Defects in Axonal Transport in Inherited Neuropathies

Abstract

Axonal transport is a highly complex process essential for sustaining proper neuronal functioning. Disturbances can result in an altered neuronal homeostasis, aggregation of cargoes, and ultimately a dying-back degeneration of neurons. The impact of dysfunction in axonal transport is shown by genetic defects in key proteins causing a broad spectrum of neurodegenerative diseases, including inherited peripheral neuropathies. In this review, we provide an overview of the cytoskeletal components, molecular motors and adaptor proteins involved in axonal transport mechanisms and their implication in neuronal functioning. In addition, we discuss the involvement of axonal transport dysfunction in neurodegenerative diseases with a particular focus on inherited peripheral neuropathies. Lastly, we address some recent scientific advances most notably in therapeutic strategies employed in the area of axonal transport, patient-derived iPSC models, in vivo animal models, antisense-oligonucleotide treatments, and novel chemical compounds.

Keywords

Axonal transport inherited peripheral neuropathies Charcot-Marie-Tooth disease genetics neurodegeneration cytoskeleton molecular motors cargoes therapeutics

COMMON ABBREVIATIONS

actin filament

ALS

amyotrophic lateral sclerosis

CMT

Charcot-Marie-Tooth

dHMN

distal hereditary motor neuropathy

endoplasmic reticulum

HDAC6

histone deacetylase 6

HSAN

hereditary sensory and autonomic neuropathy

HSP

hereditary spastic paraplegia

IPN

inherited peripheral neuropathies

iPSC

induced pluripotent stem cell

MIM

mitochondrial inner membrane

MOM

mitochondrial outer membrane

microtubule

MTA

microtubule-targeting agents

NEFL

neurofilament light

NEFM

neurofilament medium

NEFH

neurofilament heavy

neurofilament

NMJ

neuromuscular junction

Perry syndrome

PTM

post-translational modifications

RBP

RNA-binding protein

RNP

ribonucleoprotein particles

SARM1

Sterile Alpha and TIR Motif Containing 1

sHSP

small heat shock protein

SMA

spinal muscular atrophy

SMALED

spinal muscular atrophy with lower limb predominance

Axonal transport is a highly complex process allowing movement of molecules and organelles within neurons over tremendous distances towards presynaptic nerve termini (anterograde transport), and transferring material from the periphery back to the neuronal soma (retrograde transport). This intracellular trafficking is crucial for sustaining proper neuronal functioning as well as clearing misfolded proteins or damaged organelles to avoid accumulation of harmful aggregates. Neuronal homeostasis depends on an efficient axonal transport allowing neuronal growth (axonal outgrowth), repair and regeneration upon injury, endocytosis and exocytosis of large and small molecules. The complex morphology and length (which can reach over one meter) of neurons make them particularly vulnerable to changes in axonal trafficking. Transport of a molecule or organelle (cargo) within the axon implies its recognition and binding to motor proteins, followed by an ATP-dependent movement of the motors along the cytoskeleton, direction of the cargo to the correct destination and release of the cargo upon reaching its destination. Axonal transport is regulated at all of these stages in a number of different ways. Impaired axonal transport can result in an altered neuronal homeostasis, aggregation of cargoes, and ultimately a dying-back degeneration of neurons.

Here we review the cytoskeletal components, molecular motors and adaptor proteins involved in axonal transport mechanisms and their implication in neuronal functioning. Furthermore, we discuss how genetic mutations, associated with inherited peripheral neuropathies (IPN), impact on axonal transport. Finally, we focus on some of the underlying mechanisms of axonal transport disturbances causing neurodegeneration and highlight future therapeutic prospects.

CYTOSKELETAL COMPONENTS OF AXONAL TRANSPORT

Neurons become post-mitotic cells in early development and need to remain functional for a lifetime, requiring a solid structural cytoskeleton. The neuronal cytoskeleton consists of microtubules (MTs), intermediate filaments and actin filaments (AFs), each having their own intracellular distribution (Fig. 1).

Fig.1

Schematic overview of the peripheral nerve and the mechanisms directly involved in typical and pathogenic axonal transport in anterograde and retrograde direction. (A) Transport of mitochondria occurs along microtubules by dynein (purple) and kinesin (blue) motors. Mitochondria undergo a coordinated balance between mitochondrial fission and fusion. Mitochondria-ER tethering and mitochondrial Ca²⁺ flux are maintained to sustain proper axonal transport. (B) Formation of autophagosomes and endosomes with subsequent fusion with lysosomes. Anterograde transport of lysosomes and retrograde transport of endosomes and autophagosomes occurs along microtubules by kinesin and dynein motors, respectively. (C) mRNA are bound by RNA-binding proteins (RBPs) and heterogeneous nuclear ribonuclear proteins (hnRNPs), and subsequently formed into ribonucleotide particles (RNPs) capable of being transported by kinesin and dynein motors along the microtubules. Increased aggregation of RBPs causes formation of stress granules. (D) The neuronal cytoskeleton provides the tracks on which the molecular motors move to facilitate axonal transport. The dynein and kinesin motors move along the microtubules, whereas the myosin motors move along actin filaments. Both actin filaments and microtubules undergo dynamic polymerization and depolymerization mediated by adaptor proteins. (E) Synaptic vesicles are transported in anterograde direction by binding to kinesin motors, which move along microtubules. Fusion of synaptic vesicles at the nerve terminal is mediated by v-SNARE (purple) and t-SNARE (green) complexes. Here the t-SNARE complex is depicted as Syntaxin (light green) and SNAP25 (dark green).

The MTs originate from the centrosome and are the major component of the neuronal cytoskeleton. MTs consist of α- and β-tubulin polymers that form polarized tubular structures, which are subjected to a dynamic process of polymerization and depolymerization [1]. Parallel MTs form unipolar arrays with the ‘minus-ends’ orientated towards the soma and the ‘plus-ends’ towards the axon, directing motor proteins in antero- or retrograde direction respectively. The tubulin polymers are involved in spatial organization and cell shape maintenance, as well as in many features of cytoplasmic structure, including organization of specific signaling pathways [1]. Microtubule-associated proteins, motor proteins, post-translational tubulin modifications and plus-end tracking proteins regulate MT dynamics.

The major intermediate filaments of neurons are neurofilaments (NFs), which control axon diameter and thereby axonal conductance [2]. Mature NFs contain neurofilament light (NEFL), medium (NEFM) and heavy (NEFH) chains. However, NFs in the peripheral nervous system can also contain peripherin. NF monomers share a common structural organization: a central α-helical rod domain, flanked by a N-terminal head, and a C-terminal tail domain [2]. In addition to their function in maintenance of the axonal architecture, NFs are fundamental to maintain Schwann cell-axon interactions, NF complex assembly and axonal transport [3].

The AFs consist of polymers of globular actin units and have a growing end (plus-end) where actin monomers are attached, while monomers are dissociated at the minus end. AF dynamics sustain a balance between polymerization and depolymerization and provide a structured cytoskeletal network for cell support, shape and migration. Within the cell, the function of AFs is regulated by a range of actin-binding proteins such as profilins and formins.

Disturbance or alteration of cytoskeleton stability and dynamics occurs in various neurodegenerative diseases, including IPN, and can be caused by mutations in the cytoskeletal proteins themselves, in their adaptor proteins or in other regulators. Mutations in INF2, encoding formin, cause autosomal dominant focal and segmental glomerulosclerosis (FSGS) as well as dominant intermediate Charcot-Marie-Tooth (CMT) type E [4, 5].

Mutations in tubulin genes are commonly referred to as tubulinopathies, such as mutations in β-tubulin known to cause polymicrogyria, congenital fibrosis of extraocular muscle (CFEOM) or malformation of cortical development. Mutations in TUBB3 are not only associated with CFEOM, e.g. the D417N mutation can also cause a polyneuropathy by reducing kinesin localization to microtubule plus-ends and affecting the axonal transport. TUBB3 mutations can lead to microtubule overstabilization and increased depolymerization [6].

Mutations in NEFL were identified in several subtypes of axonal CMT (CMT2) neuropathies, with dominant mutations causing their aggregation and recessive mutations resulting in a loss-of-function [7 –9]. NEFL mutations can target different domains in the protein affecting the NF assembly, e.g. the Q333P mutation leads to destabilization of coiled–coil interactions resulting in reduced self-interaction and dimerization of NFs [10]. A mutation located in the NEFL head domain, P8R, affects phosphorylation and destabilization of the NF complex resulting in NF aggregation [11]. Furthermore, phosphorylation of the NEFL head domain not only regulates NF assembly and disassembly, but also its transport in the axon [11, 12]. Dominant mutations in NEFH can also cause CMT2 [7, 13]. All reported NEFH mutations are frameshifts leading to the translation of additional 3’UTR regions encoding a cryptic amyloidogenic element. This leads to toxic protein aggregation similar to mutational mechanisms in NEFL and FUS, which when recognized by the autophagic pathway in motor neurons, trigger caspase 3 activation resulting in apoptosis [7, 13].

Several small heat shock proteins (sHSP), HSPB1 and HSPB8 are associated with cytoskeletal abnormalities and neuromuscular dysfunction [14 –16]. Dominant mutations in HSPB1 are associated with CMT type 2F and distal hereditary motor neuropathy (dHMN) [16]. The S135F mutation results into a higher affinity of HSPB1 for α-tubulin and overstabilization of MTs, and several HSPB1 mutants reduce α-tubulin acetylation, affecting axonal transport [16]. HSPB1 mutations can also affect the assembly and transport of NFs [17]. Moreover, spinal motor neurons, differentiated from patient-derived induced pluripotent stem cells (iPSC), carrying HSPB1 mutations showed a significant reduction of the mitochondrial mobility along the axons [18].

MOLECULAR MOTORS AND ADAPTORS IN AXONAL TRANSPORT

Neurons depend on efficient transport of cargoes, which is enforced by several different classes of molecular motor proteins (Fig. 1) [19]. Microtubule based molecular motors are grouped into the kinesins (for anterograde transport) and cytoplasmic dyneins (for retrograde transport) [19]. The kinesin family consists of 45 genes classified into 15 subfamilies of which the kinesin-1, kinesin-2 and kinesin-3 families contribute to axonal transport [20]. Kinesin motor domains homo- and heterodimerize, and bind additional kinesin light chains forming the kinesin motor complex [20]. The dynein family is much smaller, and the main component -dynein heavy chain- is encoded by only one gene (DYNC1H1). Dynein functions as a complex consisting of two dynein heavy chains that dimerize at their tail domains. Additional dynein intermediate chains and dynein light chains will bind to these tails by forming a cargo-binding domain [20].

Many axonal cargoes have multiple motor types that can bind simultaneously, even cargoes that move steadily in either antero- or retrograde direction [21, 22]. This shows that molecular motors do not function independently from each other when pulling an organelle in opposite directions. Rather they are co-dependent, as shown by impairment of minus-end motors resulting in severe suppression of plus-end motility [22]. This simultaneous application of opposite forces does not necessarily result in stalled cargoes. Instead, it likely provides additional support for transport coordination to overcome mechanical obstacles on MT tracks [21].

The third type of molecular motors are myosins, specifically involved in the transport over actin filaments [19]. Mammals have 40 myosin genes [19]. The most essential differences are found in the C-terminal globular tail domain, which recognizes various cargoes through direct interactions or via adaptor proteins [19]. For instance, Myosin Va is able to interact with kinesin heavy chains, with MTs through its tail domain and with NEFL through its head motor domain. Doing so, Myosin Va plays a crucial role in coupling microtubule- and actin-based transport mechanisms and regulates distribution of cargoes across the cytoskeletal network [23, 24].

Post-translational modification (PTM) of cytoskeletal components includes polyglutamylation, polyglycylation, detyrosination, acetylation, phosphorylation and palmitoylation [1]. For MTs, these preferentially occur on tubulin subunits already incorporated into microtubules. Molecular motors can recognize different PTMs as signature for their recruitment; e.g. polyglutamylation recruits the kinesin-3 family member KIF1A motor, whereas tyrosination recruits kinesin-1 family member KIF5 motor proteins [25, 26]. Additionally, protein kinases regulate axonal transport through direct phosphorylation of motors, adapters and cargoes, or protein kinases can phosphorylate several factors involved in the regulation of microtubule stability [27].

Dynactin is the best known adaptor protein of the cytoplasmic dynein complex. Dynactin-dynein interaction expands the range of cargoes that dynein can move, and increases the dynein motor processivity. Independently of cytoplasmic dynein, dynactin can anchor microtubules at the centrosome. The bicaudal D homologue (BICD) proteins represent another group of activating adaptors. In mammals, there are two BICD proteins, BICD1 and BICD2, as well as two related proteins, BICDR1 and BICDR2. The BICD functioning is diverse; BICD2 mostly functions in a complex of one dynein and one dynactin, whereas BICDR1 can recruit two dynein dimers to a single dynactin, further enhancing the force and velocity of the motor complex [28 –30]. BICD2 also interacts with the dynein heavy chain and dynactin, enhancing the affinity of the dynein–dynactin interaction. Another class of microtubule adaptors are the Hook proteins, which are involved in motor-microtubule interactions as well as in cargo binding [31]. However, mutations in Hook proteins have so far not been associated with IPN or other neurodegenerative diseases.

Where the cytoskeleton functions as the tracks, the molecular motors are equally important for proper axonal transport. Disturbances in these motor complexes and their adaptors are a recognized causal mechanism for IPN and related diseases. Dominant mutations in dynein heavy chain (DYNC1H1) are linked to several distinct neurodegenerative phenotypes, e.g. CMT type 2O, spinal muscular atrophy with lower limb predominance (SMALED), hereditary spastic paraplegia (HSP) and intellectual disability with neuronal migration defects (MRD13) [reviewed in [32]].

Several dynein adaptors and modulators are associated with neurological disease; dynactin (DCTN1), Huntingtin (HTT), LIS1, BICD1 and BICD2. Mutations in DCTN1 can cause Perry Syndrome (PS) and dHMN, and susceptibility to develop amyotrophic lateral sclerosis (ALS) [33 –35]. DCTN1 mutations found in PS patients affect amino acid residues within or immediately adjacent to the p150^glued CAP-Gly GKNDG motif of dynactin. A G59S variant in DCTN1 causing HMN occurs centrally in the p150^glued CAP-Gly domain. However, both PS and HMN associated variants induce a modest decrease in MT binding. Cells transfected with DCTN1 mutants have a dramatic redistribution of dynactin and more p150^Glued aggregates [34, 35]. It remains unclear why mutations localized in different part of the DCTN1 sequence manifest such different disease phenotypes [34, 35].

BICD2 mutations cause a spectrum of phenotypes including SMALED, HSP, and distal myopathy [36 –40]. BICD2 joins dynein and dynactin in a motor protein complex capable of processive movement. Modeling of SMALED-BICD2 mutations in cells showed an increased retrograde transport, suggesting that an imbalance of anterograde and retrograde dynein motor complex motility could be the underlying pathomechanism for these mutations [41, 42]. In addition, BICD2 has been implicated in Golgi functioning as the BICD2 coiled-coil (CC) structure allows interaction with the small GTPase RAB6A located at the Golgi apparatus and fragmentation of the Golgi apparatus has been shown in BICD2-patient fibroblasts [43].

Mutations in the other major group of molecular motors (kinesins) can also give rise to a spectrum of neurodegenerative diseases. Variants in KIF1A have been associated with HSP, hereditary sensory and autonomic neuropathy (HSAN) and complex phenotypes (combining HSP, HSAN and ataxia) [44, 45]. KIF5A variants can cause CMT2, HSP and ALS [46 –49]. Genotype/phenotype correlations revealed that the site of KIF5A mutations determine the clinical phenotype. Mutations in the kinesin motor or neck domain cause CMT2 and HSP. Mutations affecting splicing (exon 27) at the C-terminus cause atypical ALS with an earlier onset and a slower disease progression [46 –49]. KIF5A mutations in HSP alter processivity and directionality of kinesin by changing order–disorder transition of the neck linker, or affecting ATPase activity/microtubule gliding. Both mechanisms can alter MT-dependent transport and net anterograde transport of cargoes [50 –52]. KIF1Bβ mutations were first implicated in CMT2A in 2001, but as only few patients were identified, the association has been controversial [37 , 54]. Functional evidence for KIF1Bβ mutations in neuropathy phenotypes suggests that different mechanisms may be at play. The initially reported Q98L mutation resides in the conserved ATP-binding site and significantly reduced ATPase activity and perinuclear accumulation of the mutant KIF1Bβ protein. Only recently a novel CMT2-associated Y1087C mutation in KIF1Bβ was identified. It specifically impairs KIF1Bβ binding capacity and transport of Insulin-like growth factor 1 receptor (IGF1R) down the axons affecting IGF-I/IGF1R signaling, which is essential for neuronal survival and axonal development [54].

CARGOES IN AXONAL TRANSPORT

Synaptic vesicles, NFs, and cytosolic proteins are cargoes transported in anterograde fashion. Signaling endosomes, autophagosomes and proteins involved in injury signaling are transported towards the cell body. Lastly, mitochondria, a variety of endosomes, lysosomes and mRNA are transported in a bi-directional manner [31]. Depending on the cargo and its destination, specific regulatory proteins and motor complexes are recruited (Fig. 1). While cargoes make use of different transport pathways with their specific components, there is clear interdependence between these pathways and many of them converge [55]. In addition to the cargoes discussed below, axons also provide possibilities to transport viruses [reviewed in [56]].

mRNA and cytosolic proteins

Axons constitute the connections between neurons and their targets allowing them to communicate and respond to environmental stimuli. A large and diverse pool of cytosolic proteins is transported slowly from the soma towards the axon [57]. With a couple of examples studied so far, current models suggest that these cytosolic proteins form spontaneously aggregated complexes that undergo ‘dynamic recruitment’ to allow short bursts of anterograde transport by hitching a ride on passing vesicles [58, 59]. However, the fast-acting mechanisms of the axon, required to promptly respond to external stimuli, could not depend solely on slow transport of proteins from the soma. As such processes in the distal axons rely on localized protein synthesis to spatially and temporally regulate protein content by localized mRNA translation. In addition, local translation allows for differential PTM variants of translated proteins according local requirements.

The mRNAs subjected to local translation bind to RNA-binding proteins (RBPs) before undergoing active transport by molecular motors. While RBPs are reported to bind to both anterograde and retrograde motor complexes, it is unclear whether these RBPs have bound mRNAs, which would allow for differentiation between the two supposed functions: (I) relocation of mRNAs within the axon or (II) delivery of RBPs to the cell body for reuse [60]. Binding of mRNAs to RBPs regulates PTMs and stress responses ensuring mRNA stability, translation efficiency, and localization to cytoplasmic granules [61]. mRNA association with RBPs also guarantees the formation of transport-competent ribonucleoprotein particles (RNPs), which are protein complexes commonly referred to as ‘RNA transport granules’. Specificity of RBP binding to mRNAs is dictated by sequences within the 5^′ and 3^′ UTR of mRNAs, which are recognized by RBPs. Although more rarely, RBP regulatory sequences can also be present in protein coding regions. RBPs can cooperate or compete for a regulatory outcome at more than one of these sites per mRNA and specific RBP binding sequences are often present in many different mRNAs. Interestingly, mRNAs encoding proteins with complementary functions were shown to bind similar sets of RBPs leading to the concept of ‘RNA regulons’, which could be involved in subcellular compartment specific transport and translation.

Mutation in genes encoding RBPs are linked with several neuromuscular diseases such as ALS, SMA, multisystem proteinopathy (MSP) and frontotemporal lobar degeneration (FTLD). Well-known examples are TDP-43 mutations causative for ALS and FTLD. TDP-43 mutations affect its subcellular localization causing accumulation of RNP granules in the cell soma and proximal axons, they show disturbance in RNP complex transport and alter the axonal content of both mRNAs and miRNAs [62]. A major subtype of RBPs are the heterogeneous nuclear ribonuclear proteins (hnRNPs), wherein mutations are responsible for a number of cases of familial ALS and FTD [63]. Purice et al., (2018) reviewed the subset of disease-causing RBPs that are hnRNPs, namely TDP-43, FUS, hnRNPA1, hnRNPA2B1, matrin-3, and TIA1 [63]. While hnRNPs have an intrinsic tendency to aggregate, a common causal mechanism for these genes is the presence of mutations in the low complexity prion-like domain, which exacerbate the propensity to form self-seeding fibrils, resulting in accumulation of persistent stress granules [64, 65].

Mitochondrial transport

Neurons require functional mitochondria that need to travel long distances to provide support, such as adequate ATP-production, at specific sites of the neuron, like synaptic termini. Mutations affecting the balance between either division (known as fission), or collision and fusion of mitochondria, can cause alterations in the mitochondrial morphology. Subsequently, a more fragmented or dense mitochondrial network impacts axonal transport. The most prominent axonal CMT subtype (CMT2A) consists of mutations in MFN2 affecting mitochondrial transport [66]. Together with the MFN1 and OPA1, MFN2 plays a major role in the mitochondrial fusion process. Mitochondrial fusion is a unique process involving two membranes, i.e. the mitochondrial outer membrane (MOM) and mitochondrial inner membrane (MIM), that requires rearrangement in a coordinated manner in order to maintain the organelle’s integrity [67]. Mitochondrial fission is mediated by the recruitment of the DNM1L/Drp1 on the MOM [68, 69]. This happens through the interaction with the mitochondrial fission factor (Mff), mitochondrial dynamics protein Mid49 and Mid51, and in a minor interaction with mitochondrial fission 1 (Fis1) [70 –72]. Mutations in GDAP1, causative for several types of CMT, alter the regulation of mitochondrial fission activity dependent on the fission factors Drp1 and Fis1. Interestingly, the effect of GDAP1 mutations is dependent on the mode of inheritance. Recessive mutations cause a reduction in the fission activity, whereas the dominant inherited mutations hamper mitochondrial fusion events [73]. Mitochondrial function and localization can be indirectly affected by mutations in the Miro/Milton complex, which mediate MT interaction, and by mutations in NEFL, which in turn alter the mitochondrial distribution [74]. Interestingly, CMT associated mutations in small heat shock proteins HSPB1 and HSPB8 indirectly affect mitochondrial function transport by altering cytoskeletal properties. Furthermore, mitochondrial functioning in neurons can also be affected by mutations causing loss of contacts between the mitochondria and the endoplasmic reticulum (ER) or ER network stability. As a consequence, the ER is hampered to initiate mitochondria to promote fission, e.g. mutations in VAPB induce the formation of abnormal ER-derived inclusions [75]. Mutations in REEP1, associated with HSP and dHMN, were shown to disrupt ER network and promote ER fragmentation [76]. Whereas, decrease of VAPB MOM protein, causes a perturbation of the uptake of Ca²⁺ by mitochondria, which is required to maintain an intracellular homeostasis as well as mitochondrial transport. Ca²⁺ is an important factor for various other functions such as cell signaling as well as regulating mitochondrial function and structure [77]. Therefore, mutations in Ca²⁺-channels such VDAC or TRPV4 lead to mitochondrial dysfunction in IPN [77].

Membrane-bound dynamin like GTPases known as atlastins (ATLs) mediate the formation of ER-mitochondria contact sites essential for calcium communication [78]. Mutations in the human isoforms ATL1 and ATL3 are associated with HSP and HSAN. Transmission electron microscopy studies reported higher ER-mitochondria contact sites upon expression of mutant ATL3, resulting in increased Ca²⁺ uptake into the mitochondria [78]. Moreover, aberrant calcium signaling affects mitochondrial trafficking through the Rho GTPases Miro1 and Miro2. Atlastin mutations demonstrate a reduced mobility of mitochondria in the cytoplasm together with altered mitochondrial localization, where the mitochondria are retained within the soma rather than distributed to the neuronal processes [78]. Furthermore, mutations in SPTLC1 and SPTLC2 produce atypical sphingolipids observed in HSAN type I. Alecu et al., (2017) demonstrated that these toxic deoxysphingolipids could localize into mitochondria disrupting the mitochondrial integrity [79].

Mutations in tRNA synthetases (e.g. GARS, HARS or KARS) also affect mitochondrial function. Mitochondrial dysfunction could contribute to neuromuscular junctions (NMJ) degeneration. E.g. a CMT2 type D mouse model expressing mutant GARS mutant displayed affected NMJs as well as muscle atrophy prior to synaptic degeneration [80].

Vesicular transport

Vesicular transport in axons occurs in different forms depending on the origin of intracellular components. Late endosomes and autophagosomes are subjected to retrograde transport to reach the lysosome, which mainly have a perinuclear localization. In contrast, synaptic vesicle precursors are produced in the soma and transported in anterograde direction towards the axon. These, mature synaptic vesicles are essential for proper neuronal growth, function and survival as they contain neurotransmitters, contribute to synapse formation and location, as well as help to sustain a balance between exo- and endocytosis [81, 82]. The transport of all vesicle types occurs through conserved mechanisms. Defects in the formation of endosomes, autophagosomes, lysosomes and synaptic vesicles, as well as impairments in their antero- and retrograde trafficking, may cause axonal degeneration [83, 84].

Rab GTPases regulate vesicular trafficking at different levels from vesicle formation, vesicle movement along actin and tubulin networks, to membrane fusion. Members of this protein family are involved in autophagy, lysosome and synaptic vesicle transport. Relevant for IPN and related neurodegenerative diseases is the small GTPase Rab5, which mediates transport and fusion of early endosomes, inducing neurite outgrowth and dendritic branching [85]. ALS2 encodes a Guanine Nucleotide Exchange Factor for Rab5 and is involved in endosomal dynamics. Mutations in ALS2, associated with the onset of HSP, ALS and primary lateral sclerosis, produce truncated forms of alsin causing alterations in Rab5 and Rab7 signaling and Rab5-to-Rab7 conversion [86]. Mutations in NIPA1 or SPG11 similarly affect endosomal trafficking. A list of other HSP genes, divided by functionality has been reviewed in [87]. In addition to Rab5, Rab7 plays a crucial role in trafficking of late endosomes to lysosomes and is responsible for bidirectional transport of autophagosomes as well as for autophagosome-lysosome fusion in all cells. Rab7 binds to effector proteins like FYCO1, responsible for plus-end, or to ORP1L and RILP, responsible for minus-end-directed transport [88].

Other classes of GTPases are involved in membrane trafficking and in vesicle formation in the endo-lysosome pathway. Dynamin2 (DNM2) polymers work by wrapping the neck of budding membranes, promoting membrane fission. Mutations in DNM2 are associated with autosomal dominant centronuclear myopathy (ADCNM) and intermediate CMT [89]. DNM2 contains actin binding sites suggesting that it may regulate actin dynamics during membrane tabulation [90].

An ensemble of kinesin-1, SKIP, Arl8, and subunits of the BLOC-one-related complex (BORC) direct anterograde transport of lysosomes into the axon. These eight BORC subunits consist of BLOS1, BLOS2, Snapin, KXD1, LOH12CR1 (myrlysin), C17orf59 (lyspersin), C10orf32 (diaskedin) and LOC729991 (MEF2BNB). Interference of the BORC function decreases lysosome transport in the axon, and its function is required for maintenance of axonal growth cone dynamics and autophagosome clearance [91].

The different functions of Rab GTPases correlated with the pathological conditions was reviewed by [86]. Nian et al., (2019) demonstrated that primary neurons from Rab^–/– mice showed a reduction in lysosomal trafficking together with the accumulation of autophagosomes, suggesting an altered autophagic vesicle transport [92]. Rab3 and Rab27 are important in synaptic vesicle exocytosis, while Rab5 is a key regulator of synaptic vesicles retrieval/endocytosis. Warburg Micro Syndrome (WARBM) and CMT2 type B neuropathy are associated with mutations in RAB3GAP1 and RAB7 respectively. Zebrafish Rohon-Beard spinal sensory neurons expressing Rab7 mutations reported defects in neurite outgrowth and branching, and a marked decrease in the speed of K157N Rab7 containing vesicles, underscoring the importance of Rab7 in endosome transport [93]. Furthermore, mutations in CHMP2B cause a reduced Rab7-endosome recruitment and are linked to a FTD phenotype with partial overlap with ALS.

Another aspect is the fusion of synaptic vesicles at the nerve terminals. The neuronal v-SNAREs, vesicle Soluble N-ethylmaleimide-sensitive factor Attachment protein Receptors, are essential for these fusion processes. Mutations in VAMP2 encoding one of the v-SNAREs, cause aberrant synaptic vesicle morphology and vesicle endocytosis often characterized by axial hypotonia [94]. Other mutations such as those in UNC13A can affect synaptic regulators and are known to be involved in neurodevelopmental disorders often in combination with involuntary movements [95].

RESEARCH AND THERAPEUTICS PROSPECTS

Axonal transport deficits are one of the most common and recurrent pathomechanisms in IPN and other neurodegenerative diseases (Table 1 and Fig. 2). For this reason, axonal transport studies are crucial, not only to understand its physiological role, but also to determine the effect of mutations that impair the network of axonal transport process and its regulation. Of note is that similar disturbances of axonal transport, or one of its key components, are also at play in other far more prevalent acquired disorders of the PNS [96]. Axonal transport in human peripheral nerves is difficult to investigate, as the tissue of interest is inaccessible. Therefore, animal models have been used especially for real-time monitoring of cargoes trafficking and axonal transport defects in order to develop novel therapeutic strategies. The wings of the fruit fly Drosophila melanogaster offer an in vivo model to study axonal cargo dynamics [97] and specific targeting of fluorescently-labelled organelles in the nematode Caenorhabditis elegans allow studying of axonal growth and synaptogenesis [98]. Furthermore, in transgenic MFN2 zebrafish (Danio rerio) it is possible to genetically label mitochondria in motor neurons [99]. Other platforms for in vivo axonal transport studies compared to ex vivo tools have been reviewed by [100].

Fig.2

Overview of the subcellular components and genes (illustrated by gene symbols) involved in axonal transport mechanisms and in which defects are associated with IPN and related neurodegenerative diseases.

Table 1

Overview of genes causative for IPN-related disorders involved in axonal transport

	Gene symbol	Protein name	Function	Mutational effects	Mode of inheritance	Disease associations	Refs
Cytoskeleton	TUBB3	Tubulin Beta 3 Class III	Structural component of neurofilaments	Impairment of tubulin heterodimer formation.	AD	CMT2	[1, 6]
						Congenital fibrosis of extraocular muscles 3
	NEFL	Neurofilament Light	Structural component of neurofilaments	Reduced self-interaction and neurofilament dimerization.	AD/AR	CMT1F	[2 , 7–12]
				Neurofilament aggregation.		CMT2E
						CMT2B5
						Nemaline rod myopathy
	NEFH	Neurofilament Heavy	Structural component of neurofilaments	Translation of cryptic amyloidogenic element causing protein aggregation.	AD	CMT2CC	[2 , 13]
	HSPB1	Heat Shock Protein Family B (Small) Member 1	Molecular chaperone for reducing protein aggregation and misfolding	Microtubule overstabilitization.	AD	CMT2F	[16–18 , 114]
				Impaired mitochondrial transport.		dHMN2B
	HSPB8	Heat Shock Protein Family B (Small) Member 8	Molecular chaperone for reducing protein aggregation and misfolding	Cytoskeletal destabilization.	AD	CMT2L	[14, 15]
				Reduced mitochondrial membrane potential.		dHMN2
				Impairment of aggregate degradation via autophagy.
	INF2	Inverted formin-2	Stimulating actin filament growth and mitochondrial fission	Perturbation of actin filament dynamics and cytoskeletal networks.	AD	Focal and segmental glomerulosclerosis	[4, 5]
						CMT intermediate type E
Motor complexes and adaptors
	DYNC1H1	Dynein Cytoplasmic 1 Heavy Chain 1	Dynein subunit for retrograde axonal transport	Inhibition of microtubule gliding.	AD	CMT2O	[31, 32]
				Compromised dynein processive movement activation.		SMALED
						MRD13
	DCTN1	Dynactin subunit 1	Dynein adaptor protein for retrograde axonal transport	Altered localization of dynactin.	AD	HMN7B	[31 , 33–35]
				Reduced dynactin-microtubule binding.		ALS susceptibility
	BICD2	Protein bicaudal D homolog 2	Dynein adaptor protein for retrograde axonal transport	Imbalance in anterograde and retrograde dynein motor complex motility.	AD	SMALED2A	[28–31 , 36–43]
						SMALED2B
						Spastic paraparesis
						Distal myopathy
	KIF1A	Kinesin-like protein KIF1A	Motor protein for anterograde axonal transport	Impaired microtubule binding.	AD/AR	HSN2C	[19 , 45]
				Impaired movement along microtubules.		CMT2 with acrodystrophy
						SPG30
	KIF1Bβ	Kinesin family member 1Bbeta isoform III	Motor protein for anterograde axonal transport	Reduced ATPase activity and perinuclear localization.	AD	CMT2A1	[19 , 54]
				Impairment of binding capacity and transport of IGF1R.
	KIF5A	Kinesin heavy chain isoform 5A	Motor protein for anterograde axonal transport	Altered processivity and directionality of kinesin dependent transport.	AD	CMT2 with pyramidal signs	[19 , 52]
						SPG10
						ALS25
mRNA and proteins
	HNRNPA1	Heterogeneous nuclear ribonucleoprotein A1	Packages mRNA into RNP particles	Increased fibrillization and self-aggregation.	AD	Myopathy (IBM) with Paget disease of Bone without Dementia (IBMPFD3)	[61 , 63–65]
				Increased formation of cytoplasmic stress granules.		ALS20
						HMN (unpublished)
Mitochondria
	ATL1	Atlastin-1	Formation of ER-mitochondria contact sites	Impairment of GTPase activity and dimer formation.	AD	HSN1D	[78]
				Reduced ER-mitochondia fusion effect.		SPG3A
	ATL3	Atlastin-3	Formation of ER-mitochondria contact sites	Higher ER-mitochondria contact sites.	AD	HSN1F	[78]
				Increased Ca²⁺ uptake into the mitochondria.
	GDAP1	Ganglioside-induced differentiation-associated protein 1	Mediates mitochondrial fission	Impairment of mitochondrial fusion (AD).	AD/AR	CMT2K	[67 , 74]
				Reduction in mitochondrial fission activity (AR).		CMT4A
	MFN2	Mitofusin-2	Mediates mitochondrial clustering and fusion	Aberrant mitochondrial morphology and altered mitochondria-ER tethering.	AD/AR	CMT2A2A	[67 , 115]
						CMT2A2B
	REEP1	Receptor Expression Enhancing Protein 1	Stabilization of ER tubules	Destabilization of ER tubules.	AD/AR	HSP	[76]
				ER fragmentation.		dHMN
				Accumulation of synaptic vesicle proteins.		Congenital axonal neuropathy with diaphragm palsy
	SPTLC1	Serine palmitoyltransferase 1	Production of sphingolipids	Production of toxic deoxysphingolipids that disrupt mitochondrial integrity.	AD	HSAN1A	[79]
	SPTLC2	Serine palmitoyltransferase 2	Production of sphingolipids	Production of toxic deoxysphingolipids that disrupt mitochondrial integrity.	AD	HSAN1C	[79]
	TRPV4	Transient receptor potential cation channel subfamily V member 4	Non-selective calcium permeant cation channel	Abnormal TRPV4-regulated Ca²⁺ influx.	AD	HMN8	[77 , 128–130]
				Higher basal intracellular Ca²⁺ levels.		SMA
						SMALED
						CMT2C
	VAPB	Vesicle-associated membrane protein-associated protein B/C	Formation of membrane contact sites between ER and other organelles	Formation of intracellular aggregates not associated with membranes.	AD	Proximal SMA	[75]
Vesicular transport
	DNM2	Dynamin-2	Production of microtubule bundles	Increased GTPase activity and oligomerization.	AD	CMT intermediate type B	[89, 90]
				Impaired autophagic degradation.		CMT2M
				Impaired DNM2 lipid binding and GTPase activity.		Centronuclear myopathy
						Lethal congenital contracture syndrome 5
	GAN	Gigoxonin	Involved in crosstalk of the cytoskeletal architecture and E3 ligase	Inhibition of autophagosome synthesis and altered fusion to the lysosome.	AR	GAN	[131, 132]
	RAB7	Ras-related protein Rab-7	Regulation of endo-lysosomal trafficking	Increased nucleotide exchange rates.	CMT2B	[93]
				Reduced hydrolysis of GTP.
	SPG11	Spatacsin	Endosomal trafficking	Alters Rab5 and Rab7 signaling.	AR	CMT2X	[87, 133]
				Alters Rab5-to-Rab7 conversion.		HSP
						ALS5

AD - autosomal dominant; ALS - amyotrophic lateral sclerosis; AR - autosomal recessive; CMT - Charcot-Marie-Tooth; (d)HMN - (distal) hereditary motor neuropathy; GAN - giant axonal neuropathy; HSAN - hereditary sensory and autonomic neuropathy; HSN - hereditary sensory neuropathy; HSP - hereditary spastic paraplegia; MRD13 - mental retardation, autosomal dominant 13; SMA - spinal muscular atrophy; SMALED - spinal muscular atrophy with lower limb predominance; SPG - spastic paraplegia.

Anatomical, metabolic and physiological differences between small animal models and human complicate the translation of many therapies into clinical trials. However, iPSC technology offers the possibility to reprogram patient-derived cell lines into pluripotent cells before differentiation into a specific cell type relevant for the disease [101]. This provides new possibilities for IPN, allowing the generation of patient-derived motor and sensory neurons, potentially Schwann cells as well, without using peripheral nerve biopsies or post-mortem tissues. Nevertheless, further improvements are required to more closely model the in vivo situation to have a more representative environment and to take the necessary steps in optimization of co-cultures to study motor neuron-Schwann cell interactions.

Working with 2D-monolayers, microfluidic chambers provide the possibility to investigate retrograde and anterograde transport of organelles with live-cell imaging. Moreover, microfluidic chambers are a flexible tool to mimic the generation of neuromuscular junctions (NMJ), when motor neurons are co-cultured with iPSC-derived muscle cells. Nowadays, bioengineering and iPSC-research work together to develop functional 3D-models to properly reconstruct the immunological, biochemical and anatomical feature of specific organs. A 3D-printed heart-like structure was recently reported using personalized hydrogel, combined with patient-derived cells to print thick, vascularized and perfusable cardiac patches [102]. This suggest that we are not far from the generation of 3D models which could include iPSC-derived peripheral nerves, Schwann cells, blood vessels and myofibers, supported by 3D-scaffolds obtained with new biomaterials and methods of fabrication [103], being entirely supplied by a regulated oxygen transfer with flow pumps. The iPSC-derived models are therefore moving closer to a preclinical application, by providing a pre-screening platform for candidate drugs before testing them in vivo. However, animal models will still remain indispensable to investigate translational research in a whole organism [104, 105].

To date, treatment of IPN is limited to supportive measures that have a partial benefit in relieving the symptoms, such as neuropathic pain or gait impairment. However, no effective interventions exist targeting the causative pathway. Thanks to the understanding of genetic causes, together with the development of more appropriate cellular and animal models, axonal transport emerged as candidate drugable pathway to restore the neuronal function. This will apply to IPN, however many of these insights will also be relevant for the more common acquired diseases of the peripheral nervous system.

The first attempts in targeting axonal degeneration have come from the field of drug repurposing of small molecules. To reduce protein aggregation and improve axonal transport, chemical compounds target MTs through direct binding, PTMs, or chaperone upregulation, aiming to reach the right balance between highly stable and hyperdynamic MTs. In HSP patient-derived neurons, low doses of microtubule-targeting agents (MTA), such as taxol and vinblastine, increased the acetylated α-tubulin levels and restored peroxisome trafficking speeds and distance travelled, effectively improving the mutant phenotype [106]. Other MTA like Epothilone D improved microtubule density, axonal density and cognition in an AD mouse model [107]. However, these compounds, already approved as anticancer drugs, have been discontinued in clinical trials for neurodegenerative diseases due to neurotoxicity [108]. This suggests that new strategies are required to effectively intervene in the treatment of IPN.

Recent studies reported that histone deacetylase 6 (HDAC6) inhibitors such as ACY-1215 (Ricolinostat), currently in clinical trials for cancer, or ACY-1083 showing a higher selectivity towards HDAC6, are able to restore nerve function. They also affect microtubule dynamics by increasing the acetylation of α-tubulin, which re-established the transport function of MTs and offer neuroprotection [109]. Miro1 has been identified and shown to act directly as a novel target of HDAC6, in which Miro1 is deacetylated, at lysine 105, resulting in reduction of mitochondrial transport and outgrowth of the axonal cone. Therefore, HDAC6 inhibitors significantly correct anterograde transport of mitochondria, supporting the therapeutic use of this class of molecules for IPN [110].

Axonal transport involves specific kinase cascade activation, which is altered in neurodegenerative diseases [111]. In ALS, overactivation of p38 MAPKα causes excessive phosphorylation of molecular motors preventing their movement along MTs and p38 MAPKα phosphorylation of NF subunits altering their transport and inducing bundling, effectively inhibiting retrograde transport. The use of p38 MAPKα inhibitors reverses these transport deficits in SOD1-G93A motor neurons, representing a novel therapeutic strategy in ALS. This represents a promising perspective in the treatment of AD, as well as in IPN [112, 113]. Failure in the protein quality control system can lead to deficits in axonal trafficking and aggregate clearance, which can drastically affect neuronal function. Therefore, upregulation of chaperones could be beneficial. Dual treatment using celastrol and arimoclomol increases the expression of a set of sHSPs (HSPA6, HSPA1A, DNAJB1, HO-1, HSPB1) in differentiated SH-SY5Y neuronal cells [114]. Despite the ability of celastrol in reducing the percentage of neuronal inclusions in the transgenic SOD1 mouse model of ALS, or in neurons expressing aggregation prone NEFL mutants, celastrol shows a motor neuron specific effect, with no effect in sensory neurons, limiting its use in IPN with sensory involvement [10]. As drugs can act on different levels of interactions (e.g. level of target, pathway, processes), co-administration of pharmaceutical active molecules should be considered to target the complexity and diversity of affected molecular pathways in IPN [115].

Despite the benefit of small molecules to rescue axonal degeneration, the risk for side effects has limited the use of these treatment strategies. However, a new class of molecules active on microtubule, molecular motors and autophagy [116], as well as antisense oligonucleotides (ASO) or gene therapy directly targeting the affected genes, have emerged and offer promising tools to treat IPN with axonal transport defects. ASO therapies have recently been developed and approved in the treatment of patients with spinal muscular atrophy (SMA) [117]. Similarly, ASO therapies could be applied to reduce PMP22 transcription levels in the CMT1A duplication [118]. Further advancements have been made in the delivery of vector-based gene therapies using adeno-associated viruses (AAV) in the treatment of SMA, consisting of delivery of a functional copy of the human SMN1 gene into motor neuron cells [119, 120]. Likewise, intrathecal injection of lentiviral vectors for Schwann cell-targeted expression has been used to restore the nodal architecture in demyelinating neuropathies. In Sh3tc2^–/– mice, a genetic model of CMT4C, or in the mutant GJB1 mouse model for CMTX1, gene delivery respectively of hSH3TC2 or WT Cx32, resulted in amelioration of motor performances together with a reduced myelin pathology [120, 121]. These few examples show that understanding dysfunctional genes operating in the axon, together with optimization of gene delivery methods, including vectors and administration routes, opens promising prospects to treat IPN and related axonopathies.

Deficits in axonal transport are inevitably linked with progressive axonal degeneration, a common feature of IPN and other neurodegenerative diseases. SARM1 (Sterile Alpha and TIR Motif Containing 1) is a mediator of axonal degeneration and initiates cellular self-destruction. Deletion of SARM1 or expression of dominant negative SARM1 mutations, both impairing its activation, showed a reduction in axonal degeneration after axonal transection, the most rapid and aggressive trigger of axonal degeneration [122]. Furthermore, Turkiew et al., (2017) reported that Sarm1^–/– mice are resistant to distal axonal degeneration in a model of chemotherapy induced peripheral neuropathy and in high fat diet induced metabolic neuropathy [123]. AAV-mediated delivery of dominant negative SARM1 in mice induces long-lasting axon protection following nerve transection. This approach may provide a new strategy to slow axon loss in chronic neurodegenerative diseases [124].

For neuropathies with toxic aggregations, therapeutic strategies are currently in development making use of gene replacement or silencing in a cell-specific manner. Recently, for CMT2A caused by dominant heterozygous mutations in MFN2, a combined therapy was tested on iPCS-derived spinal motor neurons which include the simultaneous use of RNA interference to silence the mutant allele and insertion of a mutagenized MFN2 gene, resistant to shRNA activity, encoding for the native protein [125]. Next to therapeutic strategies, the identification of biomarkers for disease is relevant to diagnose and treat pre-symptomatic patients or to follow-up ongoing treatments. Currently, measuring NEFL levels in plasma is correlated with disease severity in multiple forms of CMT neuropathies [126], and also PFN2 and GAMT were identified as molecular determinants for CMT2 neuropathy, with a possible role of PFN2 in disease progression [127].

CONCLUSION

Axonal transport is a highly dynamic process involving the movement of different types of cargoes (mRNA, proteins, mitochondria, lysosomes and synaptic vesicles) that are essential to sustain healthy neuronal functions. In this review, we have highlighted how deficits in cargo transportation and related factors affect axonal transport. The identification of numerous genetic causes for IPN-related disorders provides important insights into the underlying mechanisms of axonal degeneration. This knowledge allows the design of targeted therapeutic approaches, some of which have taken up a lead role and moved into clinical trials. Despite this, not all components and mechanisms of axonal transport have been unraveled, novel research strategies have emerged and will move towards patient-derived model systems (e.g. iPSC derived neurons in 2D and 3D-cultures) and in vivo animal models. These models will create new platforms to study and test therapeutic strategies for axonal degeneration.

CONFLICT OF INTEREST

The authors have no conflict of interest to report.

Footnotes

ACKNOWLEDGMENTS

The research in our laboratories is supported by the University of Antwerp, Flanders Fund for Scientific Research (FWO-Vlaanderen), Medical Foundation Queen Elisabeth (GSKE), Association Belge contre les Maladies Neuromusculaires (ABMM), Association Française contre les Myopathies (AFM), Muscular Dystrophy Association (MDA) and the European Union’s Horizon 2020 project Solve-RD (Solving the unsolved Rare Diseases) under grant agreement N°779257. These granting organizations provided in part support to PhD students D.B., A.S. and J.V.L. JB is supported by a Senior Clinical Researcher mandate of the Research Fund - Flanders (FWO) under grant agreement number 1805016N.

References

Chakraborti

, Natarajan

, Curiel

, Janke

, Liu

. The emerging role of the tubulin code: From the tubulin molecule to neuronal function and disease, Cytoskeleton (Hoboken). 2016;73(10):521–50. doi: 10.1002/cm.21290

Grant

, Pant

. Neurofilament protein synthesis and phosphorylation, J Neurocytol. 2000;29(11-12):843–72. doi:

Lancaster

, Li

, Hanania

, Liem

, Scheideler

, Scherer

. Myelinated axons fail to develop properly in a genetically authentic mouse model of Charcot-Marie-Tooth disease type 2E, Exp Neurol. 2018;308:13–25. doi: 10.1016/j.expneurol.2018.06.010

Brown

, Schlondorff

, Becker

, Tsukaguchi

, Tonna

, Uscinski

, et al. Mutations in the formin gene INF2 cause focal segmental glomerulosclerosis, Nat Genet. 2010;42(1):72–6. doi: 10.1038/ng.505

Boyer

, Nevo

, Plaisier

, Funalot

, Gribouval

, Benoit

, et al. INF2 mutations in Charcot-Marie-Tooth disease with glomerulopathy, N Engl J Med. 2011;365(25):2377–88. doi: 10.1056/NEJMoa1109122

Hong

, Lee

, Park

, Choi

, Hyun

, Park

, et al. A family with axonal sensorimotor polyneuropathy with TUBB3 mutation, Mol Med Rep. 2015;11(4):2729–34. doi: 10.3892/mmr.2014.3047

Rebelo

, Abrams

, Cottenie

, Horga

, Gonzalez

, Bis

, et al. Cryptic amyloidogenic elements in the 3’ UTRs of neurofilament genes trigger axonal neuropathy, Am J Hum Genet. 2016;98(4):597–614. doi: 10.1016/j.ajhg.2016.02.022

Sainio

, Ylikallio

, Maenpaa

, Lahtela

, Mattila

, Auranen

, et al. Absence of NEFL in patient-specific neurons in early-onset Charcot-Marie-Tooth neuropathy, Neurol Genet. 2018;4(3):e244. doi: 10.1212/NXG.0000000000000244

Jordanova

, De Jonghe

, Boerkoel

, Takashima

, De Vriendt

, Ceuterick

, et al. Mutations in the neurofilament light chain gene (NEFL) cause early onset severe Charcot-Marie-Tooth disease, Brain. 2003;126(Pt 3):590–7. doi: 10.1093/brain/awg059

10.

Gentil

, Mushynski

, Durham

. Heterogeneity in the properties of NEFL mutants causing Charcot-Marie-Tooth disease results in differential effects on neurofilament assembly and susceptibility to intervention by the chaperone-inducer, celastrol, Int J Biochem Cell Biol. 2013;45(7):1499–508. doi: 10.1016/j.biocel.2013.04.009

11.

Brownlees

, Ackerley

, Grierson

, Jacobsen

, Shea

, Anderton

, et al. Charcot-Marie-Tooth disease neurofilament mutations disrupt neurofilament assembly and axonal transport, Hum Mol Genet. 2002;11(23):2837–44. doi: 10.1093/hmg/11.23.2837

12.

Yates

, Manser

, De Vos

, Shaw

, McLoughlin

, Miller

. Neurofilament subunit (NFL) head domain phosphorylation regulates axonal transport of neurofilaments, Eur J Cell Biol. 2009;88(4):193–202. doi: 10.1016/j.ejcb.2008.11.004

13.

Jacquier

, Delorme

, Belotti

, Juntas-Morales

, Sole

, Dubourg

, et al. Cryptic amyloidogenic elements in mutant NEFH causing Charcot-Marie-Tooth 2 trigger aggresome formation and neuronal death, Acta Neuropathol Commun. 2017;5(1):55. doi: 10.1186/s40478-017-0457-1

14.

Bouhy

, Juneja

, Katona

, Holmgren

, Asselbergh

, De Winter

, et al. A knock-in/knock-out mouse model of HSPB8-associated distal hereditary motor neuropathy and myopathy reveals toxic gain-of-function of mutant Hspb8, Acta Neuropathol. 2018. doi:

15.

Irobi

, Holmgren

, De Winter

, Asselbergh

, Gettemans

, Adriaensen

, et al. Mutant HSPB8 causes protein aggregates and a reduced mitochondrial membrane potential in dermal fibroblasts from distal hereditary motor neuropathy patients, Neuromuscul Disord. 2012. doi:

16.

d’Ydewalle

, Krishnan

, Chiheb

, Van Damme

, Irobi

, Kozikowski

, et al. HDAC6 inhibitors reverse axonal loss in a mouse model of mutant HSPB1-induced Charcot-Marie-Tooth disease, Nat Med. 2011;17(8):968–74. doi: 10.1038/nm.2396

17.

Ackerley

, James

, Kalli

, French

, Davies

, Talbot

. A mutation in the small heat-shock protein HSPB1 leading to distal hereditary motor neuronopathy disrupts neurofilament assembly and the axonal transport of specific cellular cargoes, Hum Mol Genet. 2006;15(2):347–54. doi: 10.1093/hmg/ddi452

18.

Kalmar

, Innes

, Wanisch

, Kolaszynska

, Pandraud

, Kelly

, et al. Mitochondrial deficits and abnormal mitochondrial retrograde axonal transport play a role in the pathogenesis of mutant Hsp27-induced Charcot Marie Tooth Disease, Hum Mol Genet. 2017;26(17):3313–26. doi: 10.1093/hmg/ddx216

19.

Xiao

, Hu

, Wei

, Tam

. Cytoskeleton molecular motors: Structures and their functions in neuron, Int J Biol Sci. 2016;12(9):1083–92. doi: 10.7150/ijbs.15633

20.

De Vos

, Hafezparast

. Neurobiology of axonal transport defects in motor neuron diseases: Opportunities for translational research?Neurobiol Dis. 2017;105:283–99. doi: 10.1016/j.nbd.2017.02.004

21.

Hancock

. Bidirectional cargo transport: Moving beyond tug of war, Nat Rev Mol Cell Biol. 2014;15(9):615–28. doi: 10.1038/nrm3853

22.

Gross

, Welte

, Block

, Wieschaus

. Coordination of opposite-polarity microtubule motors, J Cell Biol. 2002;156(4):715–24. doi: 10.1083/jcb.200109047

23.

Cao

, Chang

, Masters

, Mooseker

. Myosin-Va binds to and mechanochemically couples microtubules to actin filaments, Mol Biol Cell. 2004;15(1):151–61. doi: 10.1091/mbc.e03-07-0504

24.

Rao

, Mohan

, Kumar

, Yuan

, Montagna

, Campbell

, et al. The myosin Va head domain binds to the neurofilament-L rod and modulates endoplasmic reticulum (ER) content and distribution within axons, PLoS One. 2011;6(2):e17087. doi: 10.1371/journal.pone.0017087

25.

Ikegami

, Heier

, Taruishi

, Takagi

, Mukai

, Shimma

, et al. Loss of alpha-tubulin polyglutamylation in ROSA22 mice is associated with abnormal targeting of KIF1A and modulated synaptic function, Proc Natl Acad Sci U S A. 2007;104(9):3213–8. doi: 10.1073/pnas.0611547104

26.

Konishi

, Setou

. Tubulin tyrosination navigates the kinesin-1 motor domain to axons, Nat Neurosci. 2009;12(5):559–67. doi: 10.1038/nn.2314

27.

Gibbs

, Greensmith

, Schiavo

. Regulation of axonal transport by protein kinases, Trends Biochem Sci. 2015;40(10):597–610. doi: 10.1016/j.tibs.2015.08.003

28.

Urnavicius

, Lau

, Elshenawy

, Morales-Rios

, Motz

, Yildiz

, et al. Cryo-EM shows how dynactin recruits two dyneins for faster movement, Nature. 2018;554(7691):202–6. doi: 10.1038/nature25462

29.

Grotjahn

, Chowdhury

, Xu

, McKenney

, Schroer

, Lander

. Cryo-electron tomography reveals that dynactin recruits a team of dyneins for processive motility, Nat Struct Mol Biol. 2018;25(3):203–7. doi: 10.1038/s41594-018-0027-7

30.

Schlager

, Serra-Marques

, Grigoriev

, Gumy

, Esteves da Silva

, Wulf

, et al. Bicaudal d family adaptor proteins control the velocity of Dynein-based movements, Cell Rep. 2014;8(5):1248–56. doi: 10.1016/j.celre2014.07.052

31.

Olenick

, Holzbaur

ELF

. Dynein activators and adaptors at a glance, J Cell Sci. 2019;132(6). doi: 10.1242/jcs.227132

32.

Schiavo

, Greensmith

, Hafezparast

, Fisher

. Cytoplasmic dynein heavy chain: The servant of many masters. Trends Neurosci. 2013;36(11):641–51. 10.1016/j.tins.2013.08.001

33.

Farrer

, Hulihan

, Kachergus

, Dachsel

, Stoessl

, Grantier

, et al. DCTN1 mutations in Perry syndrome, Nat Genet. 2009;41(2):163–5. doi: 10.1038/ng.293

34.

Levy

, Sumner

, Caviston

, Tokito

, Ranganathan

, Ligon

, et al. A motor neuron disease-associated mutation in p150Glued perturbs dynactin function and induces protein aggregation, J Cell Biol. 2006;172(5):733–45. doi: 10.1083/jcb.200511068

35.

Puls

, Jonnakuty

, LaMonte

, Holzbaur

, Tokito

, Mann

, et al. Mutant dynactin in motor neuron disease, Nat Genet. 2003;33(4):455–6. doi: 10.1038/ng1123

36.

Neveling

, Martinez-Carrera

, Holker

, Heister

, Verrips

, Hosseini-Barkooie

, et al. Mutations in BICD2, which encodes a golgin and important motor adaptor, cause congenital autosomal-dominant spinal muscular atrophy, Am J Hum Genet. 2013;92(6):946–54. doi: 10.1016/j.ajhg.2013.04.011

37.

Drew

, Zhu

, Kidambi

, Ly

, Tey

, Brewer

, et al. Improved inherited peripheral neuropathy genetic diagnosis by whole-exome sequencing, Mol Genet Genomic Med. 2015;3(2):143–54. doi: 10.1002/mgg3.126

38.

Koboldt

, Kastury

, Waldrop

, Kelly

, Mosher

, McLaughlin

, et al. In-frame de novo mutation in BICD2 in two patients with muscular atrophy and arthrogryposis, Cold Spring Harb Mol Case Stud. 2018;4(5). doi: 10.1101/mcs.a003160

39.

Kropatsch

, Schmidt

, Buttkereit

, Epplen

, Hoffjan

. BICD2 mutational analysis in hereditary spastic paraplegia and hereditary motor and sensory neuropathy, Muscle Nerve. 2019;59(4):484–6. doi: 10.1002/mus.26394

40.

Storbeck

, Horsberg Eriksen

, Unger

, Holker

, Aukrust

, Martinez-Carrera

, et al. Phenotypic extremes of BICD2-opathies: From lethal, congenital muscular atrophy with arthrogryposis to asymptomatic with subclinical features, Eur J Hum Genet. 2017;25(9):1040–8. doi: 10.1038/ejhg.2017.98

41.

Huynh

, Vale

. Disease-associated mutations in human BICD2 hyperactivate motility of dynein-dynactin, J Cell Biol. 2017;216(10):3051–60. doi: 10.1083/jcb.201703201

42.

Martinez Carrera

, Gabriel

, Donohoe

, Holker

, Mariappan

, Storbeck

, et al. Novel insights into SMALED2: BICD2 mutations increase microtubule stability and cause defects in axonal and NMJ development, Hum Mol Genet. 2018;27(10):1772–84. doi: 10.1093/hmg/ddy086

43.

Martinez-Carrera

, Wirth

. Dominant spinal muscular atrophy is caused by mutations in BICD2, an important golgin protein, Front Neurosci. 2015;9:401. doi: 10.3389/fnins.2015.00401

44.

Esmaeeli Nieh

, Madou

, Sirajuddin

, Fregeau

, McKnight

, Lexa

, et al. De novo mutations in KIF1A cause progressive encephalopathy and brain atrophy, Ann Clin Transl Neurol. 2015;2(6):623–35. doi: 10.1002/acn3.198

45.

Riviere

, Ramalingam

, Lavastre

, Shekarabi

, Holbert

, Lafontaine

, et al. KIF1A, an axonal transporter of synaptic vesicles, is mutated in hereditary sensory and autonomic neuropathy type 2, Am J Hum Genet. 2011;89(2):219–30. doi: 10.1016/j.ajhg.2011.06.013

46.

Brenner

, Yilmaz

, Muller

, Grehl

, Petri

, Meyer

, et al. Hot-spot KIF5A mutations cause familial ALS, Brain. 2018;141(3):688–97. doi: 10.1093/brain/awx370

47.

Citrigno

, Magariello

, Pugliese

, Di Palma

, Conforti

, Petrone

, et al. Kinesins in neurological inherited diseases: A novel motor-domain mutation in KIF5A gene in a patient from Southern Italy affected by hereditary spastic paraplegia, Acta Neurol Belg. 2018;118(4):643–6. doi: 10.1007/s13760-018-1039-0

48.

Filosto

, Piccinelli

, Palmieri

, Necchini

, Valente

, Zanella

, et al. A novel mutation in the stalk domain of KIF5A causes a slowly progressive atypical motor syndrome, J Clin Med. 2018;8(1). doi: 10.3390/jcm8010017

49.

Nam

, Yoo

, Choi

, Chung

. Wide phenotypic spectrum in axonal Charcot-Marie-Tooth neuropathy type 2 patients with KIF5A mutations, Genes Genomics. 2018;40(1):77–84. doi: 10.1007/s13258-017-0612-x

50.

Dutta

, Diehl

, Onuchic

, Jana

. Structural consequences of hereditary spastic paraplegia disease-related mutations in kinesin, Proc Natl Acad Sci U S A. 2018;115(46):E10822–E9. doi: 10.1073/pnas.1810622115

51.

Jennings

, Chenevert

, Liu

, Mottamal

, Wojcik

, Huckaba

. Characterization of kinesin switch I mutations that cause hereditary spastic paraplegia, PLoS One. 2017;12(7):e0180353. doi: 10.1371/journal.pone.0180353

52.

Ebbing

, Mann

, Starosta

, Jaud

, Schols

, Schule

, et al. Effect of spastic paraplegia mutations in KIF5A kinesin on transport activity, Hum Mol Genet. 2008;17(9):1245–52. doi: 10.1093/hmg/ddn014

53.

Zhao

, Takita

, Tanaka

, Setou

, Nakagawa

, Takeda

, et al. Charcot-Marie-Tooth disease type 2A caused by mutation in a microtubule motor KIF1Bbeta, Cell. 2001;105(5):587–97. doi:

54.

, Takahashi

, Tanaka

, Ichinose

, Niwa

, Wicklund

, et al. KIF1Bbeta mutations detected in hereditary neuropathy impair IGF1R transport and axon growth, J Cell Biol. 2018;217(10):3480–96. doi: 10.1083/jcb.201801085

55.

Jean

, Kiger

. Coordination between RAB GTPase and phosphoinositide regulation and functions, Nat Rev Mol Cell Biol. 2012;13(7):463–70. doi: 10.1038/nrm3379

56.

Taylor

, Enquist

. Axonal spread of neuroinvasive viral infections, Trends Microbiol. 2015;23(5):283–8. doi: 10.1016/j.tim.2015.01.002

57.

Roy

. Seeing the unseen: The hidden world of slow axonal transport, Neuroscientist. 2014;20(1):71–81. doi: 10.1177/1073858413498306

58.

Scott

, Das

, Tang

, Roy

. Mechanistic logic underlying the axonal transport of cytosolic proteins, Neuron. 2011;70(3):441–54. doi: 10.1016/j.neuron.2011.03.022

59.

Tang

, Scott

, Das

, Gitler

, Ganguly

, Roy

. Fast vesicle transport is required for the slow axonal transport of synapsin, J Neurosci. 2013;33(39):15362–75. doi: 10.1523/JNEUROSCI.1148-13.2013

60.

van Niekerk

, Willis

, Chang

, Reumann

, Heise

, Twiss

. Sumoylation in axons triggers retrograde transport of the RNA-binding protein La, Proc Natl Acad Sci U S A. 2007;104(31):12913–8. doi: 10.1073/pnas.0611562104

61.

Harvey

, Dezi

, Pizzinga

, Willis

. Post-transcriptional control of gene expression following stress: The role of RNA-binding proteins, Biochem Soc Trans. 2017;45(4):1007–14. doi: 10.1042/BST20160364

62.

Alami

, Smith

, Carrasco

, Williams

, Winborn

, Han

SSW

, et al. Axonal transport of TDP-43 mRNA granules is impaired by ALS-causing mutations, Neuron. 2014;81(3):536–43. doi: 10.1016/j.neuron.2013.12.018

63.

Purice

, Taylor

. Linking hnRNP function to ALS and FTD pathology, Front Neurosci. 2018;12:326. doi: 10.3389/fnins.2018.00326

64.

Geuens

, Bouhy

, Timmerman

. The hnRNP family: insights into their role in health and disease, Hum Genet. 2016;135(8):851–67. doi: 10.1007/s00439-016-1683-5

65.

, King

, Shorter

, Gitler

. Stress granules as crucibles of ALS pathogenesis, J Cell Biol. 2013;201(3):361–72. doi: 10.1083/jcb.201302044

66.

Verhoeven

, Claeys

, Zuchner

, Schroder

, Weis

, Ceuterick

, et al. MFN2 mutation distribution and genotype/phenotype correlation in Charcot-Marie-Tooth type 2, Brain. 2006;129(Pt 8):2093–102. doi: 10.1093/brain/awl126

67.

Hoppins

, Lackner

, Nunnari

. The machines that divide and fuse mitochondria, Annu Rev Biochem. 2007;76:751–80. doi: 10.1146/annurev.biochem.76.071905.090048

68.

Smirnova

, Griparic

, Shurland

, van der Bliek

. Dynamin-related protein Drp1 is required for mitochondrial division in mammalian cells, Mol Biol Cell. 2001;12(8):2245–56. doi: 10.1091/mbc.12.8.2245

69.

Wangler

, Assia Batzir

, Robak

, Koenig

, Bacino

, Scaglia

, et al. The expanding neurological phenotype of DNM1L-related disorders, Brain. 2018;141(4):e28. doi: 10.1093/brain/awy024

70.

Gandre-Babbe

, van der Bliek

. The novel tail-anchored membrane protein Mff controls mitochondrial and peroxisomal fission in mammalian cells, Mol Biol Cell. 2008;19(6):2402–12. doi: 10.1091/mbc.E07-12-1287

71.

, Huang

, Li

. Drp1-Dependent mitochondrial fission plays critical roles in physiological and pathological progresses in mammals, Int J Mol Sci. 2017;18(1). doi: 10.3390/ijms18010144

72.

Palmer

, Osellame

, Laine

, Koutsopoulos

, Frazier

, Ryan

. MiD49 and MiD51, new components of the mitochondrial fission machinery, EMBO Rep. 2011;12(6):565–73. doi: 10.1038/embor.2011.54

73.

Niemann

, Wagner

, Ruegg

, Suter

. GDAP1 mutations differ in their effects on mitochondrial dynamics and apoptosis depending on the mode of inheritance, Neurobiol Dis. 2009;36(3):509–20. doi: 10.1016/j.nbd.2009.09.011

74.

, Williams

, Ding

. Mitochondrial dynamics and mitochondrial quality control, Redox Biol. 2015;4:6–13. doi: 10.1016/j.redox.2014.11.006

75.

De Vos

, Morotz

, Stoica

, Tudor

, Lau

, Ackerley

, et al. VAPB interacts with the mitochondrial protein PTPIP51 to regulate calcium homeostasis, Hum Mol Genet. 2012;21(6):1299–311. doi: 10.1093/hmg/ddr559

76.

Yalcin

, Zhao

, Stofanko

, O’Sullivan

, Kang

, Roost

, et al. Modeling of axonal endoplasmic reticulum network by spastic paraplegia proteins, Elife. 2017;6. doi: 10.7554/eLife.23882

77.

Kumar

, Kumar Majhi

, Kanta Acharya

, Smalla

, Gundelfinger

, Goswami

. [PREPRINT] TRPV4 interacts with mitochondrial proteins and acts as a mitochondrial structure-function regulator, BioRxiv. 2018. doi: 10.1101/330993

78.

Krols

, Asselbergh

, De Rycke

, De Winter

, Seyer

, Muller

, et al. Sensory neuropathy-causing mutations in ATL3 affect ER-mitochondria contact sites and impair axonal mitochondrial distribution, Hum Mol Genet. 2019;28(4):615–27. doi: 10.1093/hmg/ddy352

79.

Alecu

, Tedeschi

, Behler

, Wunderling

, Lamberz

, Lauterbach

, et al. Localization of 1-deoxysphingolipids to mitochondria induces mitochondrial dysfunction, J Lipid Res. 2017;58(1):42–59. doi: 10.1194/jlr.M068676

80.

Spaulding

, Sleigh

, Morelli

, Pinter

, Burgess

, Seburn

. Synaptic deficits at neuromuscular junctions in two mouse models of charcot-marie-tooth type 2d, J Neurosci. 2016;36(11):3254–67. doi: 10.1523/JNEUROSCI.1762-15.2016

81.

Kratsios

, Pinan-Lucarre

, Kerk

, Weinreb

, Bessereau

, Hobert

. Transcriptional coordination of synaptogenesis and neurotransmitter signaling, Curr Biol. 2015;25(10):1282–95. doi: 10.1016/j.cub.2015.03.028

82.

Lou

. Sensing exocytosis and triggering endocytosis at synapses: Synaptic vesicle exocytosis-endocytosis coupling, Front Cell Neurosci. 2018;12:66. doi: 10.3389/fncel.2018.00066

83.

Frake

, Ricketts

, Menzies

, Rubinsztein

. Autophagy and neurodegeneration, J Clin Invest. 2015;125(1):65–74. doi: 10.1172/JCI73944

84.

Tammineni

, Ye

, Feng

, Aikal

, Cai

. Impaired retrograde transport of axonal autophagosomes contributes to autophagic stress in Alzheimer’s disease neurons, Elife. 2017;6. doi: 10.7554/eLife.21776

85.

Deinhardt

, Salinas

, Verastegui

, Watson

, Worth

, Hanrahan

, et al. Rab5 and Rab7 control endocytic sorting along the axonal retrograde transport pathway, Neuron. 2006;52(2):293–305. doi: 10.1016/j.neuron.2006.08.018

86.

Agola

, Jim

, Ward

, Basuray

, Wandinger-Ness

. Rab GTPases as regulators of endocytosis, targets of disease and therapeutic opportunities, Clin Genet. 2011;80(4):305–18. doi: 10.1111/j.1399-0004.2011.01724.x

87.

Blackstone

. Cellular pathways of hereditary spastic paraplegia, Annu Rev Neurosci. 2012;35:25–47. doi: 10.1146/annurev-neuro-062111-150400

88.

Wijdeven

, Janssen

, Nahidiazar

, Janssen

, Jalink

, Berlin

, et al. Cholesterol and ORP1L-mediated ER contact sites control autophagosome transport and fusion with the endocytic pathway, Nat Commun. 2016;7:11808. doi: 10.1038/ncomms11808

89.

Zuchner

, Noureddine

, Kennerson

, Verhoeven

, Claeys

, De Jonghe

, et al. Mutations in the pleckstrin homology domain of dynamin 2 cause dominant intermediate Charcot-Marie-Tooth disease, Nat Genet. 2005;37(3):289–94. doi: 10.1038/ng1514

90.

Gonzalez-Jamett

, Baez-Matus

, Olivares

, Hinostroza

, Guerra-Fernandez

, Vasquez-Navarrete

, et al. Dynamin-2 mutations linked to Centronuclear Myopathy impair actin-dependent trafficking in muscle cells, Sci Rep. 2017;7(1):4580. doi: 10.1038/s41598-017-04418-w

91.

Snouwaert

, Church

, Jania

, Nguyen

, Wheeler

, Saintsing

, et al. A Mutation in the Borcs7 subunit of the lysosome regulatory BORC complex results in motor deficits and dystrophic axonopathy in mice, Cell Rep. 2018;24(5):1254–65. doi: 10.1016/j.celre2018.06.118

92.

Nian

, Li

, Cheng

, Wu

, Lin

, Tang

, et al. Rab18 collaborates with Rab7 to modulate lysosomal and autophagy activities in the nervous system: An overlapping mechanism for warburg micro syndrome and charcot-marie-tooth neuropathy type 2B, Mol Neurobiol. 2019. doi: 10.1007/s12035-019-1471-z

93.

Ponomareva

, Eliceiri

, Halloran

. Charcot-Marie-Tooth 2b associated Rab7 mutations cause axon growth and guidance defects during vertebrate sensory neuron development, Neural Dev. 2016;11:2. doi: 10.1186/s13064-016-0058-x

94.

Salpietro

, Malintan

, Llano-Rivas

, Spaeth

, Efthymiou

, Striano

, et al. Mutations in the neuronal vesicular SNARE VAMP2 affect synaptic membrane fusion and impair human neurodevelopment, Am J Hum Genet. 2019;104(4):721–30. doi: 10.1016/j.ajhg.2019.02.016

95.

Lipstein

, Verhoeven-Duif

, Michelassi

, Calloway

, van Hasselt

, Pienkowska

, et al. Synaptic UNC13A protein variant causes increased neurotransmission and dyskinetic movement disorder, J Clin Invest. 2017;127(3):1005–18. doi: 10.1172/JCI90259

96.

Prior

, Van Helleputte

, Benoy

, Van Den Bosch

. Defective axonal transport: A common pathological mechanism in inherited and acquired peripheral neuropathies, Neurobiol Dis. 2017;105:300–20. doi: 10.1016/j.nbd.2017.02.009

97.

Vagnoni

, Bullock

. A simple method for imaging axonal transport in aging neurons using the adult Drosophila wing, Nat Protoc. 2016;11(9):1711–23. doi: 10.1038/nprot.2016.112

98.

Lipton

, Maeder

, Shen

. Rapid assembly of presynaptic materials behind the growth cone in dopaminergic neurons is mediated by precise regulation of axonal transport, Cell Rep. 2018;24(10):2709–22. doi: 10.1016/j.celre2018.07.096

99.

Bergamin

, Cieri

, Vazza

, Argenton

, Mostacciuolo

. Zebrafish Tg(hb9:MTS-Kaede): A new in vivo tool for studying the axonal movement of mitochondria, Biochim Biophys Acta. 2016;1860(6):1247–55. doi: 10.1016/j.bbagen.2016.03.007

100.

Sleigh

, Vagnoni

, Twelvetrees

, Schiavo

. Methodological advances in imaging intravital axonal transport, F1000Res. 2017;6:200. doi: 10.12688/f1000research.10433.1

101.

Han

, Williams

, Eggan

. Constructing and deconstructing stem cell models of neurological disease, Neuron. 2011;70(4):626–44. doi: 10.1016/j.neuron.2011.05.003

102.

Noor

, Shapira

, Edri

, Gal

, Wertheim

, Dvir

. 3D Printing of personalized thick and perfusable cardiac patches and hearts, Adv Sci (Weinh). 2019;6(11):1900344. doi: 10.1002/advs.201900344

103.

Jammalamadaka

, Tappa

. Recent advances in biomaterials for 3D printing and tissue engineering, J Funct Biomater. 2018;9(1). doi: 10.3390/jfb9010022

104.

Little

, Luft

, Mosaku

, Ketteler

, Devine

, Gissen

. High-content autophagy analysis in iPSC-derived neurons using immunofluorescence, Methods Mol Biol. 1994:165–74. doi: 10.1007/978-1-4939-9477-9_15

105.

Marrone

, Poser

, Casci

, Japtok

, Reinhardt

, Janosch

, et al. Isogenic FUS-eGFP iPSC reporter lines enable quantification of FUS stress granule pathology that is rescued by drugs inducing autophagy, Stem Cell Reports. 2018;10(2):375–89. doi: 10.1016/j.stemcr.2017.12.018

106.

Fan

, Wali

, Sutharsan

, Bellette

, Crane

, Sue

, et al. Low dose tubulin-binding drugs rescue peroxisome trafficking deficit in patient-derived stem cells in Hereditary Spastic Paraplegia, Biol Open. 2014;3(6):494–502. doi: 10.1242/bio.20147641

107.

Ballatore

, Brunden

, Huryn

, Trojanowski

, Lee

, Smith

3rd . Microtubule stabilizing agents as potential treatment for Alzheimer’s disease and related neurodegenerative tauopathies, J Med Chem. 2012;55(21):8979–96. doi: 10.1021/jm301079z

108.

Hung

, Fu

. Drug candidates in clinical trials for Alzheimer’s disease, J Biomed Sci. 2017;24(1):47. doi: 10.1186/s12929-017-0355-7

109.

Krukowski

, Ma

, Golonzhka

, Laumet

, Gutti

, van Duzer

, et al. HDAC6 inhibition effectively reverses chemotherapy-induced peripheral neuropathy, Pain. 2017;158(6):1126–37. doi: 10.1097/j.pain.0000000000000893

110.

Kalinski

, Kar

, Craver

, Tosolini

, Sleigh

, Lee

, et al. Deacetylation of Miro1 by HDAC6 blocks mitochondrial transport and mediates axon growth inhibition, J Cell Biol. 2019;218(6):1871–90. doi: 10.1083/jcb.201702187

111.

Spurrier

, Shukla

, McLinden

, Johnson

, Giniger

. Altered expression of the Cdk5 activator-like protein, Cdk5alpha, causes neurodegeneration, in part by accelerating the rate of aging, Dis Model Mech. 2018;11(3). doi: 10.1242/dmm.031161

112.

Roy

, Grum-Tokars

, Schavocky

, Saeed

, Staniszewski

, Teich

, et al. Targeting human central nervous system protein kinases: An isoform selective p38alphaMAPK inhibitor that attenuates disease progression in Alzheimer’s disease mouse models, ACS Chem Neurosci. 2015;6(4):666–80. doi: 10.1021/acschemneuro.5b00002

113.

Gibbs

, Kalmar

, Rhymes

, Fellows

, Ahmed

, Whiting

, et al. Inhibiting p38 MAPK alpha rescues axonal retrograde transport defects in a mouse model of ALS, Cell Death Dis. 2018;9(6):596. doi: 10.1038/s41419-018-0624-8

114.

Deane

, Brown

. Induction of heat shock proteins in differentiated human neuronal cells following co-application of celastrol and arimoclomol, Cell Stress Chaperones. 2016;21(5):837–48. doi: 10.1007/s12192-016-0708-2

115.

Pemovska

, Bigenzahn

, Superti-Furga

. Recent advances in combinatorial drug screening and synergy scoring, Curr Opin Pharmacol. 2018;42:102–10. doi: 10.1016/j.coph.2018.07.008

116.

White

, Banerjee

, Gunawardena

. Axonal transport and neurodegeneration: How marine drugs can be used for the development of therapeutics, Mar Drugs. 2016;14(5). doi: 10.3390/md14050102

117.

Wood

MJA

, Talbot

, Bowerman

. Spinal muscular atrophy: antisense oligonucleotide therapy opens the door to an integrated therapeutic landscape, Hum Mol Genet. 2017;26(R2):R151–9. doi: 10.1093/hmg/ddx215

118.

Zhao

, Damle

, Ikeda-Lee

, Kuntz

, Li

, Mohan

, et al. PMP22 antisense oligonucleotides reverse Charcot-Marie-Tooth disease type 1A features in rodent models, J Clin Invest. 2018;128(1):359–68. doi: 10.1172/JCI96499

119.

Food Drug Administration Center for Drugs Evaluation Research. BLA APPROVAL BL 125694/0 ZOLGENSMA (onasemnogene abeparvovec-xioi) AveXis, Inc. FDA Maryland; 2019. doi:

120.

Schiza

, Georgiou

, Kagiava

, Medard

, Richter

, Tryfonos

, et al. Gene replacement therapy in a model of Charcot-Marie-Tooth 4C neuropathy, Brain. 2019;142(5):1227–41. doi: 10.1093/brain/awz064

121.

Kagiava

, Karaiskos

, Richter

, Tryfonos

, Lapathitis

, Sargiannidou

, et al. Intrathecal gene therapy in mouse models expressing CMT1X mutations, Hum Mol Genet. 2018;27(8):1460–73. doi: 10.1093/hmg/ddy056

122.

Gerdts

, Summers

, Milbrandt

, DiAntonio

. Axon self-destruction: New links among SARM1, MAPKs, and NAD+Metabolism, Neuron. 2016;89(3):449–60. doi: 10.1016/j.neuron.2015.12.023

123.

Turkiew

, Falconer

, Reed

, Hoke

. Deletion of Sarm1 gene is neuroprotective in two models of peripheral neuropathy, J Peripher Nerv Syst. 2017;22(3):162–71. doi: 10.1111/jns.12219

124.

Geisler

, Huang

, Strickland

, Doan

, Summers

, Mao

, et al. Gene therapy targeting SARM1 blocks pathological axon degeneration in mice, J Exp Med. 2019;216(2):294–303. doi: 10.1084/jem.20181040

125.

Rizzo

, Bono

, Salani

, Bordoni

, Melzi

, Ruepp

, et al. RNAi/gene therapy combined approach as therapeutic strategy for Charcot-Marie-Tooth 2A (S58.005), Neurology. 2019;92. doi:

126.

Sandelius

, Zetterberg

, Blennow

, Adiutori

, Malaspina

, Laura

, et al. Plasma neurofilament light chain concentration in the inherited peripheral neuropathies, Neurology. 2018;90(6):e518–e24. doi: 10.1212/WNL.0000000000004932

127.

Juneja

, Burns

, Saporta

, Timmerman

. Challenges in modelling the Charcot-Marie-Tooth neuropathies for therapy development JNNP. 2018. doi:

128.

Auer-Grumbach

, Olschewski

, Papic

, Kremer

, McEntagart

, Uhrig

, et al. Alterations in the ankyrin domain of TRPV4 cause congenital distal SMA, scapuloperoneal SMA and HMSN2C, Nat Genet. 2010;42(2):160–4. doi: 10.1038/ng.508

129.

Deng

, Klein

, Yan

, Shi

, Wu

, Fecto

, et al. Scapuloperoneal spinal muscular atrophy and CMT2C are allelic disorders caused by alterations in TRPV4, Nat Genet. 2010;42(2):165–9. doi: 10.1038/ng.509

130.

Landoure

, Zdebik

, Martinez

, Burnett

, Stanescu

, Inada

, et al. Mutations in TRPV4 cause Charcot-Marie-Tooth disease type 2C, Nat Genet. 2010;42(2):170–4. doi: 10.1038/ng.512

131.

Scrivo

, Codogno

, Bomont

. Gigaxonin E3 ligase governs ATG16L1 turnover to control autophagosome production, Nat Commun. 2019;10(1):780. doi: 10.1038/s41467-019-08331-w

132.

Bomont

, Cavalier

, Blondeau

, Ben Hamida

, Belal

, Tazir

, et al. The gene encoding gigaxonin, a new member of the cytoskeletal BTB/kelch repeat family, is mutated in giant axonal neuropathy, Nat Genet. 2000;26(3):370–4. doi: 10.1038/81701

133.

Branchu

, Boutry

, Sourd

, Depp

, Leone

, Corriger

, et al. Loss of spatacsin function alters lysosomal lipid clearance leading to upper and lower motor neuron degeneration, Neurobiol Dis. 2017;102:21–37. doi: 10.1016/j.nbd.2017.02.007