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date: 03 August 2020

Mechanisms of Axonal Degeneration and Regeneration: Lessons Learned From Invertebrates

Abstract and Keywords

In the face of acute or chronic axonal damage, neurons and their axons undergo a number of molecular, cellular, and morphological changes. These changes facilitate two types of responses, axonal degeneration and regeneration, both of which are remarkably conserved in both vertebrates and invertebrates. Invertebrate model organisms, including Drosophila and C. elegans, have offered a powerful platform with accessible genetic tools for manipulation and amenable nervous system for visualization. Thus far, several critical components and pathways in axonal degeneration and regeneration have been identified in invertebrate studies, including Sarm and Wallenda/DLK. This article highlights important findings in Drosophila, C. elegans, and other invertebrate injury models that have shed light upon the mechanism in axonal injury response.

Keywords: axon damage, degeneration, adaptation, regeneration, Sarm, Nmnat, DLK/Wallenda, cytoskeleton, intracellular transport

Neurons and the connections that they make with each other typically need to persist for an animal’s entire life, even in the face of injury. How do nervous systems, including our own, cope with and respond to damage? It would be ideal if we simply could regenerate the damaged part, like a planarian which can regenerate an entire head de novo (Owlarn & Bartscherer, 2016). However, most nervous systems across the animal kingdom lack such capacity. Instead, many nervous systems do their best to repair damaged axons and synapses. And in many cases in adult nervous systems, damaged axons and synapses are simply lost.

Axons are thought to be particularly vulnerable components of neuronal circuitry. They are often exceptionally long: Human motoneuron axons can reach lengths over 10,000 times the diameter of their cell body, and even in small invertebrates such as Drosophila an axon can be 200 times the cell body diameter. Such length can be a vulnerability: A problem occurring anywhere along the axon’s length could result in lost ability to communicate with downstream cells. Furthermore, because axons connect to distant sites, it can be difficult to re-establish lost connections, especially in the adult nervous system. During development, axonal growth to specific targets is directed via a series of growth promoting and guidance cues (Tessier-Lavigne & Goodman, 1996). Many of these cues are only transiently present during nervous system development and are largely absent in the adult system.

In this chapter we review some important factors that mediate neuronal responses to axonal damage. We will emphasize here what has been learned from research using invertebrate model organisms (especially Drosophila and C. elegans), which has directed exciting discoveries of mechanisms that are conserved across the animal kingdom. (p. 576)

Overview of Acute and Chronic Models of Axonal Damage

The response(s) that neurons make to axonal damage or stress can vary according to the types of damage and neuron type (Fig. 25.1, and Table 25.1). However, a simple determinant for classification is the duration of the harmful stimulus: acute (short term, cartooned in Fig. 25.1A–C) verses chronic (long term, cartooned in Fig. 25.1D–E).

Mechanisms of Axonal Degeneration and RegenerationLessons Learned From Invertebrates

Figure 25.1 Axon regeneration and degeneration in response to acute and chronic injuries. (A) Acute injuries physically break axons into two parts: a proximal stump, which remains connected to the cell body, and a distal stump, which has lost this connection. In many cases the distal stump has presynaptic terminals (cartooned as button-shaped boutons) that are made nonfunctional by the injury. (B) In response to acute injuries, the proximal stump either (i) succeeds or (ii) fails to form a new growth cone, and the distal axon either (iii) degenerates or (iv) stays intact. Responses vary in different injury models (Table 25.1); however, in most cases, outcomes (i) and (iii) occur. (C) Ultimate outcomes of the injury responses include (i) new growth (in green) from the proximal stump to replace the lost distal stump. Alternatively, some invertebrate neurons have been observed to undergo (ii) fusion of the two stumps, which requires less new growth from the proximal stump and the ability of the distal stump to remain intact until it can be reconnected. (iii) Failure to regenerate axons, followed by degeneration, is another outcome for some neuronal injuries. (D) Chronic injuries include long-term forms of stress that neurons may experience in their axons or synaptic terminals as the result of a genetic mutation or an environmental condition. Such stresses can include perturbations that impair mitochondrial function, organization of the cytoskeleton, long-distance transport of proteins and organelles in axons, and impairments to synaptic transmission. (E) Responses to chronic injuries include degeneration of (i) synaptic terminals or (ii) entire axons and even cell death. However, (iii) neurons may also initiate stress response pathways, which may allow for an enhanced resiliency to degeneration. This likely involves transcriptional and translational events in the cell body and transport of newly synthesized proteins into axons (hence the green nucleus and red cytosol).

Acute axon injuries can be induced experimentally by directly transecting or crushing nerves, or by microsurgical cutting of individual axons using a high-energy laser. After such an injury, repair may be possible if the part of the axon that remains attached to the cell body (the proximal “stump”) can grow again. The ability of an injured axon to reinitiate new axonal growth and eventually reconnect to its target (Fig. 25.1Bi and 25.1Ci) is commonly termed axon “regeneration.” Axon regeneration has been documented and studied in many invertebrate models, including cockroach, crickets, crayfish, squid, Aplysia, Great pond snail, earthworm, leech, and more recently C. elegans and fruit flies (Table 25.1). Is the ability to regenerate axons universal for invertebrate neurons? Probably not, since failures have also been documented (Wu et al., 2007; Ayaz et al., 2008; Song et al., 2012). It is interesting that some of these failures occur after (p. 577) (p. 578) (p. 579) (p. 580) injuries in the central nervous system (CNS) (Ayaz et al., 2008; Song et al., 2012), where stalled regeneration (Fig. 25.1Bii), followed by degeneration of the proximal stump (Fig. 25.1Ciii) have been described. In the mammalian CNS, the failure to regenerate damaged axons is a major clinical impediment to recovery after brain and spinal cord injuries; hence, the possibility that some aspects of this failure are shared with invertebrates, where it can be studied in a simple model system, is interesting and potentially exciting.

Table 25.1 Axon Response to Acute Injury in Different Models





Degeneration of Distal Stump

Regeneration of Proximal Stump


Drosophila melanogaster (fruit fly)

Olfactory receptor neuron (adult)



Fragmentation starts 1 day after injury and completes by a week


Hoopfer et al., 2006; Macdonald et al., 2006

Motor neuron (larva)

Crush/laser surgery


Fragmentation initiates 4 hours after injury and continues for up to a day

Forming growth cones 10–14 hours after injury, followed by axon growth

Xiong et al., 2010; Xiong et al., 2012; Mishra et al., 2013

Sensory neuron (adult wing)

Transection/laser surgery


Fragmentation initiates within a day and completes by around a week

(Laser surgery): regeneration initiates by 3 days

Fang et al., 2012; Neukomm et al., 2014; Soares et al., 2014

DA sensory neuron (larva)

Laser surgery


Fragmentation starts 6 hours after injury for Class I neurons

Regeneration observed for Class IV neurons (3 days after injury) and some Class I neurons (1–3 days) but not for Class III; limited CNS regeneration versus prominent peripheral regeneration

Stone et al., 2010, 2012; Song et al., 2012

Mushroom body neuron (larva, pupa, and adult)

Disassociation (in vitro)



Neurites sprout 1 day after dissociation in culture

Marmor-Koller & Schuldiner, 2016

Motor neuron (larva)

Disassociation (in vitro)



Neurites sprout 2–3 days after dissociation in culture

Lu et al., 2015

sLNv neuron (adult brain explant)

Transection by microdissection device (in vitro)


Fragmentation starts 1 day after injury

Filopodia sprouts observed 2 days after injury in a portion of axons while a majority of axon do not regenerate

Ayaz et al., 2008

Periplaneta americana (cockroach)

Motor neuron



Reinnervation by 4 weeks and recovery of circuits by 7–9 weeks

Bodenstein, 1957; Case, 1957

Acheta domestica (cricket)

Medial giant interneuron (giant fiber system)

Crush or cut


Sprouting; crush near cell body induces dendritic branching

Roederer & Cohen, 1983

Sensory neuron

Transection of cerci


Sprouting; target to the same giant fiber

Edwards & Sahota, 1967

Procambarus clarkii (crayfish)

Motor neuron

Crush or transection


Not oberved for 3 months

Fusion occurs; a majority of injured axons establish reconnection by 30 days

Hoy et al., 1967

Caenorhabditis elegans

Mechanosensory neuron (ALM, PLM, AVM); chemosensory neuron (ASH, ASJ); GABAergic motor neuron (DD, VD); cholinergic motor neuron (DA/DB); HSN motor neuron

Laser surgery


Axons become thin, beaded, and invisible in ALM/PLM/DD neurons 24 hours for L1 or several days for L3 after injury

Axon growth 12–24 hours after surgery; fusion observed with some mechanosensory neuron

Yanik et al., 2004; Wu et al., 2007; Gabel et al., 2008;Chen & Chisholm, 2011; Chung et al., 2016; Nichols et al., 2016; for details, see Hammarlund & Jin, 2014

GABAergic motor neuron (β-spectrin mutant)




70% axons regenerate

Hammarlund et al., 2009

Aplysia californica (California sea hare)

Sensory neuron and motor neuron

Dissociated or crush (in vitro)

Degeneration does not occur for hours

Sprouting and regrow axons

Schacher & Proshansky, 1983; Dash et al., 1998

Sensory neuron



Reinnervation and recovery of reflex behavior within 2–3 weeks

Dulin et al., 1995; Noel et al., 1995; Steffensen et al., 1995

Buccal motor neuron



Degeneration observed 8 weeks after injury

Reinnervation to targets observed 3 weeks after injury

Ross et al., 1994

Lymnaea Stagnails (Great pond snail)




Sprouting more prominent when injured closer to soma; synaptic connections restored 3–6 days after injury

Allison & Benjamin, 1985; Benjamin & Allison, 1985

Helodrilus caliginosus and Lumbricus terrestris (earthworm)

Giant axon



Fusion with high accuracy

Hall, 1921; Birse & Bittner, 1976

Hirudo medicinalis (leech)




Starts around 1 month after injury

Sprouting and following the existing distal stump to innervate the target by 1 month and functional recovery by 2 months

Frank et al., 1975; Muller & Carbonetto, 1979

CNS, central nervous system; NA, not applicable; PNS, peripheral nervous system.

Whereas the proximal stump can either regenerate or fail to regenerate, the common fate of the “distal stump,” which is no longer connected to the cell body, is to degenerate (Fig. 25.1Biii). Most Drosophila neurons behave similarly to mammalian neurons by initiating axonal degeneration quite rapidly (within a day) after injury. This fast process, in theory, may allow for a “replacement” by new growth from a regenerating “proximal stump” (Fig. 25.1Ci). However, an injured “distal stump” has been observed in some invertebrate animals (crayfish and leech) to persist for months after injury with no signs of degeneration (Hoy et al., 1967; Frank et al., 1975). In these cases, as well as in C. elegans, a process of refusion between the two separated stumps (Fig. 25.1Cii) has been observed (Hall, 1921; Hoy et al., 1967; Birse & Bittner, 1976; Muller & Carbonetto, 1979; Neumann et al., 2011). Whether the repair is achieved by replacement (in Fig. 25.1Ci) or fusion (Fig. 25.1Cii), the two processes of axonal degeneration and regeneration need to be coordinated.

Axonal damage can also occur in other scenarios of injury and stress, which we define here as “chronic injuries.” These scenarios include prolonged presence of a neurotoxin (such as taxol and colchicine, which induce axonal loss) or the presence of a mutation, which induces a persistent “stress” to the integrity and function of the axon (Fig. 25.1D). Many mutations that are known to cause inherited neurodegenerative disorders in humans, when expressed in vertebrate and invertebrate models, lead to loss of synapses (Fig. 25.1Ei) and/or axons (Fig. 25.1Eii) prior to neuronal death (Saxena & Caroni, 2007; Charng et al., 2014; Casci & Pandey, 2015).

A unifying feature of both acute and chronic models of axonal damage is the impairment of intracellular transport processes within axons. The transport of organelles and proteins in axons relies upon the action of motor proteins, which physically carry their cargo by walking upon microtubule tracts. In acute axonal injuries, the delivery of molecules from the cell body to the distal axon is irreversibly blocked due to the fact that they are no longer physically connected. In chronic models, although the connection remains, the process of axonal transport is also thought to be persistently impaired (Fig. 25.1D). This has been shown by altered movement of fluorescently tagged organelles such as mitochondria or synaptic vesicle precursors, or by accumulations of organelles in axons or cell bodies (Millecamps & Julien, 2013). Mutations that disrupt the cytoskeleton, comprised of the microtubule tracts and associated molecules, often lead to degeneration of axons and/or synapses (Pielage et al., 2005, 2008; Schaefer et al., 2007; Bounoutas et al., 2011; Hsu et al., 2014; Ma et al., 2014; Neumann & Hilliard, 2014).

In contrast to acute injuries that completely disconnect cell bodies with their synaptic targets, neurons in a chronic injury condition have the chance to try to adapt to the stress and make their axons more resilient to degeneration (Fig. 25.1Eiii). Although adaptive changes and mechanisms are still very poorly characterized, they may potentially entail an induced expression of chaperones and transport of additional cytoskeletal components into axons. Depending upon the severity and duration of the stress, this response may or may not be enough to maintain the axon and/or to prevent cell death.

The processes of degeneration, regeneration, and adaptation are all interesting from the perspective of human health. The mechanisms that neurons engage to either delay or accelerate axonal and synaptic loss and repair after injury could be valuable therapeutic targets for treatment of traumatic injuries as well as neuropathies and even potentially neurodegenerative diseases in which axonal loss occurs.

Axon and Synapse Loss

The process of axonal degeneration after acute injury (Fig. 25.1Biii) is highly stereotyped and is termed Wallerian degeneration (WD), based on its first description by Augustus Waller in 1851. For a period of time after injury (termed the “lag phase”) the distal stump of the axon remains intact and is able to propagate action potentials (Lubińska, 1977; Beirowski et al., 2005). In most cases, the lag phase is then followed by a rapid fragmentation phase, in which the axon breaks into many individual pieces, which are then phagocytosed by glia and immune cells (Bhatheja & Field, 2006). WD likely entails a cell autonomous chain of events that occur within the distal axon itself and, hence, can be considered as a “self-destruction” pathway, akin to (p. 581) apoptosis. However, WD appears to involve a molecular pathway that is quite distinct from apoptosis (Deckwerth & Johnson, 1994; Finn et al., 2000). The pathway itself and its molecular players are still at the early phases of being described, and studies in the invertebrate model organism Drosophila have led the way in this exciting area of study.

In Drosophila, multiple approaches for inducing axonal injury and studying WD have now been described (Table 25.1; also see Fang & Bonini, 2012). Combined with the powerful genetics of this model organism, such approaches provide an ideal vehicle to uncover vital molecular mediators of the WD pathway. Many of the injury approaches (such as cutting an antenna or a wing) are simple to carry out, enabling genetic screens to identify molecules whose function is required for degeneration to occur. For some of the approaches (including laser-directed transection and nerve crush injuries in peripheral neurons of larvae that are semitransparent) the process of WD is amenable to detailed study of the cellular changes during the course of WD, including changes in intracellular calcium, mitochondria, and cytoskeleton (Avery et al., 2012; Fang et al., 2012; Kitay et al., 2013; Mishra et al., 2013). Here we briefly highlight some of the discoveries made in Drosophila that have strongly influenced our understanding of WD.

Central Molecular Regulators of Wallerian Degeneration


A heroic large-scale genetic mosaic screen in Drosophila led to the discovery of dSarm (Drosophila Sterile alpha and HEAT/Armadillo motif containing), as a critical molecular player in WD (Osterloh et al., 2012). Neurons that are mutant in dsarm fail to degenerate distal axons after axonal injury. This role in promoting degeneration was then shown to be conserved in mammalian Sarm1 and in C. elegans tir-1 (Osterloh et al., 2012; Vérièpe et al., 2015). Although Sarm1/tir-1/dSarm (which we refer to here simply as Sarm) has been previously implicated in innate immunity and neuronal development (Couillault et al., 2004; Liberati et al., 2004; Chuang & Bargmann, 2005; Chen et al., 2011), a role in axonal degeneration would have never been guessed, highlighting the importance of forward genetic approaches. With Sarm’s central role in degeneration revealed, current work is now focused upon its mechanism. Domain analysis suggests that dimerization of Sarm’s TIR domains is sufficient to activate WD in uninjured axons, and this activation leads to a rapid rundown of intracellular metabolites NAD + and ATP (Gerdts et al., 2015; Sasaki et al., 2016; Summers et al., 2016). Much work remains to be done to understand the molecular events that lead to the activation of Sarm and its downstream actions.


In contrast to the unanticipated discovery of Sarm, a potential role for Nicotinamide mononucleotide adenylyl transferase enzyme (Nmnat) had been suspected for years, ever since a gain-of-function mutation in the Nmnat1 enzyme was fortuitously discovered in the background of a mouse strain. WD fails to occur in these mutant mice, and this effect can be recapitulated in Drosophila neurons, by overexpressing the Drosophila dNmnat enzyme (MacDonald et al., 2006).

Loss-of-function studies in both Drosophila and mice suggest that endogenous versions of Nmnat enzymes in healthy neurons play a protective role to inhibit degeneration (Zhai et al., 2006; Gilley & Coleman, 2010; Wen et al., 2011; Fang & Bonini, 2012; Hicks et al., 2012; Rallis et al., 2013; Sasaki et al., 2015). In many of these studies, depletion of Nmnat function leads to spontaneous axonal degeneration even in the absence of injury. To explain these observations, it has been proposed that Nmnat is an axonal “survival” factor. Drosophila Nmnat and mammalian Nmnat2 are continuously transported into distal axons, where the protein is then rapidly turned over (Gilley & Coleman, 2010; Xiong et al., 2012; Milde et al., 2013; Fig. 25.2). Once disconnected from the cell body, the distal stump loses the supply of Nmnat from the cell body and its essential but still poorly understood survival function, which leads to the initiation of degeneration.

What exactly the Nmnat enzymes do to maintain axon integrity is still not clear, and this is the subject of much investigation and discussion (Ali et al., 2013). It is clear that Nmnat enzymes need to localize in the cytosol (Fig. 25.2A), and they become more potent at protecting axons from degeneration when they are targeted to axons (Avery et al., 2009; Beirowski et al., 2009; Sasaki et al., 2009). The enzymatic activity for NAD + synthesis also appears to be important (Araki et al., 2004; Jia et al., 2007; Sasaki et al., 2009). In line with this, a recent study has linked Sarm1’s role in degeneration to a rapid rundown in NAD + (Gerdts et al., 2015). However, whether inhibiting NAD rundown is a direct action of the Nmnat enzymes has been difficult to address, and some studies have linked other metabolites on (p. 582) the NAD biosynthesis pathway with axonal degeneration (Conforti et al., 2014).

Mechanisms of Axonal Degeneration and RegenerationLessons Learned From Invertebrates

Figure 25.2 Molecular mechanisms during axon and synapse regeneration and degeneration. (A) In uninjured neurons, several critical factors for regeneration or degeneration are present in axon and synapses. These include Nmnat (orange circle) and DLK (blue circle), which are transported (associated with vesicles) in axons and which are also turned over in axons, most likely in distal axon and synaptic locations by the Hiw ubiquitin ligase complex (gray squares). (B) Upon injury, the DLK kinase becomes activated, and in the proximal stump it can signal retrogradely to the cell body to initiate a transcriptional response (green nucleus). Retrograde signaling by DLK and other factors (indicated with stars; Rishal & Fainzilber, 2014) are required for later axon regrowth. In the distal stump, “survival” factors such as Nmnat become depleted because their turnover continues while the supply of new molecules from the cell body is cut off. Sarm (triangle) becomes activated in the injured distal stump and promotes a rapid rundown in intracellular NAD + and ultimately axonal degeneration.

A quite different idea is that Nmnat performs an additional function that is separate from NAD + synthesis, by acting as a molecular chaperone, akin to the function of heat shock proteins (Ali et al., 2013). This idea builds upon observations that Nmnat transgenes that are nonfunctional for NAD synthesis activity can still have protective effects when overexpressed in Drosophila neurons (Zhai et al., 2006, 2008). Further, endogenous Nmnat isoforms become upregulated in several models of protein-folding disorders, and in these cases Nmnat protein is observed to colocalize with protein aggregates. In addition, Nmnat can facilitate the folding of denatured luciferase in vitro, possibly via its ATPase domain (Zhai et al., 2008; Ali et al., 2011, 2012, 2016). Although it is challenging to nail down a chaperone function in vivo, the idea remains attractive since other known chaperones (p. 583) (such as TBCE, CSP, and Hsp70) are required for continued axon and synapse integrity (Fernández-Chacón et al., 2004; Schaefer et al., 2007; Rallis et al., 2013).

Potential Sarm and Nmnat-Independent Pathways

Although axonal degeneration usually initiates within hours of injury in Drosophila and mammalian neurons, crayfish and leech axons have been observed to persist for months after injury (Hoy et al., 1967; Frank et al., 1975; Ballinger & Bittner, 1980). This implies the existence of mechanisms that have been adopted by some animals to maintain the integrity of distal “stump” and/or inhibit degeneration. Do these axons fail to activate Sarm? If so, how and why is this change achieved in these neurons? Or are there additional pathways involved? Interestingly, a recent study has documented WD in C. elegans, which occurs independently of any manipulation to TIR-1, the C. elegans homologue of Sarm, or Nmnat (Nichols et al., 2016). This suggests that Sarm may not universally promote degeneration in all neuron types, or it may be utilized to different degrees in different contexts. A revisit to old observations of axonal degeneration in nonmodel organisms with contemporary techniques may help to reveal the origin and evolution of the degeneration program.

Adaptive Mechanisms to Chronic Stress

Similarly to WD after injury, chronic stressors (such as cytoskeletal toxins and/or mutations) can also cause axonal degeneration (Fig. 25.1D and 25.1Eii). Because in most cases degeneration is inhibited by manipulations that increase Nmnat activity, it is thought that degeneration in these models shares a common underlying mechanism with WD (Coleman & Freeman, 2010). However, neurons in different injury models exhibit various tolerances to chronic stressors (Conforti et al., 2014). We posit that some of these differences are determined by whether the cell is able to make an adaptive response to delay the degeneration process (Fig. 25.1Eiii).

Some recent studies in Drosophila suggest that nuclear signaling pathways may become engaged in response to chronic stress and damage with an output that serves to enhance and/or maintain axon integrity. An important example is the upregulation of Nmnat expression and alternative splicing of Nmnat isoforms, which has been observed in response to proteotoxic stress, heat shock stress, and hypoxia (Zhai et al., 2008; Ali et al., 2011; Ruan et al., 2015). An additional pathway, discussed later, is the DLK signaling pathway, which becomes activated in injured axons. In Drosophila motoneuron axons, this pathway induces cellular changes that delay the process of WD, such that an axon that has been injured once becomes more resilient to degeneration after a second injury (Xiong & Collins, 2012). The manifestation of this protective response only occurs for the proximal stump but not the distal stump, likely because the process involves the expression and transport of new molecules into axons.

It is interesting that both Nmnat and the DLK kinase share commonalities in their regulatory mechanisms. First, both are transported in axons and associated with Golgi-derived vesicles (Fig. 25.2A). Palmitoylation allows for this localization and is required for the rapid turnover of Nmnat2 (Milde et al., 2013) and the function of DLK (Holland et al., 2015) in mammalian neurons. Whereas this has yet to be tested in invertebrate neurons, Drosophila Nmnat and DLK proteins contain palmitoylation consensus sequences. Second, the protein turnover of both Nmnat and DLK are regulated in both Drosophila and mammalian neurons by a conserved ubiquitin ligase complex, whose signature component is a highly conserved PHR protein, named Highwire (Hiw) in Drosophila, RPM-1 in C. elegans, and PAM in mice (Nakata et al., 2005; Collins et al., 2006; Wu et al., 2007; Xiong et al., 2010, 2012; Babetto et al., 2013; Brace et al., 2014). Hiw therefore becomes an intriguing regulator (and is perhaps a coordinator) of adaptive responses to axonal damage (Fig. 25.2).

Finally, it is interesting to consider that many chronic paradigms of axonal injury can originate or manifest at presynaptic terminals. Several mutations that disrupt synaptic structure lead to axon and/or synapse degeneration (Fernández-Chacón et al., 2004; Pielage et al., 2005; Burgoyne & Morgan, 2011; Wishart et al., 2012). Also, in many neuropathies, the most terminal connections of the axon are lost first, suggesting a “dying-back” mechanism, which may be initiated by a toxic stimulus at the synapse (Yaron & Schuldiner, 2016). Because it has been suggested that a degeneration program may be initiated and/or restrained at synapses (Fig. 25.1Ei), the Hiw ubiquitin ligase complex gains even further cache, since Hiw and its homologues are known to localize to presynaptic terminals (Schaefer et al., 2000; Wan et al., 2000; Zhen et al., 2000) and can inhibit synaptic degeneration in at least one chronic paradigm (Massaro et al., 2009).

(p. 584) Axon and Synapse Repair

For a damaged axon to grow (or regrow) it needs to have a growth cone (Tom et al., 2004; Ertürk et al., 2007). Many early studies in cultured neurons from Aplysia and cockroach (whose giant axons are very amenable to imaging and recording after injury in culture) have helped to describe cellular events that direct a transformation of a severed axonal stump into a growth cone: Calcium influx triggered by the injury itself directs axon membrane resealing at the injury site (Yawo & Kuno, 1985; Strautman et al., 1990; Davenport & Kater, 1992; Spira et al., 1993; Ziv & Spira, 1995) and activation of local calcium-regulated proteases, which promote reconstructuring of neurofilaments and microtubules close to the stump ending (Gitler & Spira, 1998, 2002; Spira, 2003). These events ultimately lead to the formation of a growth cone that has dynamic lamelopodia and filopodia (Baas & Heidemann, 1986; Ertürk et al., 2007; Schaefer et al., 2008; Hellal et al., 2011).

The ability of the growth cone to direct new axonal growth is associated with transcriptional and translational changes in the cell body. These changes are induced by signaling pathways that become activated in injured axons, and this “injury signaling” appears to be mediated, at least in part, by molecules which are physically transported in axons (Hanz & Fainzilber, 2006). Early studies in Aplysia led to the identification of several proteins that are retrogradely transported specifically in injured neurons (Ambron et al., 1992; Schmied et al., 1993; Schmied & Ambron, 1997; Zhang et al., 2000; Sung et al., 2001; Sung et al., 2004). These findings inspired later studies in mammalian peripheral sciatic nerves, which have revealed critical components of “retrograde signaling” (Lindwall & Kanje, 2005; Perlson et al., 2005; Ben-Yaakov et al., 2012).

Upon this foundation of knowledge from Aplysia and other invertebrate studies (Table 25.1), our understanding of molecular pathways underlying regeneration was brought to an exciting new level in studies using C. elegans and Drosophila, which have enabled genetic screens and genetic dissection of pathways required for axonal growth after injury. (Detailed review can be found in Hammarlund & Jin, 2014; Brace & DiAntonio, 2016; and Byrne & Hammarlund, 2016). We’ll focus our discussion on what is perhaps the most important discovery, the elucidation of the DLK/Wallenda signaling pathway, which detects and initiates responses to axonal damage.

DLK/Wallenda Is Essential for Axonal Regeneration

A role for the DLK kinase in axonal regeneration was first identified in a cleverly designed genetic screen in C. elegans (Hammarlund et al., 2007). The screen was built upon the observation that axons in β-spectrin mutants break spontaneously; however, in response they form new growth cones. Hammarlund and colleagues screened for mutants that failed to do so and identified a signaling cascade governed by DLK kinase, which is essential for the transformation of axonal breaks into new growth cones. Importantly, dlk mutants have no obvious phenotype in axonal outgrowth during development (Nakata et al., 2005; Collins et al., 2006; Miller et al., 2009). These findings suggest that DLK carries out a specific postdevelopmental role in regulating responses to axonal injury.

Several points emphasize the importance of DLK as a central player in regulating the ability of injured axons to regenerate. First, the requirement for DLK in axonal regeneration appears conserved across multiple neuron types in C. elegans, Drosophila, and also in mammalian PNS neurons, which regenerate (Yan et al., 2009; Xiong et al., 2010; Pinan-Lucarre et al., 2012; Shin et al., 2012). Second, DLK functions as an upstream regulator of a MAP Kinase signaling cascade. A number of observations suggest that it is transported in axons and becomes acutely activated after axonal damage. Activated DLK or its downstream targets give rise to retrograde signaling to initiate a nuclear response; hence, DLK appears to function as a regulator of signaling molecules that are retrogradely transported in axons (Yan et al., 2009; Xiong et al., 2010; Bounoutas et al., 2011; Shin et al., 2012; Watkins et al., 2013). In mammalian neurons DLK has also been implicated in other processes that seem to be quite distinct from axonal regeneration: DLK promotes neuronal death after nerve growth factor withdrawal (Ghosh et al., 2011), death of retinal ganglion cells after injury of their axons within the optic nerve (Watkins et al., 2013; Welsbie et al., 2013; Fernandes et al., 2014), and neuronal death in cellular models of excitotoxicity (Pozniak et al., 2013). A shared component of all of these processes that activate DLK is the presence of stress and/or damage to the axon/synapse. The current unified view in the field is that DLK functions as a “sensor” of axonal damage, with the downstream consequences of its activation varying depending upon context.

(p. 585) The amenability of Drosophila and C. elegans to combining mutagenesis and genetic interaction analysis has provided insight into the cellular pathways and processes that appear to regulate DLK’s signaling functions in neurons. In C. elegans, a mechanism for direct regulation by intracellular calcium has been described (Yan & Jin, 2012): An isoform of DLK binds to and inhibits the full-length form of DLK, and this inhibitory binding is released in conditions that elevate intracellular calcium. However, the sequences that mediate these interactions are not conserved for mammalian or Drosophila DLK, so there are likely additional important mechanisms for its regulation. Indeed, a recent study has identified the cAMP effector kinase PKA as an important upstream activator of DLK in Drosophila and mammalian neurons (Hao et al., 2016).

Interestingly, studies in all model organisms have noted DLK’s relationship with microtubule and actin cytoskeleton. Induced cytoskeletal stresses, such as treatment with taxol or cholchicine, or mutations in cytoskeletal components (tubulin) or regulators (microtubule associated protein), lead to changes of structure and expression in neurons. Interestingly, many of these changes are suppressed when DLK is mutated (Bounoutas et al., 2011; Valakh et al., 2013; Chen et al., 2014; Marcette et al., 2014; Richardson et al., 2014). These findings imply that DLK acts downstream to these manipulations to cytoskeleton, and indeed, DLK signaling becomes activated in mammalian neurons that are treated with cytoskeletal destabilizing agents (Valakh et al., 2013, 2015). Complementary to this point, it appears that a downstream effect of DLK signaling is the induction of alterations in cytoskeleton organization (Lewcock et al., 2007; Hendricks & Jesuthasan, 2009; Eto et al., 2010; Feltrin et al., 2012; Klinedinst et al., 2013) and tubulin expression (Nadeau et al., 2005). These findings place DLK as both a sensor and effector to regulate cytoskeleton dynamics.

Is Regeneration “Programmed”?

During development, axons respond to specifically placed cues to direct their growth correctly, often over a long path that involves many intermediate targets, to find their appropriate synaptic targets. Is the same developmental process re-engaged for regeneration? The answer is likely both Yes and No. Extracellular factors are important for both development and regeneration. However, in mammalian PNS regeneration, where motor neurons can reinnervate their targets accurately after crush (Nguyen et al., 2002), nerve growth factors are released by Schwann cells and microphages rather than targets, which are the main source during development. Schwann cell “tubes” can physically confine axon outgrowth during regeneration, but not during development (Bhatheja & Field, 2006; Scheib & Höke, 2013). In invertebrates, little is known about the mechanisms that govern pathfinding and reinnervation. However it is striking that reconnection to original targets after axonal injury has been observed in many species, including cockroaches, crickets, leech, crayfish, Aplysia, and snails (Bodenstein, 1957; Case, 1957; Edwards & Sahota, 1967; Hoy et al., 1967; Muller & Carbonetto, 1979; Allison & Benjamin, 1985; Benjamin & Allison, 1985). In instances where regeneration has been studied on a molecular level (in C. elegans and Drosophila), pathways that are distinctly required for regeneration and not for development have been most notable and well characterized. These include the DLK/Wallenda and PTEN/PI3K signaling pathways. In C. elegans, DLK is dispensable for axon outgrowth during development (Hammarlund et al., 2009). Likewise, in the Drosophila mushroom body, neither PI3K nor DLK is required for axon outgrowth during development (Marmor-Kollet & Schuldiner, 2016). On the other hand, unc-40/DCC is required for axon initial outgrowth during development (Chan et al., 1996; Keino-Masu et al., 1996), but not axon regeneration (Gabel et al., 2008). These “regeneration-specific” pathways suggest that axon outgrowth and innervation after injury are intrinsically and uniquely programmed.

The functional endpoint for axonal regeneration is to re-establish a functional circuit. Unless refusion occurs (as in Fig. 25.1Cii), this requires a newly formed axon to reform lost synaptic contacts (Fig. 25.1Ci). Although mechanisms that promote the growth of injured axons have been the topic of much investigation, there are very few studies to characterize whether and how synapses can be formed by regenerating axons. Studies of mammalian NMJ regeneration have provided insights into roles of extracellular matrix proteins (Skouras et al., 2011). But little is known about the intracellular pathways in neurons that promote regeneration of synapses. Synapse regeneration may share similarities in “programming” with axon regeneration, with mechanisms that are both shared and distinct from developmental pathways. The field simply needs more studies and more information (p. 586) on this topic. Invertebrate model systems in which functional regeneration can occur (which can result in readily screenable behavioral phenotypes) have important contributions to make for these important future questions.


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