that enable functional recovery of touch sensation following injury to sensory neuron terminals in the CNS (Li et al. 2020; Wang et al. 2020).
For the versatility of laser-induced microsurgery in Drosophila larvae, we describe a protocol for introducing focal injuries to axons and dendrites in the larval PNS using a MicroPoint laser (see Protocol: Laser Microsurgery on Drosophila Larvae Using the MicroPoint Ablation System [Smithson et al. 2024b]).The MicroPoint laser is a versatile pulsed dye laser that can be mounted onto any standard microscope and therefore is an affordable alternative to two-photon lasers for inducing high-powered focal ablations. Laser surgery in intact animals is enabled by noninvasive methods to immobilize live larvae for imaging. For this we describe a method for immobilizing larvae for imaging using a readily obtainable polydimethylsiloxane (PDMS) larva chip (see Protocol: Immobilizing Second-Instar Drosophila Larvae for Imaging and Surgery Using the Larva Chip [Smithson et al. 2024a]).
ACKNOWLEDGMENTS
This work was supported by the National Institutes of Health (RO1 NS069844 to C.A.C.) and the Canadian Institutes of Health Research (postdoctoral fellowship to L.J.S.).
Footnotes
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↵3 Co-first authors.
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From the Drosophila Neurobiology collection, edited by Bing Zhang, Ellie Heckscher, Alex C. Keene, and Scott Waddell.
