Due to the COVID-19 crisis, the transition into subscription mode of the Oxford Research Encyclopedia of Neuroscience has been temporarily postponed. Please watch this space for updates as we work toward launching in the near future. Visit About to learn more, meet the editorial board, or learn how to subscribe.

Show Summary Details

Page of

PRINTED FROM the OXFORD RESEARCH ENCYCLOPEDIA, NEUROSCIENCE (oxfordre.com/neuroscience). (c) Oxford University Press USA, 2020. All Rights Reserved. Personal use only; commercial use is strictly prohibited (for details see Privacy Policy and Legal Notice).

date: 09 April 2020

Enhancing the Regeneration of Neurons in the Central Nervous System

Summary and Keywords

Injured axons fail to regenerate in the adult mammalian central nervous system, representing a major barrier for effective neural repair. Both extrinsic inhibitory environments and neuron-intrinsic mechanisms contribute to such regeneration failure. In the past decade, there has been an explosion in our understanding of neuronal injury responses and regeneration regulations. As a result, several strategies have been developed to promote axon regeneration with the potential of restoring functions after injury. This article will highlight these new developments, with an emphasis on cellular and molecular mechanisms from a neuron-centric perspective, and discuss the challenges to be addressed toward developing effective functional restoration strategies.

Keywords: axon regeneration, DLK, mTOR, injury responses, cytoskeleton, microtubule, growth cone, optic nerve injury, spinal cord injury


Enhancing the Regeneration of Neurons in the Central Nervous System

Figure 1. Different steps for regenerating axons to regenerate and reconnect with their targets. After axotomy, a set of injury signals generated in injured axons trigger injury responses in the compartments of axons and their cell bodies. This occurs naturally to most injured axons. However, depending on the regenerative competence, injured neurons exhibit distinct outcomes. For successful regeneration, injured axons need to reorganize local cytoskeletons and subcellular structures and their cell bodies need to turn on axon growth programs. It is possible that multiple signals and programs are involved in these processes. Thus combinatorial manipulations may allow optimized regenerative growth and reconnection with their targets.

In the adult mammalian central nervous system (CNS), spontaneous regeneration usually does not occur (Byrne & Hammarlund, 2017; Hilton & Bradke, 2017; Rasmussen & Sagasti, 2017). Although much progress has been made on deciphering this lack of regenerative ability, there is still no effective neural repair strategy for CNS injuries such as spinal cord injury, traumatic brain injury, or stroke. One underlying cause is the extraordinary complexity of interconnected cellular and molecular injury responses that involve different neuronal compartments. In general, axonal injury elicits an injury signal that propagates from the site of injury to the cell body of the neuron. Such signals often trigger a set of homeostatic adaptations. In some cases, axon insults could lead to cell death. However, preventing neuronal death after a CNS insult is not sufficient to promote axonal regeneration. For regrowing axons, the inhibitory injury site is often a tremendous hurdle. Even if this is overcome, they will face further challenges in finding their way to their appropriate targets. Following this chronology of events from the injury to the functional connection of regenerating axons (Figure 1), we will go through the current understanding of the mechanisms of axonal regeneration from a neuron-centric perspective. Readers are referred to recent reviews that focus on neuron-extrinsic influences (Geoffroy & Zheng, 2014; Silver, Schwab, & Popovich, 2015).

Injury Response

Neurons have a unique morphology with a cell body or soma and elaborated processes including both axons and dendrites. In humans, axons could project up to one meter from their cell bodies. Such structural properties require a tight regulation of the intra-cellular communication between different subcellular compartments. For the interaction between axon and soma, the cell body adjusts the gene expression programs based on information received from the axons. Conversely, the axon needs to know how the cell body is adapting to a situation such as injury. While the injury responses have been mainly studied in more permissive systems such as peripheral sensory neurons (Lieberman, 1971) and C. elegans neurons (Hammarlund & Jin, 2014; Yanik et al., 2004), the mechanisms of the generation and propagation of injury signals appear to be conserved in CNS (Bradke, Fawcett, & Spira, 2012; He & Jin, 2016).

Upon an axotomy, three sequential steps, probably to some extent overlapping, have been proposed as a model for injury site–soma communication (Abe & Cavalli, 2008). First, the breakage of the axonal membrane induces a disruption of membrane potential. Second, the normal flux of retrograde transport is interrupted. Finally, signaling molecules will be actively transported from the injury site to the neuron soma with axonal transport machinery (Ambron & Walters, 1996; Rishal & Fainzilber, 2013). Among these, a key event is the locally activated kinases which phosphorylate a variety of substrates and trigger the retrograde transport (Cavalli, 2005; Hanz et al., 2003; Lindwall & Kanje, 2005; Perlson et al., 2005; Sung, Chiu, & Ambron, 2006; Yudin et al., 2008).

Conducted on C. elegans beta spectrin mutant (unc-70) whose axons break because of locomotion-induced mechanical stress, an elegant RNAi screen revealed a key sensor of axon injury called the dual leucine-zipper kinase (DLK) (Hammarlund, Nix, Hauth, Jorgensen, & Bastiani, 2009). DLK was also independently identified as a crucial axon regeneration regulator (Yan, Wu, Chisholm, & Jin, 2009). As a MAP kinase kinase kinase (MAPKKK), DLK is locally activated by axonal insults in a calcium-dependent manner and activates its substrates MAP kinase kinase MKK-4 and in turn the MAP kinase PMK-3, therefore triggering the signaling cascade. These activated signaling components are retrogradely transported back to the cell bodies to initiate regenerative programs. Indeed, DLK knock-down or knock-out inhibits axonal regeneration and its over-expression improves growth cone initiation. Interestingly, it appears that the DLK effect on axonal regeneration was linked to its capacity to stabilize CCAAT-enhancer-binding protein 1 (CEBP-1) mRNA to ensure local translation in axons (Yan et al., 2009). In Drosophila, the deletion of the actin and microtubule protein binding spectraplakin short stop (Shot) activates the early injury response protein DLK (Valakh, Walker, Skeath, & DiAntonio, 2013), consistent with the notion of DLK as an axonal sensor for injury and disorganization. Importantly, the role of DLK is conserved in mammals in both PNS (peripheral nervous system) (Shin et al., 2012) and CNS (Watkins et al., 2013).

Similar to the DLK/MAP pathway, cytokine-dependent Janus kinase (JAK)/Signal transducer and activator of transcription (STAT) signaling is another avenue to relay injury signals to the cell body (Sun & He, 2010). In this case, the signal, cytokines, such as leukemia inhibitory factor (LIF) (Banner & Patterson, 1994; Sun & Zigmond, 1996), interleukin-6 (IL6) (Fregnan, Muratori, Simões, Giacobini-Robecchi, & Raimondo, 2012; Yang, Wen, Ou, Cui, & Fan, 2012), and ciliary neurotrophic factor (CNTF) (Park et al., 2009; Smith et al., 2009), could be released from neighboring cells after injury, and activate neuronal signaling in a non-cell autonomous way. Specifically, they will interact with their specific receptors on neuronal surface and activate JAK kinases and phosphorylated signal transducer and activator of transcription 3 (STAT3) will be transported to the nucleus (Curtis et al., 1993, 1994; O’Brien & Nathanson, 2007).

For all aspects of injury responses, calcium entry triggered by axonal injury is pivotal. It has been well studied in all species that a calcium wave propagates from the injury site and contributes to other key cellular regeneration events including the activation of pro-regenerative signaling pathways, cytoskeletal remodeling, and growth cone formation (Chierzi, Ratto, Verma, & Fawcett, 2005; Ghosh-Roy, Wu, Goncharov, Jin, & Chisholm, 2010; Gitler & Spira, 1998; Sun et al., 2014). In C. elegans, the extent of regeneration is positively correlated with the amplitude of the initial calcium wave (Ghosh-Roy et al., 2010) and internal calcium release from the endoplasmic reticulum is necessary for axonal regeneration (Sun et al., 2014). However, calcium overload could cause neuronal death (Pivovarova & Andrews, 2010) and therefore its entrance and release in the axons need to be tightly regulated. For example, after a spinal cord contusion, the better the axons mitigate the influx of calcium, the less probability they have to be affected by axonal degeneration (Williams et al., 2014). Interestingly, deletion of the gene encoding the Alpha2delta2 subunit of voltage-gated calcium channels (VGCCs) promotes axonal growth in vitro and dorsal column sensory axon regeneration after spinal cord injury (Tedeschi et al., 2016). Along the same line, calcium channel inhibitors prevent injury-induced intra-axonal calcium increase and attenuate axonal degeneration in retinal ganglion cells (RGCs) (Knöferle et al., 2010) and promote limited axonal regeneration in vivo (Ribas, Koch, Michel, Bähr, & Lingor, 2016). As many cellular events are tightly regulated by calcium, future studies should monitor calcium dynamics with high temporal and spatial resolution and assess their specific outcomes.

Regeneration Initiation

With incoming injury signals, an injured neuron has to make a decision to either initiate or abort axonal regrowth, which involves a number of molecular and cellular responses in both axonal and soma compartments. This is influenced by the axonal immediate environment in which reactive glial/fibrotic scars and myelin debris could inhibit or support axon regeneration (He & Koprivica, 2004; Yiu, 2003; Yiu & He, 2006). On the other hand, the other contributing factor is the intrinsic regenerative ability associated with individual neurons (He & Jin, 2016). Deciphering this regenerative growth program is currently under intense investigation.

As an essential step for axon growth, a lesioned axonal tip needs to be reorganized and reform the growth cone (Bradke et al., 2012). Although the morphology of regenerated growth cones could be very different from their counterparts during early development, the basic function is the same: as a leading driving force for axonal extension. In the CNS, injured axons often form retraction bulbs with disorganized microtubules and accumulation of mitochondria (Ertürk, Hellal, Enes, & Bradke, 2007), an indicator of regenerative failure. In this regard, the stabilization of microtubules and actins appears to be necessary for growth cone reformation (Bradke et al., 2012) and this appears to be evolutionarily conserved in different species (Chen et al., 2011; Ertürk et al., 2007; Nawabi et al., 2015; Ruschel et al., 2015). Importantly, microtubule-stabilizing compounds such as taxol, epothilone B and D, widely used for cancer chemotherapy, have been shown to promote axon regeneration and some degree of functional recovery in spinal cord injury models (Hellal et al., 2011; Ruschel & Bradke, 2018; Ruschel et al., 2015; Sandner et al., 2018). Interestingly, microtubule stabilizing agents have dual effects on axons (intrinsic) and scar tissues (extrinsic). In addition to the axonal terminal, many types of CNS axons exhibit an injury-induced retrograde retraction with disruption of the f-actin and microtubules cytoskeleton away from the injury site (Chen et al., 2011, 2015; Erez & Spira, 2008). Thus, preserving the cytoskeleton along the axonal shaft is also important for the regeneration process.

Recent studies have begun to reveal molecular mechanisms for such cytoskeleton regulation in injured axons and their correlation with axon regeneration. Besides the well-characterized role of calcium (Bradke et al., 2012), several other molecules have been implicated in regulating cytoskeleton stabilization and growth cone reformation after axonal injury. For example, injury down-regulates the expression of the members of the doublecortin-like kinases family (DCLK1/2, DCX), which all have the ability to bind microtubules and actin. Forced expression of these molecules promotes growth cone formation and axonal regeneration in RGCs, for which both functional domains for binding microtubules and actin are required to work together. Intriguingly, other types of microtubule-binding proteins such as EB3 and Tau fail to mimic the effects of DCX members (Nawabi et al., 2015). These examples place the cytoskeleton at the center of the mechanisms that promote growth cone formation and the subsequent regeneration.

In addition to cytoskeleton, the subcellular organelles also need to reorganize and adapt to injury-induced alterations in injured axons. Being at the crossroads of many of these axonal adaptations to injury, it is possible that mitochondria play an important role in the initiation of axonal regeneration. Some of the most important second messengers such as calcium and reactive oxygen species are mainly derived from mitochondria. Additionally, growth cone formation, microtubule dynamics, and phosphorylation by kinases require a large amount of ATP. Recent studies in different models have begun to reveal the role of mitochondria in axonal regeneration (Cartoni et al., 2016; Han, Baig, & Hammarlund, 2016; Zhou et al., 2016). Of note, in RGCs, increasing mitochondrial transport by over-expression of the mammalian-specific Armadillo repeat-containing X-linked protein 1 (Armcx1) leads to a high level of short-distance regeneration but a weaker effect for long-distance extension (Cartoni et al., 2016), suggesting a preferential involvement in the initiation phase of regeneration. However, how mitochondria regulate other functions requires further investigation.

Sustaining Axonal Re-Regrowth

Similar to the initiation, sustained growth requires the localized actions in the lesioned axons. However, continuous axon extension faces additional challenges to synthesize more building blocks and transport them to the correct locations. Toward the completion of neural development, mature neurons need to switch from axon-growth mode to dendrite/synapse growth mode. Thus, de novo axon growth programs need to be activated to coordinate the actions between the soma and the axonal process. Recent studies have begun to reveal the genetic programs of axon regeneration.

An important realization is that axon regeneration could be activated by inflammation that occurs in the vicinity of injured neurons. For example, a lens injury prior to optic nerve crush resulted in a significant increase in axonal regeneration (Fischer, Pavlidis, & Thanos, 2000; Mansour-Robaey, Clarke, Wang, Bray, & Aguayo, 1994). Subsequent studies of the mechanism revealed that the lens injury induces an inflammatory response that leads to the infiltration and activation of macrophages, Müller cells, and increased expression of the growth-associated protein GAP-43 (Leon, Yin, Nguyen, Irwin, & Benowitz, 2000; Yin et al., 2003). Similarly, inducing inflammation via intravitreal injection of zymosan, a yeast wall component, also promotes axonal regeneration, suggesting direct links between inflammation, innate-immune response, and axonal regeneration (Kurimoto et al., 2010; Yin et al., 2006, 2009). Interestingly, whereas both injection of zymosan and the bacterial cell wall component lipopolysaccharide (LPS) induces similar inflammatory response, axonal regeneration was only observed in the zymosan-treated group and this effect is abolished in mouse after deletion of the receptor dectin-1 (Baldwin, Carbajal, Segal, & Giger, 2015). However, inflammation is a double-edged sword with both reparative and pathological properties (Gensel et al., 2015) and challenges remain to find approaches to fine-tune these processes.

In further support of regenerative growth as a neuronal response to inflammation and stresses, genetic studies revealed that the suppressor of cytokine signaling 3 (SOCS3) (Baker, Akhtar, & Benveniste, 2009), an important regulator of the inflammatory and immune response in CNS, acts as a negative regulator of axonal regeneration (Smith et al., 2009). Induced by cytokine-activated JAK/STAT pathways, SOCS3 acts as a negative regulator of this pathway, preventing the over-activation of inflammatory responses. Releasing this inhibition promotes STAT3 responsive genes, including many pro-regenerative ones, leading to axonal regeneration (Smith et al., 2009).

The archetype of the intrinsic and cell-autonomous regenerative program regulator is the phosphatase and tensin homolog (PTEN), a negative regulator of the mTOR pathway. PTEN is a phosphatase that converts, via inhibition of the phosphoinositide 3-kinases (PI3K), phosphatidylinositol trisphosphate (PIP3) to phosphatidylinositol bisphosphate (PIP2) (Knafo & Esteban, 2017; Worby & Dixon, 2014). The role of the mTOR pathway in cellular growth has been extensively described (Mathieu Laplante, 2012). Correlated with development-dependent decline of axon growth ability, the activation level of this pathway is gradually decreased (Park et al., 2008). Remarkably, increasing the mTOR pathway via PTEN inhibition results in profound increases of neuronal survival and axonal regeneration after optic nerve crush. This effect appears to be conserved in other neuronal types in rodents (Liu et al., 2010) and across different species (Byrne et al., 2014; Hu, 2015). Increasing this pathway via specific deletion of the tuberous sclerosis protein 1 (TSC1) gene also increases axonal regeneration of RGCs, although to a lesser extent, underscoring the central role played by mTOR as well as other related pathways in adult regeneration of CNS neurons (Park et al., 2008). Despite aging-dependent decrease (Geoffroy, Hilton, Tetzlaff, & Zheng, 2016), deleting PTEN up to 12 months post-injury still promotes injured corticospinal axons to regenerate albeit at a reduced speed (Du et al., 2015). It is important to note that mTOR regulates a general growth pathway and uncontrolled over-activation could lead to side effects. Indeed, long-term deletion of PTEN in the sensorimotor cortex leads to an increase in the cortical thickness and a disruption of cortical lamination (Gutilla, Buyukozturk, & Steward, 2016).

Because of limited numbers and distances of axon regeneration observed in individual manipulations, it is conceivable that an individual signaling pathway acts on certain populations of neurons and/or certain aspects of regenerative mechanisms. In this regard, PTEN deletion selectively promotes the regeneration from a specific type of RGCs, alpha-RGCs (Duan et al., 2015). Thus, co-manipulations of these pathways may maximize their regenerative effects. Indeed, a number of manipulations have been shown to exhibit synergistic effects with PTEN deletion/inhibition. For example, co-deleted PTEN and SOCS3 resulted in much-increased extents of axonal regrowth. The synergistic effects were characterized by transcriptome analysis, which showed that co-deleting PTEN and SOCS3 triggered the up-regulation of a specific set of genes whose expression was unchanged in single knock-outs (Sun et al., 2012). Similar additive or synergistic effects have been documented for other manipulations with PTEN deletion (see, e.g., Norsworthy et al., 2017; Luo et al., 2016; Bei et al., 2016; Cartoni et al., 2016; Belin et al., 2015; Nawabi et al., 2015; de Lima et al., 2012; Sun et al., 2012; Kurimoto et al., 2010).

Different axon growth ability in immature and mature neurons has provided mechanistic insights into the mechanisms of axon regeneration. Another transcriptional repressor for axonal growth and regeneration, the krüppel-like factor-4 (KLF4), was discovered when comparing gene expression patterns of RGCs at different developmental stages (Moore et al., 2009). KLF4 knock-out increases axonal outgrowth of RGCs in culture and promotes axonal regeneration after optic nerve crush. The effects of KLF family members appear to be conserved in other species, as these molecules have been shown to regulate axon regeneration in both mammals and zebra fish (Moore et al., 2009; Veldman, Bemben, Thompson, & Goldman, 2007). On the other hand, Sox11, a transcription factor involved in axon growth during development (Chang et al., 2017), recently has been shown to promote axon regeneration after optic nerve injury (Norsworthy et al., 2017). Intriguingly, in contrast to PTEN deletion that promotes axon regeneration from alpha-RGCs (Duan et al., 2015), Sox11 kills most alpha-RGCs and promotes the regeneration from other types (Norsworthy et al., 2017). These results revealed unexpected cell type specificity in regulating axon regeneration, which should be a focus of the next frontier of axon regeneration research.

Reaching the Targets and Beyond

To restore function, regenerated axons need to establish functional connection in the targets. So far, most of our knowledge on this issue comes from the optic nerve crush model in mice. In this model, regenerating axons are initially confined within injured optic nerves for about 2 millimeters. However, at the chiasm, a classical model of choice points for retinal axons during development (Petros, Rebsam, & Mason, 2008; Sitko, Kuwajima, & Mason, 2018; Wang, Marcucci, Cerullo, & Mason, 2016), regenerating axons have to make choices, either stopping growing, continuing growing with or without crossing to the contralateral side of the optic tract, or even toward contralateral optic nerves. Robust axon regeneration observed with different combinatorial treatments offered opportunities of assessing their projection patterns. While some studies suggested precise targeting (de Lima et al., 2012; Lim et al., 2016), others found that many axons make mistakes at the chiasm and over-shoot or completely miss their original target after crossing the chiasm (Belin et al., 2015; Luo et al., 2013). Such projection errors are not restricted to regenerating axons from RGCs, as similar mistargeting has been observed in regenerating axons in peripheral nerves (Kelamangalath et al., 2015) and in C. elegans (Gabel, Antoine, Chuang, Samuel, & Chang, 2008; Wu et al., 2007). Thus, guiding regenerating axons to appropriate targets remains an important challenge.

On the other hand, once reaching the targets, regenerating axons are able to make functional synapses, but surprisingly, fail to be myelinated (Bei et al., 2016). As a result, even when regenerated axons had made functional synapses in the target, the mice failed to recover their visual acuity unless they were treated with voltage-gated potassium channel blockers that restore conduction (Bei et al., 2016). These results revealed another barrier for functional restoration: the re-myelination of regenerating axons. Thus, further studies are needed to understand the mechanisms by which regenerated axons fail to be myelinated, which could provide insights into ways to overcome this and other possible hurdles for functional recovery.


Extensive studies in the past decade led to unprecedented progress in our understanding of the cellular and molecular mechanisms of axon regeneration in different models. Some of these findings showed clear translational potential. For example, clinically used microtubule-stabilizing compounds are promising to promote axon regeneration and functional recovery. Similarly, based on mTOR effects on axon regeneration, more translational approaches could be devised. For example, osteopontin was discovered to possess the ability to sensitize neuronal responses to IGF1 and BDNF, resulting in axon regrowth and functional recovery (Duan et al., 2015). However, even with the best combinations, only subsets of injured axons are able to regenerate for relatively short distances. A possible reason is that different types of neurons have distinct abilities to respond to individual treatments. Thus, understanding the molecular basis for these different types of neurons should be extremely informative.


Abe, N., & Cavalli, V. (2008). Nerve injury signaling. Current Opinion in Neurobiology, 18, 276–283.Find this resource:

Ambron, R. T., & Walters, E. T. (1996). Priming events and retrograde injury signals: A new perspective on the cellular and molecular biology of nerve regeneration. Molecular Neurobiology, 13, 61–79.Find this resource:

Baker, B. J., Akhtar, L. N., & Benveniste, E. N. (2009). SOCS1 and SOCS3 in the control of CNS immunity. Trends in Immunology, 30, 392–400.Find this resource:

Baldwin, K. T., Carbajal, K. S., Segal, B. M., & Giger, R. J. (2015). Neuroinflammation triggered by β‎-glucan/dectin-1 signaling enables CNS axon regeneration. Proceedings of the National Academy of Sciences, 112, 2581–2586.Find this resource:

Banner, L. R., & Patterson, P. H. (1994). Major changes in the expression of the mRNAs for cholinergic differentiation factor/leukemia inhibitory factor and its receptor after injury to adult peripheral nerves and ganglia. Proceedings of the National Academy of Sciences, 91, 7109–7113.Find this resource:

Bei, F., Lee, H. H. C., Liu, X., Gunner, G., Jin, H., Ma, L., . . . Frank, E. (2016). Restoration of visual function by enhancing conduction in regenerated axons. Cell, 164, 219–232.Find this resource:

Belin, S., Nawabi, H., Wang, C., Tang, S., Latremoliere, A., Warren, P., . . . He, Z. (2015). Injury-induced decline of intrinsic regenerative ability revealed by quantitative proteomics. Neuron, 86, 1000–1014.Find this resource:

Bradke, F., Fawcett, J. W., & Spira, M. E. (2012). Assembly of a new growth cone after axotomy: The precursor to axon regeneration. Nature Reviews Neuroscience, 13, 183–193.Find this resource:

Byrne, A. B., & Hammarlund, M. (2017). Axon regeneration in C. elegans: Worming our way to mechanisms of axon regeneration. Experimental Neurology, 287, 300–309.Find this resource:

Byrne, A. B., Walradt, T., Gardner, K. E., Hubbert, A., Reinke, V., & Hammarlund, M. (2014). Insulin/IGF1 signaling inhibits age-dependent axon regeneration. Neuron, 81, 1–13.Find this resource:

Cartoni, R., Norsworthy, M. W., Bei, F., Wang, C., Li, S., Zhang, Y., . . . He, Z. (2016). The mammalian-specific protein armcx1 regulates mitochondrial transport during axon regeneration. Neuron, 92, 1294–1307.Find this resource:

Cavalli, V. (2005). Sunday Driver links axonal transport to damage signaling. Journal of Cell Biology, 168, 775–787.Find this resource:

Chang, K.-C., Hertz, J., Zhang, X., Jin, X.-L., Shaw, P., Derosa, B. A., . . . Patel, R. D. (2017). Novel regulatory mechanisms for the SoxC transcriptional network required for visual pathway development. Journal of Neuroscience, 37, 4967–4981.Find this resource:

Chen, L., Chuang, M., Koorman, T., Boxem, M., Jin, Y., & Chisholm, A. D. (2015). Axon injury triggers EFA-6 mediated destabilization of axonal microtubules via TACC and doublecortin like kinase. eLife Sciences, 4, 2177.Find this resource:

Chen, L., Wang, Z., Ghosh-Roy, A., Hubert, T., Yan, D., O’Rourke, S., . . . Chisholm, A. D. (2011). Axon regeneration pathways identifiedby systematic genetic screening in C. elegans. Neuron, 71, 1043–1057.Find this resource:

Chierzi, S., Ratto, G. M., Verma, P., & Fawcett, J. W. (2005). The ability of axons to regenerate their growth cones depends on axonal type and age, and is regulated by calcium, cAMP and ERK. European Journal of Neuroscience, 21, 2051–2062.Find this resource:

Curtis, R., Adryan, K. M., Zhu, Y., Harkness, P. J., Lindsay, R. M., & DiStefano, P. S. (1993). Retrograde axonal transport of ciliary neurotrophic factor is increased by peripheral nerve injury. Nature, 365, 253–255.Find this resource:

Curtis, R., Scherer, S. S., Somogyi, R., Adryan, K. M., Ip, N. Y., Zhu, Y., . . . DiStefano, P. S. (1994). Retrograde axonal transport of LIF is increased by peripheral nerve injury: Correlation with increased LIF expression in distal nerve. Neuron, 12, 191–204.Find this resource:

de Lima, S., Koriyama, Y., Kurimoto, T., Oliveira, J. T., Yin, Y., Li, Y., . . . Benowitz, L. (2012). Full-length axon regeneration in the adult mouse optic nerve and partial recovery of simple visual behaviors. Proceedings of the National Academy of Sciences, 109, 9149–9154.Find this resource:

Du, K., Zheng, S., Zhang, Q., Li, S., Gao, X., Wang, J., . . . Liu, K. (2015). PTEN deletion promotes regrowth of corticospinal tract axons 1 year after spinal cord injury. Journal of Neuroscience, 35, 9754–9763.Find this resource:

Duan, X., Qiao, M., Bei, F., Kim, I.-J., He, Z., & Sanes, J. R. (2015). Subtype-specific regeneration of retinal ganglion cells following axotomy: Effects of Osteopontin and mTOR signaling. Neuron, 85, 1244–1256.Find this resource:

Erez, H., & Spira, M. E. (2008). Local self-assembly mechanisms underlie the differential transformation of the proximal and distal cut axonal ends into functional and aberrant growth cones. Journal of Comparative Neurology, 507, 1019–1030.Find this resource:

Ertürk, A., Hellal, F., Enes, J., & Bradke, F. (2007). Disorganized microtubules underlie the formation of retraction bulbs and the failure of axonal regeneration. Journal of Neuroscience, 27, 9169–9180.Find this resource:

Fischer, D., Pavlidis, M., & Thanos, S. (2000). Cataractogenic lens injury prevents traumatic ganglion cell death and promotes axonal regeneration both in vivo and in culture. Investigative Ophthalmology & Visual Science, 41, 3943–3954.Find this resource:

Fregnan, F., Muratori, L., Simões, A. R., Giacobini-Robecchi, M. G., & Raimondo, S. (2012). Role of inflammatory cytokines in peripheral nerve injury. Neural Regeneration Research, 7, 2259–2266.Find this resource:

Gabel, C. V., Antoine, F., Chuang, C.-F., Samuel, A. D. T., & Chang, C. (2008). Distinct cellular and molecular mechanisms mediate initial axon development and adult-stage axon regeneration in C. elegans. Development, 135, 1129–1136.Find this resource:

Gensel, J. C., & Zhang, B. (2015). Macrophage activation and its role in repair and pathology after spinal cord injury. Brain Research, 1619, 1–11.Find this resource:

Geoffroy, C. G., & Zheng, B. (2014). Myelin-associated inhibitors in axonal growth after CNS injury. Current Opinion in Neurobiology, 27, 31–38.Find this resource:

Geoffroy, C. G., Hilton, B. J., Tetzlaff, W., & Zheng, B. (2016). Evidence for an age-dependent decline in axon regeneration in the adult mammalian central nervous system. Cell Reports, 15, 238–246.Find this resource:

Ghosh-Roy, A., Wu, Z., Goncharov, A., Jin, Y., & Chisholm, A. D. (2010). Calcium and cyclic AMP promote axonal regeneration in Caenorhabditis elegans and require DLK-1 kinase. Journal of Neuroscience, 30, 3175–3183.Find this resource:

Gitler, D., & Spira, M. E. (1998). Real time imaging of calcium-induced localized proteolytic activity after axotomy and its relation to growth cone formation. Neuron, 20, 1123–1135.Find this resource:

Gutilla, E. A., Buyukozturk, M. M., & Steward, O. (2016). Long-term consequences of conditional genetic deletion of PTEN in the sensorimotor cortex of neonatal mice. Experimental Neurology, 279, 27–39.Find this resource:

Hammarlund, M., & Jin, Y. (2014). Axon regeneration in C. elegans. Current Opinion in Neurobiology, 27, 199–207.Find this resource:

Hammarlund, M., Nix, P., Hauth, L., Jorgensen, E. M., & Bastiani, M. (2009). Axon regeneration requires a conserved MAP kinase pathway. Science, 323, 802–806.Find this resource:

Han, S. M., Baig, H. S., & Hammarlund, M. (2016). Mitochondria localize to injured axons to support regeneration. Neuron, 92, 1308–1323.Find this resource:

Hanz, S., Perlson, E., Willis, D., Zheng, J.-Q., Massarwa, R., Huerta, J. J., . . . Twiss, J. L. (2003). Axoplasmic importins enable retrograde injury signaling in lesioned nerve. Neuron, 40, 1095–1104.Find this resource:

He, Z., & Jin, Y. (2016). Intrinsic control of axon regeneration. Neuron, 90, 437–451.Find this resource:

He, Z., & Koprivica, V. (2004). The nogo signaling pathway for regeneration block. Annual Review of Neuroscience, 27, 341–368.Find this resource:

Hellal, F., Hurtado, A., Ruschel, J., Flynn, K. C., Laskowski, C. J., Umlauf, M., . . . Bixby, J. (2011). Microtubule stabilization reduces scarring and causes axon regeneration after spinal cord injury. Science, 331, 928–931.Find this resource:

Hilton, B. J., & Bradke, F. (2017). Can injured adult CNS axons regenerate by recapitulating development? Development, 144, 3417–3429.Find this resource:

Hu, Y. (2015). The necessary role of mTORC1 in central nervous system axon regeneration. Neural Regeneration Research, 10, 186.Find this resource:

Kelamangalath, L., Tang, X., Bezik, K., Sterling, N., Son, Y.-J., & Smith, G. M. (2015). Neurotrophin selectivity in organizing topographic regeneration of nociceptive afferents. Experimental Neurology, 271, 262–278.Find this resource:

Knafo, S., & Esteban, J. A. (2017). PTEN: Local and global modulation of neuronal function in health and disease. Trends in Neurosciences, 40, 83–91.Find this resource:

Knöferle, J., Koch, J. C., Ostendorf, T., Michel, U., Planchamp, V., Vutova, P., . . . Bähr, M. (2010). Mechanisms of acute axonal degeneration in the optic nerve in vivo. Proceedings of the National Academy of Sciences, 107, 6064–6069.Find this resource:

Kurimoto, T., Yin, Y., Omura, K., Gilbert, H. Y., Kim, D., Cen, L. P., . . . Benowitz, L. I. (2010). Long-distance axon regeneration in the mature optic nerve: Contributions of Oncomodulin, cAMP, and PTEN gene deletion. Journal of Neuroscience, 30, 15654–15663.Find this resource:

Leon, S., Yin, Y., Nguyen, J., Irwin, N., & Benowitz, L. I. (2000). Lens injury stimulates axon regeneration in the mature rat optic nerve. Journal of Neuroscience, 20, 4615–4626.Find this resource:

Lieberman, A. R. (1971). The axon reaction: A review of the principal features of perikaryal responses to axon injury. International Review of Neurobiology, 14, 49–124.Find this resource:

Lim, J.-H. A., Stafford, B. K., Nguyen, P. L., Lien, B. V., Wang, C., Zukor, K., . . . Huberman, A. D. (2016). Neural activity promotes long-distance, target-specific regeneration of adult retinal axons. Nature Neuroscience, 19, 1073–1084.Find this resource:

Lindwall, C., & Kanje, M. (2005). Retrograde axonal transport of JNK signaling molecules influence injury induced nuclear changes in p-c-Jun and ATF3 in adult rat sensory neurons. Molecular and Cellular Neuroscience, 29, 269–282.Find this resource:

Liu, K., Lu, Y., Lee, J. K., Samara, R., Willenberg, R., Sears-Kraxberger, I., . . . Bin Cai (2010). PTEN deletion enhances the regenerative ability of adult corticospinal neurons. Nature Neuroscience, 13, 1075–1081.Find this resource:

Luo, X., Ribeiro, M., Bray, E. R., Lee, D.-H., Yungher, B. J., Mehta, S. T., . . . Moraes, C. T. (2016). Enhanced transcriptional activity and mitochondrial localization of STAT3 co-induce axon regrowth in the adult central nervous system. Cell Reports, 15, 1–13.Find this resource:

Luo, X., Salgueiro, Y., Beckerman, S. R., Lemmon, V. P., Tsoulfas, P., & Park, K. K. (2013). Three-dimensional evaluation of retinal ganglion cell axon regeneration and pathfinding in whole mouse tissue after injury. Experimental Neurology, 247, 653–662.Find this resource:

Mansour-Robaey, S., Clarke, D. B., Wang, Y. C., Bray, G. M., & Aguayo, A. J. (1994). Effects of ocular injury and administration of brain-derived neurotrophic factor on survival and regrowth of axotomized retinal ganglion cells. Proceedings of the National Academy of Sciences, 91, 1632–1636.Find this resource:

Mathieu Laplante, D. M. S. (2012). mTOR signaling in growth control and disease. Cell, 149, 274–293.Find this resource:

Moore, D. L., Blackmore, M. G., Hu, Y., Kaestner, K. H., Bixby, J. L., Lemmon, V. P., & Goldberg, J. L. (2009). KLF family members regulate intrinsic axon regeneration ability. Science, 326, 298–301.Find this resource:

Nawabi, H., Belin, S., Cartoni, R., Williams, P. R., Wang, C., Latremoliere, A., . . . Fu, X. (2015). Doublecortin-like kinases promote neuronal survival and induce growth cone reformation via distinct mechanisms. Neuron, 88, 704–719.Find this resource:

Norsworthy, M. W., Bei, F., Kawaguchi, R., Wang, Q., Tran, N. M., Li, Y., . . . Sanes, J. R. (2017). Sox11 expression promotes regeneration of some retinal ganglion cell types but kills others. Neuron, 94, 1112–1120.e1114.Find this resource:

O’Brien, J. J., & Nathanson, N. M. (2007). Retrograde activation of STAT3 by leukemia inhibitory factor in sympathetic neurons. Journal of Neurochemistry, 103, 288–302.Find this resource:

Park, K. K., Hu, Y., Muhling, J., Pollett, M. A., Dallimore, E. J., Turnley, A. M., . . . Harvey, A. R. (2009). Cytokine-induced SOCS expression is inhibited by cAMP analogue: Impact on regeneration in injured retina. Molecular and Cellular Neuroscience, 41, 313–324.Find this resource:

Park, K. K., Liu, K., Hu, Y., Smith, P. D., Wang, C., Cai, B., . . . Sahin, M. (2008). Promoting axon regeneration in the adult CNS by modulation of the PTEN/mTOR pathway. Science, 322, 963–966.Find this resource:

Perlson, E., Hanz, S., Ben-Yaakov, K., Segal-Ruder, Y., Seger, R., & Fainzilber, M. (2005). Vimentin-dependent spatial translocation of an activated MAP kinase in injured nerve. Neuron, 45, 715–726.Find this resource:

Petros, T. J., Rebsam, A., & Mason, C. A. (2008). Retinal axon growth at the optic chiasm: To cross or not to cross. Annual Review of Neuroscience, 31, 295–315.Find this resource:

Pivovarova, N. B., & Andrews, S. B. (2010). Calcium-dependent mitochondrial function and dysfunction in neurons. FEBS Journal, 277, 3622–3636.Find this resource:

Rasmussen, J. P., & Sagasti, A. (2017). Learning to swim, again: Axon regeneration in fish. Experimental Neurology, 287, 318–330.Find this resource:

Ribas, V. T., Koch, J. C., Michel, U., Bähr, M., & Lingor, P. (2016). Attenuation of axonal degeneration by calcium channel inhibitors improves retinal ganglion cell survival and regeneration after optic nerve crush. Molecular Neurobiology, 54, 72–86.Find this resource:

Rishal, I., & Fainzilber, M. (2013). Axon–soma communication in neuronal injury. Nature Reviews Neuroscience, 15, 32–42.Find this resource:

Ruschel, J., & Bradke, F. (2018). Systemic administration of epothilone D improves functional recovery of walking after rat spinal cord contusion injury. Experimental Neurology, 306, 243–249.Find this resource:

Ruschel, J., Hellal, F., Flynn, K. C., Dupraz, S., Elliott, D. A., Tedeschi, A., . . . Dobrint, K. (2015). Systemic administration of epothilone B promotes axon regeneration after spinal cord injury. Science, 348(6232), 347–352.Find this resource:

Sandner, B., Puttagunta, R., Motsch, M., Bradke, F., Ruschel, J., Blesch, A., & Weidner, N. (2018). Systemic epothilone D improves hindlimb function after spinal cord contusion injury in rats. Experimental Neurology, 306, 250–259.Find this resource:

Shin, J. E., Cho, Y., Beirowski, B., Milbrandt, J., Cavalli, V., & DiAntonio, A. (2012). Dual Leucine zipper kinase is required for retrograde injury signaling and axonal regeneration. Neuron, 74, 1015–1022.Find this resource:

Silver, J., Schwab, M. E., & Popovich, P. G. (2015). Central nervous system regenerative failure: Role of oligodendrocytes, astrocytes, and microglia. Cold Spring Harbor Perspectives in Biology, 7, a020602.Find this resource:

Sitko, A. A., Kuwajima, T., & Mason, C. A. (2018). Eye-specific segregation and differential fasciculation of developing retinal ganglion cell axons in the mouse visual pathway. Journal of Comparative Neurology, 130, 4999.Find this resource:

Smith, P., Sun, F., Park, K. K., Cai, B., Wang, C., Kuwako, K., . . . He, Z. (2009). SOCS3 deletion promotes optic nerve regeneration in vivo. Neuron, 64, 617–623.Find this resource:

Sun, F., & He, Z. (2010). Neuronal intrinsic barriers for axon regeneration in the adult CNS. Current Opinion in Neurobiology, 20, 510–518.Find this resource:

Sun, F., Park, K. K., Belin, S., Wang, D., Lu, T., Chen, G., . . . Yankner, B. A. (2012). Sustained axon regeneration induced by co-deletion of PTEN and SOCS3. Nature, 480, 372–375.Find this resource:

Sun, L., Shay, J., McLoed, M., Roodhouse, K., Chung, S. H., Clark, C. M., . . . Gabel, C. V. (2014). Neuronal regeneration in C. elegans requires subcellular calcium release by ryanodine receptor channels and can be enhanced by optogenetic stimulation. Journal of Neuroscience, 34, 15947–15956.Find this resource:

Sun, Y., & Zigmond, R. E. (1996). Leukaemia inhibitory factor induced in the sciatic nerve after axotomy is involved in the induction of galanin in sensory neurons. European Journal of Neuroscience, 8, 2213–2220.Find this resource:

Sung, Y. J., Chiu, D. T. W., & Ambron, R. T. (2006). Activation and retrograde transport of protein kinase G in rat nociceptive neurons after nerve injury and inflammation. Neuroscience, 141, 697–709.Find this resource:

Tedeschi, A., Dupraz, S., Laskowski, C. J., Xue, J., Ulas, T., Beyer, M., . . . Bradke, F. (2016). The calcium channel subunit Alpha2delta2 suppresses axon regeneration in the adult CNS. Neuron, 92, 1–16.Find this resource:

Valakh, V., Walker, L. J., Skeath, J. B., & DiAntonio, A. (2013). Loss of the spectraplakin short stop activates the DLK injury response pathway in Drosophila. Journal of Neuroscience, 33, 17863–17873.Find this resource:

Veldman, M. B., Bemben, M. A., Thompson, R. C., & Goldman, D. (2007). Gene expression analysis of zebrafish retinal ganglion cells during optic nerve regeneration identifies KLF6a and KLF7a as important regulators of axon regeneration. Developmental Biology, 312, 596–612.Find this resource:

Wang, Q., Marcucci, F., Cerullo, I., & Mason, C. (2016). Ipsilateral and contralateral retinal ganglion cells express distinct genes during decussation at the optic chiasm. Eneuro, 3, ENEURO.0169–16.2016.Find this resource:

Watkins, T. A., Wang, B., Huntwork-Rodriguez, S., Yang, J., Jiang, Z., Eastham-Anderson, J., . . . Lewcock, J. W. (2013). DLK initiates a transcriptional program that couples apoptotic and regenerative responses to axonal injury. Proceedings of the National Academy of Sciences, 110, 4039–4044.Find this resource:

Williams, P. R., Marincu, B.-N., Sorbara, C. D., Mahler, C. F., Schumacher, A.-M., Griesbeck, O., . . . Misgeld, T. (2014). A recoverable state of axon injury persists for hours after spinal cord contusion in vivo. Nature Communications, 5, 5683.Find this resource:

Worby, C. A., & Dixon, J. E. (2014). PTEN. Annual Review of Biochemistry, 83, 641–669.Find this resource:

Wu, Z., Ghosh-Roy, A., Yanik, M. F., Zhang, J. Z., Jin, Y., & Chisholm, A. D. (2007). Caenorhabditis elegans neuronal regeneration is influenced by life stage, ephrin signaling, and synaptic branching. Proceedings of the National Academy of Sciences, 104, 15132–15137.Find this resource:

Yan, D., Wu, Z., Chisholm, A. D., & Jin, Y. (2009). The DLK-1 kinase promotes mRNA stability and local translation in C. elegans synapses and axon regeneration. Cell, 138, 1005–1018.Find this resource:

Yang, P., Wen, H., Ou, S., Cui, J., & Fan, D. (2012). IL-6 promotes regeneration and functional recovery after cortical spinal tract injury by reactivating intrinsic growth program of neurons and enhancing synapse formation. Experimental Neurology, 236, 19–27.Find this resource:

Yanik, M. F., Cinar, H., Cinar, H. N., Chisholm, A. D., Jin, Y., & Ben-Yakar, A. (2004). Neurosurgery: Functional regeneration after laser axotomy. Nature, 432, 822–822.Find this resource:

Yin, Y., Cui, Q., Gilbert, H.-Y., Yang, Y., Yang, Z., Berlinicke, C., . . . Petkova, V. (2009). Oncomodulin links inflammation to optic nerve regeneration. Proceedings of the National Academy of Sciences, 106, 19587–19592.Find this resource:

Yin, Y., Cui, Q., Li, Y., Irwin, N., Fischer, D., Harvey, A. R., & Benowitz, L. I. (2003). Macrophage-derived factors stimulate optic nerve regeneration. Journal of Neuroscience, 23, 2284–2293.Find this resource:

Yin, Y., Henzl, M. T., Lorber, B., Nakazawa, T., Thomas, T. T., Jiang, F., . . . Benowitz, L. I. (2006). Oncomodulin is a macrophage-derived signal for axon regeneration in retinal ganglion cells. Nature Neuroscience, 9, 843–852.Find this resource:

Yiu, G. (2003). Signaling mechanisms of the myelin inhibitors of axon regeneration. Current Opinion in Neurobiology, 13, 545–551.Find this resource:

Yiu, G., & He, Z. (2006). Glial inhibition of CNS axon regeneration. Nature Review Neuroscience, 7, 617–627.Find this resource:

Yudin, D., Hanz, S., Yoo, S., Iavnilovitch, E., Willis, D., Gradus, T., . . . Hieda, M. (2008). Localized regulation of axonal RanGTPase controls retrograde injury signaling in peripheral nerve. Neuron, 59, 241–252.Find this resource:

Zhou, B., Yu, P., Lin, M.-Y., Sun, T., Chen, Y., & Sheng, Z.-H. (2016). Facilitation of axon regeneration by enhancing mitochondrial transport and rescuing energy deficits. Journal of Cell Biology, 104, jcb.201605101.Find this resource: