Intra-axonal mechanisms driving axon regeneration (original) (raw)

. Author manuscript; available in PMC: 2021 Aug 1.

Abstract

Traumatic injury to the peripheral and central nervous systems very often causes axotomy, where an axon loses connections with its target resulting in loss of function. The axon segments distal to the injury site lose connection with the cell body and degenerate. Axotomized neurons in the periphery can spontaneously mount a regenerative response and reconnect to their denervated target tissues, though this is rarely complete in humans. In contrast, spontaneous regeneration rarely occurs after axotomy in the spinal cord and brain. Here, we concentrate on the mechanisms underlying this spontaneous regeneration in the peripheral nervous system, focusing on events initiated from the axon that support regenerative growth. We contrast this with what is known for axonal injury responses in the central nervous system. Considering the neuropathy focus of this special issue, we further draw parallels and distinctions between the injury-response mechanisms that initiate regenerative gene expression programs and those that are known to trigger axon degeneration.

INTRODUCTION

It is well established that axons in the peripheral nervous system (PNS) regenerate more effectively than those of the central nervous system (CNS). The poor regeneration in the CNS is due to both extrinsic and intrinsic factors, and much effort has been devoted to understanding how PNS neurons regenerate with the goal of targeting those mechanisms to increase regeneration after brain and spinal cord injuries. Injury to PNS axons initiates a robust intrinsic growth program (Huebner and Strittmatter, 2009). The differences in intrinsic growth capacity between PNS and CNS axons become quite clear when examining dorsal root ganglion (DRG) neurons, which have both peripherally and centrally projecting axons. Injuring the peripherally projecting axon of these neurons activates a regeneration program but injuring the centrally projecting DRG axons does not activate this regeneration program (McQuarrie and Grafstein, 1973; Neumann and Woolf, 1999). However, if the peripheral axon is injured first (termed a ‘conditioning lesion’), the central axon is able to regenerate after a subsequent injury (Chong et al., 1999; Neumann and Woolf, 1999; Richardson and Issa, 1984). This conditioning effect is also seen in the PNS when a nerve is injured proximal to the initial conditioning lesion, indicating that the growth potential can be broadly changed by peripheral nerve injury (Forman et al., 1980). Using a similar in vivo conditioning PNS axotomy followed by sensory neuron culture, Smith and Skene showed that a transcriptional switch underlies this conditioning-effect by activating a growth-promoting gene expression program (Smith and Skene, 1997). Consistent with this, injection of non-hydrolysable, cell-permeable cyclic AMP into the DRG, allows injured DRG neurons to regenerate axons after severing their central axons (Neumann et al., 2002; Qiu et al., 2002). These early observations compiled with many other lines of evidence argue that initiation of a neuronal growth program involves a sequence of events that transforms the neuron from a normal maintenance/function mode to a re-growth mode. Axonal injury is the first event in this sequence, and PNS axotomy precipitates axon-to-soma signals that support the transition from maintenance/function to growth mode. Localized events within the axon contribute to transformation to this growth mode, and soma-to-axon signaling further support axon regeneration after injury. We will address these axon-initiated mechanisms in the paragraphs below, focusing on how injury is signaled to the soma (axon-to-soma signaling) and how the soma alters axon growth (soma-to-axon signaling) with a particular focus on PNS neurons.

I. Axoplasmic increase in Calcium is an early response to initiate regenerative growth programs.

Calcium ion (Ca2+) influx into axons is one of the earliest changes following axotomy. This increase in axoplasmic Ca2+ is needed to reseal the injured axonal membrane, locally activate the Ca2+-dependent protease calpain, and initiate retrograde signaling mechanisms to communicate with the cell body (Figure 1A) (Bradke et al., 2012; Detrait et al., 2000; Rishal and Fainzilber, 2010). In cultured adult DRG neurons, removing Ca2+ from the media reduces neurite growth showing that Ca2+ influx has an important role in regulating axonal growth (Chierzi et al., 2005). In vivo, Ca2+ chelation significantly reduces axon regeneration in C. elegans (Ghosh-Roy et al., 2010). Despite the importance of Ca2+ entry, the regenerative effects of Ca2+ on PNS axons is not seen in the centrally projecting DRG axons (Hervera et al., 2019), so there may be discrete differences between PNS and CNS axons in this regard. Ca2+ increase occurs through openings in the membrane caused by the physical injury to the axon and by reversal of the sodium/calcium exchanger and voltage gated ion channels (Kamber et al., 2009; Kulbatski et al., 2004; Ziv and Spira, 1997). Ca2+ entry through voltage-gated calcium channels after injury is caused by membrane depolarization due to activation of gamma aminobutyric acid A (GABAA) and N-methyl-D-aspartate (NMDA) receptors (Toyoda et al., 2003), and the voltage-gated ion channel function is needed for restoring axon membrane integrity shortly after injury (Nehrt et al., 2007). In the cockroach giant axon, it was found that Ca2+, through the activation of phospholipase A2, was required for membrane re-sealing after transection (Yawo and Kuno, 1985). Ca2+-mediated activation of cysteine proteases, like calpain and calmodulin, also plays an important role in resealing the axon membrane (Xie and Barrett, 1991).

Figure 1 – Schematic of axon intrinsic signaling mechanisms that contribute to axon regeneration.

A, Depiction of back-propagating Ca2+ wave and localized effects of injury-induced Ca2+ influx into the axon are shown on top. Ca2+ entry triggers a retrograde Ca2+-mediated/dependent axon-to-soma signal (1) and activates localized protein synthesis (2) that provides a second means for axon-to-soma signaling through protein transport (3).

B, Ensuing regeneration-promoting mechanisms are depicted on the bottom, with soma-to-axon signaling through anterograde transport of mRNAs and proteins (4) with subsequent translational regulation of these axonal mRNAs. In addition to Ca2+ influx and release from intra-cellular stores, the extracellular environment, including non-neuronal cells resident and infiltrating into the nerve after injury (5), have the potential modulate localized translation events and mRNA transport.

Increase in axoplasmic Ca2+ is also a critical mediator of Wallerian degeneration in the severed distal axon stump (Conforti et al., 2014). The first phase of Wallerian degeneration begins with a short term Ca2+ wave which leads to a cascade of events including the depletion of nicotinamide adenine dinucleotide (NAD+) and adenosine triphosphate (ATP) as well as disruption of mitochondrial function and release of reactive oxygen species (ROS) (For review see Rosell and Neukomm, 2019). A second feed-forward release of Ca2+ stores from mitochondria and endoplasmic reticulum (ER) has been posited to instigate axon fragmentation during Wallerian degeneration (Barrientos et al., 2011; Villegas et al., 2014). This implies that the proximal axon must be able to buffer the Ca2+ increase relative to the distal axon stump. Work in cultured Aplysia neurons has shown that the proximal tip of severed axons have a Ca2+ concentration gradient with higher levels typical of extracellular spaces near the injury and lower levels more typical of the intracellular environment moving a few microns back towards the soma. This leads to a gradient of protease activity that is required for growth cone formation, and a new growth cone forms approximately mid-way along this Ca2+ gradient (Khoutorsky and Spira, 2005; Spira et al., 2003; Ziv and Spira, 1997). Interestingly, depletion of ER Ca2+ is well known to activate the unfolded protein response (UPR), since ER chaperone proteins needed for folding nascently synthesized proteins are Ca2+-dependent (Harding et al., 2002). Several lines of evidence indicate that localized activation of UPR in axons is needed for initiating regenerative responses (Onate et al., 2016; Ying et al., 2015). Attenuating Ca2+ release by blocking ER inositol triphosphate receptors (IP3R) and ryanodine receptors (RyR) reduces axonal outgrowth after peripheral nerve injury suggesting that regeneration of PNS axons is, at least in part, dependent on the release of ER Ca2+ stores (Ohtake et al., 2018). A decrease in mRNA translation is a critical component of the UPR, as UPR induces an inhibitory phosphorylation of the translation factor eIF2α that has been shown to occur locally in axons (Vuppalanchi et al., 2012). Similar to non-neuronal cells, translation of some mRNAs is paradoxically upregulated despite (or because of) eIF2α phosphorylation. Intra-axonal translation of the mRNA encoding the transcription factor Luman/CREB3 under UPR conditions provides an axon-to-soma signal for altering neuronal gene expression (Ying et al., 2014; Ying et al., 2015). Intra-axonal translation of Calreticulin (Calr) mRNA through UPR-associated eIF2α phosphorylation locally supports growth cone extension (Pacheco et al., 2020). As we outline below, increase in axoplasmic Ca2+ following injury is also needed for translation of axonal mRNAs whose protein products form a retrograde signaling complex to link signaling components to retrograde transport motors (Figure 1A).

Increased intracellular Ca2+ contributes to the generation of cyclic adenosine monophosphate (cAMP) and activation of protein kinase A (PKA), protein kinase C (PKC), and dual leucine zipper kinase 1 (DLK-1) that all positively contribute to axon regeneration (Mar et al., 2014). Ca2+ release from internal stores is also needed for a back-propagating Ca2+ wave that provides very rapid axon-to-soma signaling to activate PKCμ in the soma resulting in phosphorylation and nuclear exit of histone deacetylase (HDAC) 5 (Figure 1A) (Cho et al., 2013). HDAC5 subsequently localizes to axons where it can deacetylate α-tubulin, suggesting that axon-to-soma Ca2+ signaling also supports the dynamic cytoskeleton locally needed for axon regeneration (Cho and Cavalli, 2012; Cho et al., 2013). Ca2+ signaling from axons can also modulate phosphatase activity needed for dephosphorylating HDAC3 (Hervera et al., 2019). These changes in HDAC3 and HDAC5 phosphorylation status (and localization for HDAC5) provide a Ca2+-dependent axon-to-soma regulation of gene expression through modifying the acetylation status of histones (Cho et al., 2013; Hervera et al., 2019).

Despite the clearly positive effects of Ca2+ entry and intracellular store release on the early stages of axon regeneration, it should be noted that increases in axoplasmic Ca2+ can be detrimental for axon regeneration. Excessive levels of intra-axonal Ca2 can trigger axon degeneration in intact axons reminiscent of Wallerian degeneration (Stirling and Stys, 2010). Continued expression of the α2δ2 subunit of voltage-gated Ca2+ channel (Cacna2d2) in axotomized PNS neurons negatively affects their regeneration, and blocking the channel function with pregabalin facilitates axon growth and regeneration in both PNS and CNS axons (Tedeschi et al., 2016). ER Ca2+ release subsequent to IP3R activation triggers degeneration of the distal severed axon segments (Barrientos et al., 2011; Villegas et al., 2014), and work from the Segal lab points to IP3R-mediated ER Ca2+ release as a mechanism of axon degeneration in chemotherapy-induced peripheral neuropathies (Figure 2) (Pease-Raissi et al., 2017). It has also been shown that loss of the sterile alpha and TIR motif-containing 1 (SARM1) protein, which plays an important role in axon degeneration associated with Wallerian degeneration, prevents a second wave of Ca2+ elevation thereby attenuating axon degeneration after injury (Tian et al., 2020). Taken together, these observations suggest that elevations of axonal Ca2+ must be critically balanced and timed to initiate regenerative responses after axonal injury, as too much or too long of a Ca2+ elevation could precipitate axon degeneration.

Figure 2 – Schematic of axon intrinsic signaling mechanisms that contribute to axon degeneration.

Schematic representing potential overlapping mechanisms for preventing Ca2+ store release from ER in mature functioning axons (top) and the loss of this mechanism with paclitaxel-induced peripheral neuropathy (Pease-Raissi et al., 2017). Axonally synthesized Bclw prevents or attenuates activation of IP3R along the ER (A, 1–2). Activation of IP3R (B, 3) was shown to be involved in degeneration Wallerian in severed axons (Villegas et al., 2014) and depletion of ER Ca2+ stores can trigger the UPR (Malhotra and Kaufman, 2007). As outlined in the text, UPR activation in axons is linked to some of the injury-associated translation events seen after the traumatic axonal injury depicted in Figure 1.

II. Axotomy alters axon-to-soma signals to initiate a neuronal regeneration program.

Retrograde signaling through transport of proteins provides another axon-to-soma signaling mechanism to regulate gene expression (Figure 1A). Importin-β1 mRNA (Kpnb1) is locally translated in axons after injury. The newly synthesized Importin-β1 forms an obligate heterodimer with Importin-α family proteins, with the heterodimer linking signaling cargoes containing nuclear localization signals (NLS) to the retrograde transport motor protein dynein (Hanz et al., 2003). This was a surprising finding as these importin proteins (also known as Karyopherins) were thought to only transport cargo proteins across the nuclear pore. Nonetheless, the mRNA encoding the Ran binding protein 1 (RanBP1) is also locally translated after axotomy, and RanBP1 protein facilitates Importin-β/α heterodimer formation by releasing Importin-α from other interacting proteins, a mechanism that was also thought to be restricted to the nuclear pore region (Yudin et al., 2008). Intra-axonal translation of Kpnb1 and RanBP1 mRNAs requires the injury-induced axoplasmic Ca2+ increase outlined above, thereby providing a sequence of elevated Ca2+ → axonal mRNA translation → axon-to-soma signaling (Hanz et al., 2003; Yudin et al., 2008).

Transcription factors are one of the likely cargos for retrograde transport motor proteins and complexes after axonal injury. Though it is not known if Importin-β/α system is involved in retrograde transport for all, some of these retrogradely transported transcription factors are clearly synthesized in axons after injury. Coupled with nuclear transport, these provide an on-demand axon-to-nucleus signaling mechanism for changing transcriptional programs after PNS axotomy. Signal transducer and activator of transcription 3 (STAT3) is one of those locally synthesized transcription factors. Nascently synthesized, retrogradely transported phospho-STAT3 supports survival of DRG neurons following PNS axotomy (Ben-Yaakov et al., 2012). Obviously, survival of the injured neuron is needed for functional regeneration so axonally synthesized STAT3 supports regeneration. Peroxisome proliferator-activated receptor gamma (PPARγ), which has well characterized roles in adipogenesis and glucose homeostasis, is similarly synthesized in PNS axons after axotomy and the retrogradely transported protein is needed for axon regeneration (Lezana et al., 2016). UPR induction after axotomy triggers translation of axonal Luman and Xbp1 mRNAs (Onate et al., 2016; Ying et al., 2014). Luman protein is retrogradely transported to the soma, where its nuclear functions support new gene transcription for axon growth (Ying et al., 2014). Interestingly, Xbp1 mRNA must be spliced before it can encode a functional transcription factor, and Ohate et al. (2016) show that this splicing event occurs in axons after injury, with overexpression of the spliced Xbp1 increasing axon regeneration. However, it should be emphasized that UPR-induced translational activation in axons is not limited to transcription factor mRNAs, as the mRNAs encoding the ER chaperone proteins Calr and Grp78 (also called BiP) are also translated with UPR activation in axons (Vuppalanchi et al., 2012) and recent work has shown that Protein kinase RNA-like endoplasmic reticulum kinase (PERK) is needed for Calr mRNA translation in axons after injury (Pacheco et al., 2020). Intra-axonal synthesis of transcription factor proteins for axonto-nuclear signaling is also not limited to regenerating axons. For example, nerve growth factor (NGF) activates translation of cAMP-responsive element-binding protein (Creb) mRNA in developing sensory axons, and the retrogradely transported CREB protein is needed for NGF-dependent neuronal survival (Cox et al., 2008). However, Creb mRNA was not detected in sympathetic axons (Andreassi et al., 2010), and it does not appear to be a prominent player in adult DRG axons in our hands. Thus, which axon-to-nuclear signals generated is likely unique to different physiological settings and different neuronal types.

Retrogradely transported kinases, which are resident in axons before injury, can also modulate activity of nuclear transcription factors. Axotomy induced activation of extracellular-regulated kinase (ERK) provides one such signal. Here, phospho-ERK binds to importin β1 through a scaffold provided by a proteolytic fragment of Vimentin (Vim) protein, which also prevents ERK dephosphorylation during its transport to the soma where it activates the ETS Transcription Factor ELK1 (Perlson et al., 2005). Vim mRNA is known to be locally translated in axons (Willis et al., 2005), and the proteolytic Vim fragment is generated through a Ca2+-dependent activation of calpain in axons after nerve injury (Perlson et al., 2005). DLK provides an axon-to-soma link to the nuclear transcription factor c-Jun and helps to modulate transport of other injury-induced signals. Sustained phosphorylation of c-Jun after PNS nerve injury is associated with increased regeneration (Kenney and Kocsis, 1998), and DLK knockdown decreases the axotomy-induced c-Jun phosphorylation in DRG after sciatic nerve injury (Holland et al., 2016). Further DLK activation after injury contributes to the retrograde transport of phosphorylated STAT3 (Shin et al., 2012). Notably, Ca2+-dependent axon-to-soma signaling through local translation of Vim mRNA has also been detected in CNS neurons after exposure to Aβ1–42 (Walker et al., 2018), so at least some of these axon intrinsic signaling mechanisms may extend to the CNS.

III. Mechanisms for gene regulation by axotomy-induced axon-to-nucleus signaling.

The molecular consequences for injury-induced axon-to-soma signaling have been demonstrated for both back-propagating Ca2+ wave and Importin-β/α transport, with large cohorts of genes upregulated or down regulated after axotomy altered by disruption of HDAC5 phosphorylation/nuclear export or Importin-β/α transport (Cho et al., 2013; Perry et al., 2012). This indicates that axon-to-soma signaling not only activates gene programs for axon growth but also represses axon gene expression programs associated with neuronal maintenance/function. Presumably, the axon-derived cargoes of the Importin-β/α complex provide these functions by directly or secondarily regulating transcription factor activity. Histone modification and changes in DNA methylation after injury provide a more indirect modulation of neuronal gene expression. Modifications of histones and DNA have been shown to play important roles in axon regeneration by affecting chromatin structure, thereby modifying chromatin access for transcriptional machinery following injury in both the PNS and CNS (Palmisano and Di Giovanni, 2018). Further, the Ca2+-dependent loss of phospho-HDAC5 from the nucleus was correlated with an increase in the expression of genes associated with transcriptional regulation, so this axon-to-nuclear signaling affects transcription of transcriptional regulators (Cho et al., 2013).

The histone acetyltransferases have also been shown to acetylate transcription factors. Acetylation of lysine residues in the C-terminus of the transcription factor p53 causes an upregulation of Gap43 mRNA during facial motor neuron axon regeneration (Tedeschi et al., 2009). This p53 effect required acetyl transferase activity of the CREB binding protein (CBP). It has also been shown that inducing hyperacetylation of histones by inhibiting HDACs in cultured CNS neurons increases neurite outgrowth in a transcription-dependent fashion (Gaub et al., 2010). Also, axonal HDAC targets can extend beyond histones, as indicated above for HDAC5 targeting acetyl-α-tubulin. HDAC6 is also known to deacetylate α-tubulin, but in contrast to HDAC5 its activity attenuates rather than supports axon growth (Rivieccio et al., 2009). We recently showed that HDAC6 also deacetylates Miro1 in axons, and decreases axon growth by attenuating mitochondrial transport and membrane potential (Kalinski et al., 2019).

Considering that nuclear depletion of HDAC5 induces growth-promoting gene expression, it follows that an upregulation of histone acetyltransferases could similarly increase axon regeneration. Consistent with this notion, overexpression of the histone acetyltransferase p300 in RGCs increases regeneration after optic nerve injury (Gaub et al., 2011). Work from the di Giovanni lab recently showed activation of the related acetyl-transferase CBP increases axon regeneration in both PNS and CNS neurons through modulation of gene expression (Hutson et al., 2019). Interestingly, environmental enrichment triggered an increase in CBP activity that also supported axon regeneration after spinal cord injury (Hutson et al., 2019). It is not clear if this is a retrograde signal derived from the environmental enrichment or soma-restricted mechanism, but these observations emphasize that modulation of histone acetylation after injury brings a potential therapeutic strategy for improving axon regeneration. It should be noted that histone methylation can also impact transcriptional regulation by either inducing or repressing transcription depending on the residues methylated (Wahane et al., 2019). Axon-to-soma modulation of histone methylation status has not been addressed in neurons, but repression of the polycomb complex that catalyzes methylation of H3 at lysine 27 histone attenuates expression of regeneration promoting genes in Schwann cells (Ma et al., 2018).

DNA methylation also contributes to axon regeneration potential and this epigenetic mechanism is specifically regulated by PNS axon injury. In general, DNA hypermethylation is associated with gene repression while hypomethylation is associated with gene activation; however, this does not hold true for all differentially methylated genes after axotomy (Lindner et al., 2014; Puttagunta et al., 2014). Axotomy of the peripheral axons of DRG neurons, not the central axons, leads to an increase in expression of Ten-eleven translocation 3 protein (Tet3), which demethylates DNA by catalyzing the conversion of 5-methylcytosine to 5-hydroxymethylcytosine (5hmC) (Loh et al., 2017; Weng et al., 2017). This injury-induced upregulation of Tet3 requires retrograde Ca2+ signaling, and it leads to an increase in the expression of regeneration-associated genes (RAGs) and axon regeneration (Weng et al., 2017). As mentioned above, DNA methylation can lead to gene silencing and this mechanism has been recently linked to the development of neuropathic pain after nerve injury. After a neuropathic pain inducing peripheral nerve injury, DNA methyltransferase 3α (DNMT3α) mRNA and protein are significantly increased in DRG neurons during the early phase after injury, which in turn contributes to the development of neuropathic pain by down regulating the expression of the Kv1.2 potassium channel (Zhao et al., 2017). In a bone cancer pain model, DNMTs are also increased in the spinal dorsal horn ipsilateral to the bone cancer afflicted limb and inhibiting DNMTs in the spinal cord attenuates the development of pain hypersensitivity associated with a bone cancer pain model by increasing Kv1.2 expression further providing a link between gene silencing of potassium channels due to DNA methylation and neuropathic pain (Miao et al., 2017). DNMT3α is also responsible for silencing the expression of mu and kappa opioid receptors in DRG neurons after neuropathic pain inducing peripheral nerve injury, thereby reducing the response of these animals to opioid analgesics (Sun et al., 2017).

Levels of the epigenetic regulator ubiquitin-like containing PHD ring finger 1 (UHRF1), which plays an important role in DNA methylation and subsequently axon regeneration, increase after peripheral nerve injury. Knocking down UHRF1 in spot cultures of DRG neurons leads to an impairment in axon regeneration, suggesting that some gene silencing is required for axon regeneration (Oh et al., 2018). UHRF1 works to promote axon regeneration after peripheral axotomy by effectively silencing the transcription of phosphatase and tensin homolog (PTEN) and RE1 silencing transcription factor (REST) through methylation of their promoter regions (Oh et al., 2018). Taken together, these and other observations indicate that injury-induced axon-to-soma signaling alters somal gene expression through both direct transcriptional and epigenetic mechanisms, with the combination undoubtedly responsible for the broad change in gene expression programs needed to shift from a maintenance/function to regeneration phenotype.

IV. Post-transcriptional regulation of gene expression during nerve regeneration.

Role of localized mRNAs in axon growth and regeneration –

Regulation of gene expression extends to synthesis of proteins, and many lines of evidence indicate that neuronal mRNAs are subjected to several different levels of post-transcriptional control. Indeed, localizing mRNAs to axons, modifying an mRNA’s translational efficiency, or altering survival of an mRNA can dictate when a protein is introduced, where the proteins is introduced, and how much of that protein is generated. Intra-axonal translation of mRNAs was mentioned above for the Kpnb1, Ranbp1, Vim, and some transcription factors, but this mechanism also extends to mRNAs whose proteins products directly regulate axon growth (Figure 1A). For example, axonally synthesized proteins are needed for growth cone formation after in vitro axotomy in DRG neurons (Chierzi et al., 2005). Though it is not clear which nascently synthesized proteins are needed for the growth cone formation, Pacheco et al. (2020) recently showed that translation of axonal Calr mRNA supports initiation of axon growth in axotomized DRG neurons (Pacheco et al., 2020). Axonal Calr translation is increased by phosphorylation of eIF2α (Vuppalanchi et al., 2012), and, as an avid Ca2+ binding protein (Gelebart et al., 2005), this raises the intriguing possibility that axonal Calr protein allows growth cone formation by chelating axoplasmic Ca2+. The injury induced axon-to-soma signaling can also change the repertoire of mRNAs in axons, with mRNAs encoding RAGs showing increased transport into axons following axotomy (Figure 1A) (Kar et al., 2018). Subcellular localization of mRNAs is dictated by sequence motifs within the mRNAs (_cis_-elements) that are recognized by RNA Binding Proteins (RBPs; _trans_-factors) to form ribonucleotide protein (RNP) complex. These RNPs associate with kinesin for anterograde transport into axons, with the moving RNPs referred to ‘transport granules’. So far, no unique themes for _cis_-element mRNA sequences have been identified that allow predicting whether an mRNA will localize, and studies in cultured neurons, and recently in vivo, have identified hundreds to thousands of axonally localized mRNAs (Briese et al., 2016; Gumy et al., 2011; Hafner et al., 2019; Kar et al., 2018; Minis et al., 2014; Rotem et al., 2017; Shigeoka et al., 2016; Zivraj et al., 2010). However, relatively few RBPs have been defined for axonal mRNAs. We recently used experimentally defined _cis_-elements from several axonal mRNAs to expand knowledge of axonal RBPs. These proteomics results showed that multiple axonal RBPs can interact with the same _cis_-element, but also emphasize that many RBPs that were thought to be restricted to the nucleus localize into PNS axons (Lee et al., 2018). Interestingly, axonal localization hnRNPs K, H1, and F increases after axotomy and knockdown of these and related RBPs points to roles in axon growth (Lee et al., 2018), further emphasizing the role of axonal mRNA-RBP interactions in axon regeneration. Table 1 summarizes several of the identified axonal mRNA localizing _cis-_elements and corresponding RBPs. A few of these warrant further description for their potential roles in axon growth and regeneration and to emphasize the complexities of RBP-mRNA and RBP-RBP interactions in axons.

Table 1:

Known axonal RNA motifs determining axonal mRNA localization. Beyond the localizing _cis_-elements outlined below, Martinez et al. (2019) recently reported that RNA binding motifs for Pumilo proteins (Pum1 and Pum2) serve as signals to retain mRNAs in cell bodies of PNS and CNS neurons.

Zip Code Binding Protein 1 (ZBP1, also called IGF1 binding protein 1 [IMP1]) was shown to bind a 54 nucleotide element in the 3’UTR of β-Actin (Actb) mRNA for localization to the periphery of migrating fibroblasts and growing axons (Kislauskis et al., 1994; Zhang et al., 1999). Introducing a transgene with the Actb 3’UTR into mouse PNS neurons decreased axonal levels of Actb and Gap43 mRNAs and decreased axon regeneration after axotomy (Donnelly et al., 2011). Thus, ZBP1 is needed for axonal transport of both of these mRNAs, and limited availability of ZBP1 in adult PNS neurons restricts axon regeneration capacity (Donnelly et al., 2011). This also emphasizes that ZBP1 binds to multiple mRNAs, and RNA co-IP studies in non-neuronal cells support this idea but also emphasize that ZBP1 interacts with many other RBPs (Jonson et al., 2007). Such RNA-RBP interactomes in other cellular systems have been defined as “RNA regulons” (Keene, 2007). Splicing Factor Proline And Glutamine Rich (SFPQ), which co-assembles with Bcl w and LaminB2 mRNAs into RNPs for axonal localization (Cosker et al., 2016), and Nucleolin, which is needed for transport of Kpnb1 and mTor mRNAs (Perry et al., 2016; Terenzio et al., 2018), provide two more examples. Bclw and LaminB2 proteins support survival of axons (Cosker et al., 2016; Yoon et al., 2012), while Importin-β1 and mTOR are injury response proteins that help to initiate axon regeneration responses (Perry et al., 2016; Terenzio et al., 2018). Though some headway has been made in identifying axonal RNA regulons, much more systematic analyses of the axonal mRNAs bound to axonal RBPs are needed to fully understand how RBP-mRNA interactions contribute to axon regeneration.

RNA-seq analyses of axonal hnRNP K, F, and H1 immunoprecipitates define RNA regulons supporting axon growth by bioinformatics (Lee et al., 2018), but these need to be distinguished for direct vs. indirect interactions as the transport RNPs can have many different RBPs. For example, axonal Actb mRNA has been shown to interact with ZBP1, hnRNP R, spinal motor neuron (SMN) and HuD (also called Elavl4) proteins (Glinka et al., 2010; Kim et al., 2015b; Rossoll et al., 2003; Tiruchinapalli et al., 2003). While hnRNP R and ZBP1 contribute to Actb mRNA axonal localization in motor neurons, ZBP1 and HuD are necessary for axonal localization in sensory neurons (Glinka et al., 2010; Kim et al., 2015b; Rossoll et al., 2002). ZBP1’s interaction with Gap43 mRNA requires HuD binding to the AU rich element (ARE) in Gap43 mRNA’s 3’UTR (Yoo et al., 2013). The complement of proteins binding to the individual mRNAs likely determine its fate for subcellular localization, but mRNAs may behave differently in different neuron types as suggested by hnRNP R and HuD roles in Actb localization in motor vs. sensory neurons. Further differences in the axonal transcriptome of neuron types and physiological conditions will undoubtedly be uncovered by systematic analyses of transcriptomes and axonal RNA regulons. For example, the mRNA encoding γ-Actin (Actg) does not localize to axons of cortical or sensory neurons (Bassell et al., 1998; Willis et al., 2005), but it does localize to axons of embryonic motor neurons (Moradi et al., 2017). Similar to Actb and other axonal mRNAs, the 3’UTR of Actg drives its localization in motor neurons (Moradi et al., 2017). The molecular basis of this differential Actg localization is not known beyond the functions of its 3’UTR, but this will likely be explained by different populations of RBPs interacting with this Actg 3’UTR in motor vs. sensory and cortical neurons. Also of interest is that the Hengst lab very recently uncovered a mechanism restricting mRNAs from axonal localization; specifically, they found that the Pumilio 1 and 2 (Pum1 and Pum2) binding element (PBE)-containing mRNAs are restricted to the soma of developing neurons (Martinez et al., 2019). It will be telling to see if this PBE-based mechanism is differentially regulated in different neuron types and physiological conditions.

Translational control for axon growth and regeneration –

Several lines of evidence indicate that mRNAs are stored until needed in PNS axons, which provides an on-demand system for generating new proteins locally in response to different stimuli. Evidence for this is seen with near immediate axonal translation of the injury-response gene mRNAs after axotomy that are outlined above (Figure 1A) (e.g., Importin-β1, RanBP1, STAT3 (Ben-Yaakov et al., 2012; Hanz et al., 2003; Yudin et al., 2008)). Extracellular stimuli that axons encounter have also been shown to activate translation of different axonal mRNAs. For example, translation of axonal RhoA and Cofilin mRNAs is up regulated by growth inhibitory signals (Piper et al., 2006; Wu et al., 2005) and Actb, the actin nucleating Arp2/3 complex activator WAVE1, and the actin complex stabilizer Cortactin mRNAs are upregulated by neurotrophins (Spillane et al., 2011; Willis et al., 2007). Exposure and withdrawal of NGF was shown to trigger the translation of axonal Pafah1b1 (Lis1) mRNA, with the end result determined by whether or not the mRNA is bound to Adenomatous Polyposis Coli (APC) protein (Villarin et al., 2016). APC has been shown to localize mRNAs to microtubule plus-ends, and spatially orchestrate protein synthesis in axons and growth cones (Preitner et al., 2014).

Two general mechanisms have been uncovered beyond the direct RBP-mRNA interactions to account for how stimuli, including axonal injury, can recruit mRNAs into translation. The Flanagan lab published a provocative mechanism where mRNAs and translational machinery interact with the cytoplasmic tail of the transmembrane receptor deleted in colon cancer (DCC). Binding of Netrin ligand to DCC receptor led to dissociation of these mRNAs and formation of actively elongating polysomes (Tcherkezian et al., 2010). Elegant work from the Holt lab very recently extended this observation to other axon guidance receptors (Koppers et al., 2019). We recently showed that the axonal mRNA interaction with the stress granule (SG) associated protein, Ras-GAP SH3 domain Binding Protein1 (G3BP1), provides a mechanism for activating translation of quiescent axonal mRNAs after axotomy (Sahoo et al., 2018a). Given the link to injury and regeneration, we will expand on this mechanism below; however, the possibility that receptor-mediated axonal mRNA storage contributes to regeneration has not been tested, so we do not intend to imply that mechanism is not relevant for regeneration (or axon degeneration).

SGs are used by other cell types to store untranslated mRNAs after exposure to oxidative or metabolic stress (Anderson and Kedersha, 2008). Several SG proteins have been documented in PNS axons (Murashov et al., 2007; Price et al., 2006). We found that the classic SG proteins G3BP1, Fragile X mental retardation syndrome-related protein 1 (FXR1), Fragile X mental retardation protein (FMRP), T-cell restricted intra-cellular antigen 1 (TIA1), and Human antigen-R (HuR) form SG-like structures in PNS axons under basal conditions and mRNAs are associated with these axonal SG-like structures (Sahoo et al., 2018a). Much effort for the study of SG proteins and their aggregation has been focused on neurodegenerative diseases, particularly amyotrophic lateral sclerosis and frontotemporal dementia (Mackenzie et al., 2017). G3BP1-containing granules were also recently shown to hitchhike on moving lysosomes to reach distal axons (Liao et al., 2019). In both mammalian and C. elegans axons, the SG protein granules rapidly increase in size after axotomy. For C. elegans blocking this aggregation through depletion of TIAR2, the orthologue of mammalian TIA1, increased axonal regeneration (Andrusiak et al., 2019). For mammalian PNS neurons, blocking G3BP1 aggregation by expression of G3BP1 acidic domain or a cell-permeable peptide from the acidic domain increased axonal protein synthesis and accelerated in vivo axon regeneration (Sahoo et al., 2018a). These data emphasize that SG proteins act to attenuate axon growth rates. Interestingly, aggregation of SG proteins has been demonstrated in several neurodegenerative conditions, including aggregation in axons (Khalfallah et al., 2018; Maziuk et al., 2017; McDonald et al., 2011), and mutations in TIA1 protein in amyotrophic lateral sclerosis suggest SG proteins aggregation can cause neuron loss (Mackenzie et al., 2017). With regards to the topic of this special issue, the possibility that SG protein aggregation occurs in peripheral neuropathies has not been addressed.

As implied by the text above and Table 1, regulatory elements or motifs within mRNAs determine their post-translational regulation, including recognition by the non-coding RNAs (ncRNAs) outlined in the next section. Stimuli from the environment of the axon have been shown to impact axonal mRNA transport and translation. This has been documented for axon guidance cues in developing axons and for trophic factor stimulation of mature axons (Sahoo et al., 2018b). For example, NGF-stimulated axon growth requires translation of axonal Par3 mRNA in developing sensory neurons (Hengst et al., 2009) and growth attenuation by CSPGs requires translation of axonal RhoA mRNA (Walker et al., 2012). Willis et al. (2007) showed altered transport for entire cohorts of mRNAs in adult sensory neurons in response to growth promoting vs. growth-inhibiting stimuli, with specific mRNAs accumulating or being depleted from focal stimulation points (Willis et al., 2007). The signals triggered by these extracellular stimuli likely impact functions of the RBPs or combinations of RBPs bound to these mRNAs. It will be important to understand how RBP-driven mechanisms intersect with the translational control pathways discussed herein and considering how we might utilize these mechanisms for neural repair strategies.

Contributions of non-coding RNAs to axon growth and regeneration –

Numerous studies have shown the role of non-coding RNA (ncRNA)-mediated mechanisms, particularly through micro-RNAs (miRs), in neuronal development, including critical roles in neural induction/patterning, neural precursor cell proliferation, migration, differentiation, axon growth and synaptogenesis (Cho et al., 2019; Motti et al., 2012). miRs work by inhibiting translation of target mRNAs or triggering degradation of target mRNAs, but it should be noted that in silico predicted mRNA targets for a given miR must be validated to ensure that the mRNA is actually a target (e.g., secondary/tertiary RNA structure or interacting proteins could hide an miR binding site in an mRNA). Nonetheless, an extensive body of evidence also supports roles for miRs in neuronal injury and disease (Cho et al., 2019). The Murashov lab showed that genetic ablation of dicer, a central regulator of miR biogenesis, impeded sciatic nerve regeneration, attenuating anatomical, physiological and functional recovery (Wu et al., 2012). A number of studies using both microarray and deep sequencing approaches have identified miRs whose levels are affected in both spinal cord and sciatic nerve injury (Table 2) (Motti et al., 2017; Strickland et al., 2011a; Strickland et al., 2011b; Wu and Murashov, 2013; Yu et al., 2011; Zhang et al., 2011; Zhou et al., 2011). By integrating the results from the miR profiles with the transcriptomics data from injured DRGs, candidate molecular pathways affected by the regulated miRs were identified. These pathways represent functions in neuronal differentiation and activation of stem cell niches intrinsic growth capacity, Schwann cellneuron communication, cellular morphogenesis, cell polarity and cytoskeletal reorganization (Motti et al., 2017; Strickland et al., 2011a; Yu et al., 2011). We outline a few examples of miRs with validated neuronal mRNA targets involved in axon growth and regeneration below, but refer the reader to the references herein and in Table 2 for a more formative description of the injury-regulated miRs and putative targets.

Table 2:

Micro-RNAs affecting axonal growth

Several validated miR-mRNA target pairs have been identified for neural injury and regeneration responses. miR-21 that is upregulated in DRGs after injury has been shown to modulate Sprouty2 mRNA levels (Strickland et al., 2011b). Sprouty2 is a Ras/Raf/ERK pathway inhibitor that blocks brain-derived neurotrophic factor (BDNF)-induced neuronal differentiation and survival (Gross et al., 2007), so decreasing Sprouty2 would increase pro-growth signaling from BDNF. miR-21 was also shown to be upregulated in injured spinal cord after exercise, which is known to increase functional recovery after spinal cord injury (Liu et al., 2012). Peripheral nerve injury led to down regulation of miR-145 and −214 that regulate the levels of Robo2 and srGAP2 proteins to increase neurite growth and regeneration (Zhang et al., 2011). PNS nerve injury increases miR-222 levels that promotes neurite growth by regulating levels of PTEN (Zhou et al., 2012). Previous studies from the He lab have shown that PTEN knockout supports axon regeneration after optic nerve injury and spinal cord injury (Park et al., 2010). miR-21 mentioned above is also predicated to target PTEN (Liu et al., 2012), though this has not been validated with miR overexpression and knockdown studies. miR-431 is upregulated in DRGs after sciatic nerve crush and targets the Wnt signaling inhibitor Kermen 1 to promote axon regeneration in DRGs (Wu and Murashov, 2013). Wnt signaling has been linked to spinal cord injury and regeneration, so it will be of interest to determine if miR-431 contributes to CNS axon regeneration (Fernandez-Martos et al., 2011; Gonzalez-Fernandez et al., 2014). Together, these and other studies emphasize the role(s) for ncRNAs in PNS and CNS injury responses and regeneration.

The above-mentioned studies focused on miRNAs present in soma of injured neurons or did not distinguish between somal and axonal roles. Work in cultured neurons has clearly shown that miRs localize into axons (Gumy et al., 2011; Natera-Naranjo et al., 2010). Phay et al. (2015) employed deep sequencing (RNA-seq) to identify miRs present in axoplasmic extracts from injured and regenerating rodent sciatic nerves; they found 141 axonal miRNAs with altered levels in injured relative to uninjured axons. Previous studies have shown that axons contain enzymes needed for miR biogenesis and precursor-miRNAs (pre-miR) can be locally processed and function to modulate axonal activity and function (Aschrafi et al., 2008; Aschrafi et al., 2012; Hengst and Jaffrey, 2007; Kar et al., 2013). This raises the appealing hypothesis that axonal injury may locally control miR function by regulating processing of pre-miR into mature miRs within axons. Consistent with this, Kim et al. (2015) showed the presence of pre-miRs in sciatic nerve axons, and they found that injury can indeed trigger processing to generate mature miRs from these precursors (2015). Recently it was shown that some pre-miRs are actively trafficked to distal axons of RGC neurons by hitchhiking on late endosomes/lysosomes, and these pre-miRs are then processed into mature miRs upon exposure to axon guidance cues (Corradi et al., 2020). As we point out below, this endosome/lysosome mediated transport of RNAs, including pre-miRs, could be affected in peripheral neuropathies. Taken together, there is substantial evidence that, changes in both the axonal and somal miR populations play an important role in modulating regenerative capacity after injury and may serve as potential targets for increasing growth of injured axons. Further systematic studies will be needed to fully understand the dynamics of miR function in neurons and how these miRs impact injury-induced gene expression at both global and local levels. The post-transcriptional mechanisms outlined in this section and above emphasize that mRNAs lead a rich life after transcription and restricting study to only levels of RAG mRNAs gives an incomplete picture of the protein changes that are driving regeneration.

V. Growth cone formation and cytoskeletal dynamics for axon growth.

A critical feature of successful axonal regeneration is the ability to initiate and support axon extension. Although there are reports of axon regeneration in the apparent absence of a growth cone (Jin et al., 2016), a general assumption is that the injured axon forms a growth cone that is the site of axon extension. Indeed, one aspect that differentiates injured PNS from CNS axons is the ability to form growth cones in the PNS rather than retraction bulbs as seen in the CNS (Tom et al., 2004). As outlined above, growth cone formation requires Ca2+ influx and in vitro work suggests that intra-axonal protein synthesis is also needed (Blanquie and Bradke, 2018; Chierzi et al., 2005; Erturk et al., 2007; Kamber et al., 2009). The growth cone is comprised of the following components: the peripheral domain (P domain), which includes dynamic F-actin filled filopodia and lamellipodia, a more stable central domain (C domain) containing microtubules, organelles and vesicles, and a transitional domain between the P and C domains containing the motor protein myosin II (Dent et al., 2011). Axonal outgrowth through the growth cone involves three stages: protrusion, engorgement, and consolidation. For protrusion, F-actin filled filopodia and lamellipodia extend from the leading edge of the growth cone (Lowery and Van Vactor, 2009). During engorgement, dynamic microtubules in the C domain enter the newly formed protrusions bringing along vesicles and organelles (Lowery and Van Vactor, 2009). Consolidation occurs with depolymerization of F-actin in the neck of the growth cone with the axon shaft established by compaction of microtubules (Lowery and Van Vactor, 2009). Regulation of the dynamics of these cytoskeletal components by extracellular stimuli is a prominent feature of developmental axon guidance and contributes to axon regeneration; further, work linking the tubulin binding protein APC to mRNAs brings a potential mechanism to alter the proteome of the growth cone (Preitner et al., 2014).

Cytoskeletal reorganization and dynamic cytoskeletal elements in the axon are essential for growth cone formation after nerve injury; local availability of monomer proteins for polymerization of microfilaments and microtubules, levels and activity of binding proteins, and post-translational modifications drive polymerization and stabilization/destabilization of these cytoskeletal elements in the distal axon/growth cone (Dent et al., 2011). Actin dynamics in the filopodia and lamellipodia are needed for growth cone navigation (Geraldo and Gordon-Weeks, 2009) and microfilament turnover in the P domain is necessary for growth cone motility (Difato et al., 2011). Actin depolymerizing factor (ADF)/cofilin is one of the proteins responsible for actin depolymerization in axons and is necessary for regeneration of the centrally projecting DRG axons after a conditioning lesion (Tedeschi et al., 2019), and notably ADF/Cofilin is locally translated in axons (Bellon et al., 2017; Piper et al., 2006; Willis et al., 2007). Axotomy of RGCs causes F-actin destabilization and the Doublecortin and Doublecortin-like kinases help stabilize F-actin in injured axons (Nawabi et al., 2015).

Microtubule dynamics in the growth cone are essential for axon extension (Hur et al., 2012). Acetylated α-tubulin tends to be more prevalent in regions of stable microtubules, while tyrosinated α-tubulin is more prevalent in regions with dynamic microtubules. In PNS neurons, levels of acetylated α-tubulin decrease in axons near the injury site with a concomitant increase in tyrosinated α-tubulin (Cho and Cavalli, 2012). Attenuating tyrosination of α-tubulin significantly reduces PNS sensory axon regeneration by decreasing retrograde injury signaling and delaying the activation of a pro-regenerative program (Song et al., 2015). HDAC inhibitors can block tubulin deacetylation and reduce PNS axon growth; as noted above, HDAC5 localization into distal axons after injury is thought to trigger deacetylation of axonal α-tubulin in the PNS. Tubulin deacetylation is responsible for supporting dynamic microtubules thereby contributing to growth cone dynamics needed to support axon outgrowth, at least in PNS neurons (Cho and Cavalli, 2012). This injury-induced tubulin deacetylation is specific to PNS neurons and does not occur to the same extent in the CNS (Cho and Cavalli, 2012). Intriguingly, microtubule stabilization has been shown to increase regeneration in CNS axons following spinal cord injury (Ruschel et al., 2015) and inhibition of HDAC6, which increases axonal levels of acetylated α-tubulin, promotes axon growth on non-permissive substrates that are part of the growth inhibitory environment of the injured CNS (Rivieccio et al., 2009). The recent report that HDAC6 deacetylates Miro1 in axons (Kalinski et al., 2019) emphasizes that HDAC6, and likely HDAC5, has other axonal substrates that contribute to its effects on axon growth. Also, the divergent effects of microtubule stability and post-translational modifications in PNS and CNS neurons may reflect differences between neuron types or distinct requisites of regenerating in permissive (PNS) vs. non-permissive environments (CNS). In addition, changes in the status of acetylation and tyrosination of α-tubulin also affects the interaction of motor proteins – e.g., Kinesin-1 and-3 have been reported to differentially prefer acetylated or tyrosinated microtubules, respectively (Cai et al., 2009; Guardia et al., 2016; Lipka et al., 2016). These findings raise the possibility that post-translational modifications of α-tubulin affect the subcellular transport and localization of organelles and RNP granules, which in turn affects the local proteome and influence axon integrity and regenerative ability. Notably, inhibition of HDAC6 is axon protective in both inherited and acquired peripheral neuropathies (Benoy et al., 2018; d’Ydewalle et al., 2011; Krukowski et al., 2017; Van Helleputte et al., 2018), and HDAC6 has been shown to interact with as well as target protein synthesis machinery and RBPs outlined above (Cao et al., 2019; Kwon et al., 2007; Mo et al., 2018). Thus, HDAC6 provides a clear mechanistic intersection for pathways involved in axon regeneration and axon degeneration.

VI. Contributions of non-neuronal cells in nerve injury and axon regeneration.

The sections above regard the axon as an entity dependent solely on the soma for responses to injury and ability to regenerate. However, the axon clearly is impacted by and senses its local environment, and non-neuronal cells as well as factors secreted by those clearly impact injury responses and axon growth (Figure 1A). For example, Schwann cells have a major impact on regeneration. Indeed, preventing Schwann cells from migrating in regenerating nerves impairs axon regeneration (Chen et al., 2005). Work from Toth et al. (2009) showed that secretion of calcitonin gene-related polypeptide (CGRP) from axons supports Schwann cell migration (Toth et al., 2009). Growth factors secreted by denervated Schwann cells serve as tropic stimuli for axon regrowth (Gordon, 2009). For example, Schwann cell secretion of BDNF in injured nerves activates axonal Tropomyosin Receptor Kinase B (TrkB) to support regenerative growth of motor axons through axon-to-soma signaling (English et al., 2014). Interestingly, evidence points to unique gene expression signatures of Schwann cells from motor vs. sensory nerves (Brushart et al., 2013; Hoke et al., 2006), so the Schwann cell may be tuned to support growth of its surrounding axons. Schwann cells in the PNS as well as oligodendrocytes in the CNS serve as a nutrient source for the distal axon by providing lactate and pyruvate to feed into axonal ATP production (Jha and Morrison, 2018). Finally, there is some evidence that Schwann cells can transfer macromolecules to axons, as Schwann cell-derived ribosomes have been detected in injured and regenerating axons (Court et al., 2008; Court et al., 2011), and a recent study suggested that glia are a major source of axonal ribosomes after axotomy (Muller et al., 2018). Schwann cell-derived exosomes have further been shown to increase axon growth in PNS neurons (Lopez-Verrilli et al., 2013). It is appealing to hypothesize that these exosomes transfer mRNAs or ncRNAs to axons, though what exactly is the growth promoting content of these Schwann cell packets can only be inferred at this point (Lopez-Leal and Court, 2016). Together, the above and other observations point to Schwann cell-to-axon signals that can support axon growth, function, and survival, and raise the potential for reciprocal axon-to-Schwann cell signaling.

Inflammatory cells infiltrate the nerve and directly or indirectly impact nerve regeneration. An inflammatory response can occur within a few minutes to hours after injury and is important for axon regeneration (Benowitz and Popovich, 2011). Granulocyte and monocyte infiltration into injured PNS nerves was suggested to contribute to regeneration by producing growth-supporting neurotrophic factors (Barrette et al., 2008). In line with this, exogenous interleukin (IL)-6 and interferon γ (IFNγ) increase neurotrophin-mediated axon outgrowth in DRG explants (Golz et al., 2006). Macrophage-derived IL-1 promotes Schwann cell synthesis of NGF in the distal nerve after injury (Lindholm et al., 1987). Macrophage infiltration into the DRG after peripheral nerve injury neurons also impacts sensory axon regeneration, and release of the C-C motif chemokine ligand 2 (CCL2) recruits macrophages to denervated DRGs (Lindborg et al., 2018; Niemi et al., 2013). Inflammatory cell-derived factors can also directly affect regeneration by acting on neurons. Leukemia inhibitory factor (LIF), a member of the gp130 cytokine family, plays an important role in signaling the immediate inflammatory cell infiltration into the injury site (Sugiura et al., 2000). The increased axon regeneration seen after a conditioning-injury is blunted in LIF knockout mice (Cafferty et al., 2001). LIF was shown to be retrogradely transported in sensory neurons after injection into the sciatic nerve indicating LIF could directly drive axon-to-soma injury signaling effects (Lindholm et al., 1987). Optic nerve injury causes a significant increase in IL-6 expression, and it can increase RGC survival and neurite outgrowth (Leibinger et al., 2013) as well as contribute to injury-conditioning responses in the PNS (Cafferty et al., 2004). IL-6 administration led to increased levels of regeneration-associated genes RAGs such as Sprr1a, Galanin, and Gap43 mRNAs in RGCs raising the possibility for IL-6 activating axon-to-nuclear signaling (Leibinger et al., 2013). Inflammatory cell-derived IL-2, IL-10, tumor necrosis factor-β, and transforming growth factor-β have also been linked to peripheral nerve injury (Fregnan et al., 2012), and some of these factors may affect CNS regeneration (Tsarouchas et al., 2018). Of note, work from the Price lab has shown that IL-6 can affect protein synthesis in intact sensory axons and trigger axon-to-nucleus signaling through localized CREB synthesis after application of neuropathic pain-invoking stimuli (Melemedjian et al., 2010; Melemedjian et al., 2014 ). Thus, regeneration-supportive inflammatory vs. pathologic inflammatory responses in the PNS could be determined by synergy between cytokine and neurotrophin signaling pathways as suggested by Vidal et al. (2013).

VII. Axon intrinsic signaling mechanisms and peripheral neuropathies

With reference to this issue of Brain Research, it should be noted that the injury response mechanisms outlined above overlap with the pathophysiology of some neuropathic conditions. We have attempted to raise these in the sections above, but appealing links between axonal mRNA transport/translation and axon function warrant further attention. In particular, alterations in axonal mRNA transport have been demonstrated in a chemotherapy-induced peripheral neuropathy. The Segal lab found that axonal transport of RNA granules containing the RBP SFPQ is decreased with paclitaxel exposure, thereby decreasing axonal synthesis of the anti-apoptotic protein Bclw whose RNA these granules transport (Figure 2) (Pease-Raissi et al., 2017). Axonal Bclw contributes to axon survival by binding to IP3R on ER preventing Ca2+ release (Figure 2) (Pease-Raissi et al., 2017). Axonal levels of the mRNAs for Mitofusin 2 (Mfn2), Opa1 Mitochondrial Dynamin Like GTPase, and Dynamin-1-like protein (Drp1), proteins that function in mitochondrial fusion and fission, are similarly decreased after exposure to paclitaxel (Bobylev et al., 2015). These observations indicate axonal protein synthesis can be disrupted in acquired neuropathies, and considering the proteins involved, this can directly affect axon survival and function. As we outline below, cellular mechanisms impacted in inherited neuropathies, including the varying forms of Charcot Marie Tooth (CMT) neuropathies, could have direct impact on the growth-associated axon-intrinsic functions outlined above.

Work in several labs now points to co-transport of axonal mRNAs and translational machinery on endocytic and lysosomal vesicles (Cioni et al., 2019; Liao et al., 2019; Rangaraju et al., 2019). This may help to explain earlier works pointing to association of RBPs (e.g., SMN and HTT) with membrane-bound organelles and organelle associated proteins (e.g., CopB1) binding to localized mRNAs (Bi et al., 2007; Custer et al., 2013; Custer et al., 2019; Savas et al., 2010; White et al., 2015). With regards to neuropathies, mutations of a number of proteins that cause demyelinating CMT are engaged in varying steps of the endocytic pathway that generates vesicles, including endocytosis, endosomal recycling, and endosome-to-lysosome trafficking (For review see Lee et al., 2017). Disrupted endocytic trafficking due to CMT-associated mutations leads to dysregulation of a number of critical intracellular signaling pathways (e.g., ERK, JNK1, AKT, and PI3K/AKT) that in turn can impact axonal maintenance/function through Schwann cell-to-axon signaling mechanisms (Fledrich et al., 2014; Kohl et al., 2010; Massa et al., 2006). More directly, the Holt lab recently showed that Rab7a mutants associated with CMT2B in RGC axons directly impairs axonal protein synthesis with loss of axon integrity (Cioni et al., 2019). The causative gene for CMT4J, Fat-induced Gene 4 (Fig4 also called Sac3) that encodes a phosphoinostide 5-phosphatase that regulates intracellular levels of PI(3,5)P2 and PI(5)P, plays a critical role in endosome-to-lysosome maturation and trafficking. Fig4-deficient animals show abnormal endosomes and lysosomes, suggesting the involvement of impaired endolysosomal trafficking in CMT4J pathogenesis (Chow et al., 2007; Lenk et al., 2011; Winters et al., 2011). It will be intriguing to see if RNA trafficking and axonal translation are disrupted in CMT4J and other inherited neuropathies.

Mitochondrial function is critical for maintaining axon integrity and mutations in proteins that cause altered mitochondrial transport and function can result in axonal forms of CMT (Kalmar et al., 2017; Rocha et al., 2018). These include MFN2 and HSPB1 whose mRNAs are also localized into PNS (Bobylev et al., 2018; Willis et al., 2005; Willis et al., 2007). mRNAs for several nuclear-encoded mitochondrial proteins also localize into the axons and their localized translation helps to maintain mitochondrial function in distal axons (Aschrafi et al., 2016; Fazal et al., 2019; Kaplan et al., 2009; Yoon et al., 2012). Similarly axonal synthesis of LaminB supports axonal integrity at least in part by supporting mitochondrial function (Yoon et al., 2012). Decreased mitochondrial respiration with resulting fall in nicotinamide adenine dinucleotide (NAD) has long been linked to Wallerian degeneration and likely contributes to other forms of axon degeneration (Conforti et al., 2014). Observations that mitochondrial ATP synthesis supports local mRNA translation during plasticity and axon branching (Rangaraju et al., 2019; Spillane et al., 2012) and axonally synthesized proteins supporting mitochondrial function (Aschrafi et al., 2012; Cioni et al., 2019; Natera-Naranjo et al., 2012) bring an interesting bidirectional network that can support axon integrity, function, and branching. The possibility that this axon-intrinsic network plays a role in the pathophysiology of peripheral neuropathy may bring new therapeutic targets to prevent axon loss and/or to encourage branching of residual axons that can reinnervate target tissues and help to restore some function.

CONCLUSION

Much progress has been made to understand how the intrinsic growth program of axons is initiated after injury and the critical mechanisms that allow the PNS to regenerate. Injury of the distal axon provides multiple paths for axon-to-soma and axon-to-nucleus signaling that shift the neuron’s phenotype to one supporting regeneration. This subsequently provides soma-to-axon signaling that is needed for effective regeneration and eventual functional recovery. Clearly more effort is needed to determine the intricacies of how these mechanisms support regeneration and how they are regulated. This is particularly true if this knowledge will be leveraged to develop effective therapies to improve neural repair in the PNS and CNS. As we note above, loss of some of these mechanisms can also contribute to peripheral neuropathic conditions, with clear links between axonal mRNA transport and translation to chemotherapy-induced peripheral neuropathy and very intriguing potential links to inherited peripheral neuropathies like CMT. Therapies targeting some of these mechanisms may indeed provide dual functions for neuropathic conditions by supporting axon survival/integrity as well as encouraging axon branching for reinnervation of denervated target tissues. However, we need a much better understanding for how broadly axonal RNA transport/translation or other axon-intrinsic mechanisms outlined above extend to inherited and acquired neuropathic conditions.

HIGHLIGHTS FOR REVIEW.

• Axonal injury in the PNS invokes axon-to-soma and axon-to-nucleus signals through back-propogating wave of Ca2+ and retrograte transport of signaling proteins that change gene expression to support regeneration through transcriptional and epigenetic mechansims.
• Part of the retrograde signaling is generated through localized synthesis Importin-β protein and transcription factors, whose intra-axonal translation is activated by increased axoplasmic Ca2+
• Subsequent soma-to-axon signaling helps to support axon growth by providing both proteins and mRNAs, with the latter used to locally generate proteins to support growth.
• Injured CNS axons share some of the same signaling mechanisms but do not appear to invoke these with traumatic injury, which may help to explain the decreased regeneration capacity of CNS as compared to PNS neurons.
• Axon degeneration can also be triggered by increase in axoplasmic Ca2+, particularly by release from intracellular stores including the ER. Interestingly, depletion of ER Ca2+ can trigger an unfolded protein response locally in axons which helps to initiate axon regeneration mechanisms with traumatic injury but also leads to axon death in chemotherapy induced neuropathy.

Our manuscript discusses these mechanisms and draws parallels between injury-response that leads to successful axon regeneration vs. known mechanisms of axon degeneration.

Acknowledgements –

The authors have been supported by the following grants from the following agencies in research directly related to the topic of this review: NIH/NINDS (K01-NS105879 to TS; R01-NS089633 and R01-NS041596 to JLT), NSF (MCB-1020970 to JLT), Department of Defense, US Army Medical Research and Development (W81XWH-13-1-0308 to JLT), the Dr. Miriam and Sheldon G. Adelson Medical Research Foundation (to JLT), South Carolina Spinal Cord Injury Research Fund (2018 PD-01 to PKS), and the UofSC Research Office ASPIRE program (to ANK). JLT is the incumbent South Carolina SmartState Endowed Chair in Childhood Neurotherapeutics at the Univ. of South Carolina.

Footnotes

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