Axonal Transport Impairment in Chemotherapy-Induced Peripheral Neuropathy.

1. Axonal Transport

Axonal transport is a key process of neuronal functionality, due to the peculiar feature of neurons, where the soma can be a long distance from the distal cellular parts of the neurites. At the soma level all the fundamental processes occur, such as DNA replication, transcription, and overall the mRNA translation into proteins, which afterwards must be driven to and along axons to exploit their function. Indeed, the function of axonal transport is to drive proteins and organelles to the distal parts of the cells, thus connecting the heart of the cell with the periphery [ 1 ]. However, the description of axonal transport as a simple refilling method to guarantee the correct axonal composition, even if pivotal, will be much too restrictive, since the communication from periphery to soma is also extremely important. Axonal transport is composed of two distinct mechanisms, a fast axonal and a slow component [ 1 ]. The fast mechanism is a vesicular transport based on microtubules. It can be anterograde, when directed from soma to axons, and in this case it uses kinesin as motor protein, or retrograde, going from axons to soma, and using dynein as motor protein. To guarantee the correct progress of axonal transport, all the involved molecules must be precisely regulated, and the regulation may occur by post-translational modifications of motor proteins [ 2 ]. Moreover, the microtubules are also pivotal to axonal transport, and as well as the motor proteins, they can be precisely regulated by the involvement of Microtubule-Associated Proteins (MAPs). It has been extensively demonstrated that changes in microtubule dynamics (and therefore in axonal transport) are largely due to post-translational modifications of both motor proteins and MAPs ( i.e. , tau) [ 3 ].

Differently from the fast axonal transport, the slow mechanism can only be anterograde, and it mainly concerns cytoskeletal elements [ 11 ]. For neuron structural integrity, the anterograde transport is important, but neurons are also characterized by the ability to respond to external stimuli, in particular to neurotrophic factors, selecting the neurons which have to survive and determining their function. To this aim, the neurotrophic factor signaling pathway exploits the retrograde axonal transport [ 1 ]. Since the axonal transport is essential for neuronal survival, the alteration of its functionality has devastating effects. For this reason, any drug able to interfere with this process, both directly and indirectly by acting on its post-translational regulation, can virtually have a neurotoxic action ( Table 1 ). Confirming this hypothesis, the axonal transport is one of the targets of a class of drugs, the chemotherapeutic agents, which frequently share the induction of a dose-limiting peripheral neuropathy, known in literature as Chemotherapy-Induced Peripheral Neuropathy (CIPN) [ 12 , 13 ].

2. Chemotherapy-Induced Peripheral Neuropathy (CIPN)

CIPN is a side effect that occurs in a large percentage (30%–80%) of cancer patients, very often representing the dose-limiting side effect [ 14 ]. CIPN is generally a sensory dose-dependent neuropathy characterized by paresthesia, sensory loss, dysesthesia, numbness and sometimes by neuropathic pain [ 15 , 16 ]. Less frequently CIPN shows motor or autonomic signs. It has been reported in antitubulin agents, platinum derivatives, proteasome inhibitors and Thalidomide treatment [ 17 ]. It is noteworthy that even inside each class of drugs, CIPN features and severity could be different [ 18 ]. CIPN includes both neuronopathy, axonopathy and myelinopathy [ 13 , 19 , 20 ]. The exact CIPN pathophysiology has not been clarified yet, even if several possible mechanisms have been identified [ 13 ]. DNA damage, mitochondrial derangement, Reactive Oxygen Species (ROS) over-production, inflammation, glutamate toxicity, proteasome or channel dysfunction and axonal transport defects are mechanisms reported to be correlated to peripheral neuropathy induced by different classes of antineoplastic drugs [ 12 , 18 ]. Dorsal Root Ganglia (DRG) are the main target of antineoplastic drug. DRG, in fact, vascularized by fenestrated endothelial cells lacking tight junction, are unprotected by a barrier comparable to the Blood Brain Barrier (BBB) and are exposed to different classes of low and high molecular weight drugs [ 21 ]. These data explain why CIPN is predominantly a sensory pathology. Moreover, although in all neurons axon structure and functions are strictly dependent on perikaryon organelles activity, it is widely demonstrated that the longest nerves are generally the most damaged (extensively reviewed in reference [ 17 , 22 ]) and CIPN is usually characterized by a dying-back neuropathy [ 13 ]. Epidemiological data suggest that diabetes mellitus might be a risk factor associated with CIPN development [ 23 ] and in the last few years a growing number of pharmacogenomic studies have reported genes and single nucleotide polymorphism (SNPs) correlated to CIPN development. In particular, polymorphisms have been demonstrated in DNA repair, growth factors, metal transporter, chemotherapy resistance and glutathione metabolism genes [ 24 , 25 , 26 , 27 ], but no data have been reported to clarify the greater vulnerability of longest fibers.

To date, despite important improvements in cancer treatment and many preclinical and clinical trials, no effective strategy for neuroprotection has been developed [ 17 , 28 ].

3. Antitubulin Agents

Microtubules are essential components of eukaryotic cells cytoskeleton. The basic units of microtubules are αβ-tubulin heterodimers that polymerizing produce 13 protofilaments organized to form the microtubule cylinder. In neuronal cells, besides their central role in mitotic spindle and centrioles formation, microtubules form the tracks on which bidirectional fast axonal transport is based [ 1 ]. The right balance of treadmilling and dynamic instability is crucial for microtubule functions. Consequently, antitubulin agents, both improving polymerization and/or depolymerization, alter cell mitosis and axonal transport. Considering cancer cells high mitotic index, antitubulin agents represent a widely used class of drugs, though characterized by a very high neurotoxicity.

4. Taxanes

Taxanes are plant derived, and Paclitaxel (a natural compound) and Docetaxel (a semisynthetic compound) are the most effective members of this antineoplastic family in the treatment of several solid tumors such as ovarian, lung, breast, prostate and head and neck cancer [ 29 ].

It has been demonstrated that taxanes antineoplastic effect is due to their ability to interfere with mitotic spindle formation in tumor cells [ 4 ]. However, this mode of action cannot be the responsible for well-known taxanes-induced peripheral neurotoxicity [ 30 ]. Paclitaxel-induced peripheral neuropathy is characterized by a predominant sensory neuropathy manifesting with bilateral paresthesia, numbness, tingling and burning pain. Moreover, deep tendon reflex loss is an early symptom of Paclitaxel neurotoxicity [ 19 , 31 ]. Docetaxel causes symptoms similar to those of Paclitaxel but generally they are less severe, probably due to the lower dose used [ 32 ].

The high binding affinity of taxanes to β-tubulin, in addition to blocking mitosis, can perturb microtubule dynamics and stability, inducing microtubule stabilization and determining neuronal damage by different mechanisms. Several experimental models have demonstrated that taxanes induce microtubule polymerization and inhibit depolymerization at the minus end of microtubule [ 4 , 33 ]. Along the microtubule length, Paclitaxel binds to β-tubulin inside the microtubules scaffold in a site including M-loop (β281–293) [ 34 ]. The binding stabilizes GDP-bound αβ-tubulin heterodimers [ 35 ] in a specific and reversible manner. In addition to stabilizing each protofilament structure, Paclitaxel binding also increases the interaction between neighboring protofilaments promoting over stable microtubule bundles formation [ 34 , 36 ] and altering the number of protofilaments into each microtubule [ 37 , 38 ].

An in vivo rat model has demonstrated that DRG are Paclitaxel main site of accumulation [ 21 ] and in vitro experiments have investigated Paclitaxel toxicity in organotypic DRG and in sensitive DRG primary culture [ 39 ]. Several studies have investigated the effect of Paclitaxel-induced polymerization on axonal transport in both in vitro and in vivo experiments. Theiss and Meller [ 40 ] have shown that in vitro 10 µM Paclitaxel decreases anterograde axonal transport of horseradish (HPR) peroxidase in chick DRG neurons. Simultaneously, in axons abnormal aggregation of microtubules appears. Paclitaxel (200 µM) impaired anterograde and retrograde transports have also been demonstrated by Nakata and Yorifujj [ 41 ] in Paclitaxel-bathed rat sciatic nerve using rhodamine-labeled wheat germ agglutinin (WGA-rhodamine). According to these data, but using lower concentrations of Paclitaxel (1, 10, 100 nM) more comparable to clinical doses, Goshima et al. [ 42 ] have demonstrated that Paclitaxel is able to reduce both the anterograde and the retrograde transport of chloro-methylbenzamido dialkylcarbocyanine (CM-Dil)-labeled organelles in chick DRG neurons. Shemesh and Spira [ 43 ] have demonstrated that Paclitaxel concentrations (10, 100 nM) similar to those used by Goshima et al. [ 42 ] are able to interfere with axonal transport also in an in vitro model of neurons isolated from buccal ganglia of the marine mollusk Aplysia Californica. Retrograde axonal transport impairment in this in vitro non mammalian model resulted irreversible and associated with microtubular polar reconfiguration that determines a new chaotic steady state. Moreover, 10 and 100 nM Paclitaxel also reduced the bidirectional transport of mitochondria. Shemesh and Spira [ 43 ] have suggested several hypotheses about how the polar reconfiguration may affect transport, but at the moment none of these has been demonstrated and no data regarding microtubules polar reconfiguration in mammalian peripheral neurons models have been published. However, it is noteworthy that Aplysia Californica and human tubulin share 96% of sequences [ 44 ] and this suggests that the mechanisms through which Paclitaxel acts on microtubule could be the same. In a different model, using both vesicle transport/squid axoplasma and microtubule gliding assay, La Pointe et al. [ 5 ] have shown that the interaction of Paclitaxel with microtubules slightly reduces the anterograde transport (12% and 17%, respectively at 1 µM and 10 µM), but it is not able to alter the retrograde transport. Moreover, in the same paper the authors have demonstrated that Paclitaxel is able to reduce kinesin-dependent transport velocities in isolated axoplasm, but it does not perturb the kinesin-1 activity in microtubule gliding assay. These data suggest that Paclitaxel action could target non-motor MAPs or regulatory proteins of motor proteins ( i.e ., kinases) that were not present in gliding assay. Among these latter there are regulatory molecules, such as kinases and phophatases that inducing post-translational modifications of kinesin or dynein influence their ability to bind microtubule and consequently affect the axonal transport [ 45 , 46 , 47 ]. Such a regulation has been already demonstrated in several neurodegenerative diseases [ 48 ] and could not be excluded in taxanes neurotoxicity. In fact, phosphorylation is a typical post-translational modification after taxanes treatment [ 49 ]. In non-cancer cells, Figueroa-Masot et al. [ 50 ] have highlighted activation of a sub-pool of N -terminal c-Jun protein kinase (JNK) in nucleus of cortical neurons and Nicolini et al. [ 51 ] have demonstrated a sustained phosphorylation of JNK in human neuroblastoma cell line after exposure to Paclitaxel. Moreover, it is known that JNK is able to negatively regulate fast axonal transport perturbing the motor protein kinesin activity [ 52 , 53 ].

Furthermore, taxanes binding sites are located on the inner surface of microtubule and consequently drug binding cannot directly affect the motor MAPs interaction with microtubule. According to this hypothesis taxanes binding should induce conformational change of microtubule that, in turn, affects the axonal transport. Hammond et al. [ 54 ] have demonstrated that Paclitaxel, inducing tubulin post-translational modification (acetylation, detyrosination and polyglutamination), is able to perturb the motor protein kinesin-1 ability to interact with tubulin. Among non-motor MAPs, a key role in neural microtubule development and regulation is played by tau protein [ 55 , 56 ]. Tau protein induces bundling and stabilization of axonal microtubules. Several studies both in vitro and in vivo have investigated the effect of tau protein overexpression on axonal transport, demonstrating impairment of cargo transport along axon [ 57 , 58 , 59 ]. Tau protein is able to bind microtubules in two sites, one of which partially coincides with that for Paclitaxel [ 60 , 61 ]. Treatment with Paclitaxel may then modify the binding of tau protein to microtubules thus affecting axonal transport. Samsonov et al. [ 62 ] have demonstrated that Paclitaxel treatment reduces tau binding to microtubule in Xenopus neurons. On the other hand, Black [ 63 ] in dissociated cultures of rat sympathetic neurons treated with Paclitaxel has not found decrease in tau protein interaction rate with microtubule. Theiss and Meller [ 40 ] have shown that Paclitaxel restricts tau protein to the soma in chick embryo DRG neurons.

5. Vinca-Alkaloids

Vinca alkaloids are derived from Catharanthus roseus plant (formerly, vinca rosea). This group of chemotherapeutic drugs includes both natural alkaloids, such as Vincristine and Vinblastine, and semi-synthetic ones, such as Vinorelbine, Vindesine and Vinflunine. These chemotherapeutic agents have a broad spectrum of activity against hematologic and lymphatic malignancies, as well as solid tumors such as ovarian, testicular, brain, non-small lung cell tumors and sarcomas [ 17 ].

Vinca alkaloids share the antineoplastic effect, based on inhibitory effect on microtubule assembly, while differ in antitumoral spectra and toxicity. Vinca alkaloids form a stable complex with β-tubulin, binding its GTPase domain so that GTP hydrolysis is inhibited and as a consequence β-tubulin is not able to polymerize into microtubules. Despite apparent minimal structural differences among Vinca alkaloids derivatives, they induce significantly dissimilar conformational changes of β-tubulin, due to their different affinity for β-tubulin, (Vincristine > Vinblastine > Vinorelbine > Vinoflunine), and this could explain the distinct neurotoxic profiles: Vincristine is the most effective drug but it is also the most neurotoxic [ 6 , 17 ]. Vincristine-induced peripheral neuropathy is characterized by disturbance in sensory, motor and autonomic functions [ 18 ], and the severity of symptoms depends on several factors: (i) patient age [ 64 ]; (ii) concomitant use of other neurotoxic modalities, such as radiation therapy and chemotherapeutic agents [ 65 ]; (iii) dosage regimen [ 66 ]; (iv) method of calculation of total dose [ 67 ].

The mechanism that underlies Vinca–alkaloid-induced neuropathy remains still unclear, but many possible causes could be involved, such as changes in gene expression, membrane excitability, inflammation and also axonal transport disruption [ 16 , 68 , 69 ]. The Vincristine inhibitory effect on β-tubulin polymerization leads to a severe alteration in axonal microtubules, which entails axonal swelling in myelinated and unmyelinated fibers [ 70 ], due to a progressive accumulation of axoplasmic organelles and vesicles [ 71 ] and damage to nervous fibers. Vincristine also induces chronic neuropathic pain, because C-fibers nociceptors enhance their responsiveness [ 72 ]. Vinblastine induces a less severe peripheral neuropathy in comparison with Vincristine, but a hematological toxicity is observed. Vinorelbine and Vinflunine administration causes a mild sensory-motor neuropathy, which is reversible after drug discontinuation [ 73 ]. Furthermore, C-fiber dysfunction is not observed, saving patients from chronic pain.

Vincristine limitations have prompted the improvement of Vincristine formulation, and liposomal technology has been proposed; the use of Vincristine sulfate liposome is associated with a more favorable PK profile and clinical outcomes [ 74 ]. As reported for Exabepilone, Vincristine has severe inhibitory effects on fast axonal transport, both on anterograde and retrograde direction. In common with Eribulin, Vincristine suppresses microtubule dynamic instability at low concentration and promotes microtubule disassembly at high concentration [ 75 , 76 , 77 ]. Since Vincristine binds along microtubule length, the interactions between microtubules and motors (in particular kinesin-1) are affected, and even dimeric tubulin conformation is compromised [ 5 ].

6. Epothilones

Epothilones are anti-tubulin agents derived from the myxobacterium Sorangium cellulosum, which show high resistance to multidrug resistant (MDR) mechanisms [ 78 ] and are effective against cells that express the multidrug resistance gene MDR-1 and have acquired tubulin mutations. Among epothilones the two major fermentation products are Epothilone A and Epothilone B.

Ixabepilone (BMS-227550) is a second-generation semisynthetic analogue of Epothilone B that shows high metabolic stability, low plasma protein binding and activity especially against taxanes-resistant cell lines [ 79 , 80 ]. Despite Ixabepilone being a good therapeutic option for advanced or metastatic breast cancer, 72% of patients manifest neuropathic symptoms [ 81 ]. However, it is noteworthy that the incidence of its sensory neuropathy is very variable among different administration schedules [ 82 , 83 ].

Sagopilone (ZK-EPO) is a fully synthetic third-generation Epothilone B derivative [ 84 ]. It has been demonstrated to be active against breast cancer, non–small cell lung cancer, cholangiocarcinoma, melanoma and adrenal carcinoma [ 85 ]. Moreover, Hoffmann et al. [ 86 ] have demonstrated that Sagopilone is able to cross the blood-brain barrier. Although two different phase II trials failed to demonstrate Sagopilone’s efficacy against glioblastoma and brain metastasis [ 87 , 88 ].

Epothilones antineoplastic activity is related to their ability to bind microtubules, leading to aberrant spindle formation that determines mitotic arrest and apoptosis through Bcl-2 phosphorylation [ 85 ]. In particular, epothilones bind to tubulin in a site next to taxanes site on β-tubulin inside the microtubule and along the microtubule length. Epothilones binding determines interaction between tubulin dimers intra and inter protofilaments promoting microtubule assembly and reducing their dynamic instability [ 89 ].

Epothilones-induced peripheral neuropathy is primarily an axonal, dose-dependent, sensory distal neuropathy which is reversible. Paresthesia, dysesthesia, numbness and/or burning neuropathic pain in a stocking-glove distributions are commonly reported symptoms [ 83 ].

La Pointe et al. [ 5 ] have demonstrated that Ixabepilone reduces both kinesin-dependent transport velocities in isolated axoplasma and kinesin activity in microtubule gliding assay. The observation that inhibition in entire axoplasm is higher than in in vitro gliding assay suggests that the inhibition of Ixabepilone on axonal transport could be due to its effect on non-motor MAPs present in axoplasma.

Moreover, the greater effect of Ixabepilone on axonal transport and the more severe peripheral neuropathy with respect to those induced by Paclitaxel suggest different post-translational modification of tubulin and different effect on microtubule organization. In fact, Meurer-Grob [ 90 ] has demonstrated that epothilones (like Paclitaxel) are able to modify the microtubule protofilament number and Khrapunovich-Baine et al. [ 36 ] have shown that both epothilones and Paclitaxel affect the COOH-terminus of tubulin influencing the binding of motor and non-motor MAPs and in particular the regulation of kinesin-1 activity.

7. Eribulin

Eribulin mesylate (E7389) is an anticancer drug belonging to the anti-microtubule agents and successfully administered in patients with advanced metastatic breast cancer [ 91 , 92 ]. Moreover, Eribulin is active also against Cisplatin-resistant ovarian cancer [ 93 ]. Structurally it is a simplified macrocyclic, ketone analogue of halicondrin B [ 94 ]. Its antitumoral efficacy, demonstrated in in vitro experiments on several cell lines (U937, Jurkat, HL-60, and HeLa cells) and in four different xenograft models (breast, colon, ovarian cancer and melanoma), is due to irreversible mitotic blockade, related to sustained Bcl-2 phosphorylation [ 95 , 96 ]. Eribulin mesylate binds to the plus end of microtubules (affecting polymerization), but its binding does not influence microtubule depolymeralization [ 77 , 97 ]. Furthermore, after Eribulin mesylate treatment, abnormal mitotic spindles unable to undergo metaphase/anaphase checkpoint are detectable. Formation of mitotic spindle is due to non-functional aggregates of tubulin after Eribulin [ 75 , 98 ].

Regarding CIPN induced by Eribulin, low incidence (21%–26%) of severe neuropathy has been reported [ 97 ]. According to this clinical observation, La Pointe et al. [ 5 ], using vesicle motility assay in isolated squid axoplasma, have demonstrated that Eribulin has a mild effect on fast axonal transport. In particular, Eribulin is able to slightly reduce anterograde fast axonal transport (10% and 13% respectively at concentrations of 1 µM and 10 µM), but it does not inhibit retrograde fast axonal transport. In contrast, Eribulin is ineffective in reducing kinesin-1 activity in microtubule gliding assay. This mild effect could be due to the Eribulin binding only to the plus end and not along the entire microtubule.

Other drugs, differently from taxanes and Vinca alkaloids, which clearly affect the inner structure of axonal transport, other classes of anticancer drugs do not have the microtubule system as main target, but anyway they induce the onset of a peripheral neuropathy, simply through different mechanisms.

8. Platinum Compounds

Platinum compounds are a family of several antineoplastic drugs all characterized by the presence of a Platinum (Pt) atom in their chemical structure. Their common antineoplastic action is based on the formation of adducts between Pt and DNA and/or proteins, which induce the apoptotic death in replicating cells. However, also the post-mitotic neuronal cells are damaged by the drugs, in particular the DRG sensory neurons [ 15 ]. The DRG become an accumulation site for the drugs, and the target of their neurotoxic action [ 15 ]. Although similar, these drugs have different efficacy, and also show a different and in some case peculiar neurotoxic profile, so the consequences of the neurotoxic effect depend on the drug, more serious with Cisplatin and milder with Carboplatin [ 99 ].

9. Cisplatin

Cisplatin ( cis -diamminedichloroplatinum(II)) is the first drug of the platinum compounds family entered in clinical practice, effectively used for the treatment of several tumors, such as ovarian, testicular, and small cell lung cancers [ 99 ]. Starting from a cumulative dose of 300 mg/m 2 neuropathic signs appear, and over the cumulative dose of 500 mg/m 2 a severe sensory neuropathy has been reported in a large amount of patients, with distal paresthesia and sensory ataxia [ 15 , 100 ]. Sometimes these symptoms progress for several months after the end of drug treatment, an event known as “coasting” [ 100 ].

Initially it has been hypothesized an axonopathy, with a “dying back” degeneration mechanism, since neurophysiological studies have evidenced a reduction of Sensory Nerve Action Potentials (SNAPs) amplitude [ 101 ]. However, several authors have demonstrated that Cisplatin induces neuronal cells degeneration through different mechanisms. Surely the formation of Pt-DNA and Pt-proteins adducts has adverse effects also for neurons, but Gill and Windebank [ 102 ] have demonstrated that these cells after Cisplatin exposure undergo to an attempt to re-enter into the cell cycle from G0 phase, and this event is the prelude that triggers the apoptotic neuronal death.

It is noteworthy that an in vitro study reported Cisplatin ability to directly damage the motor proteins kinesin and dynein by adduct formation, and so to directly impair the fast axonal transport [ 7 ], but the available evidence suggests that Cisplatin exposure more likely induces the onset of a neuronopathy. Therefore, axonal degeneration could be only the consequence of the anterograde transport impairment and it may also explain the Cisplatin “coasting” effect, as the latency needed by damage to reach the distal part of the axons.

10. Carboplatin

Carboplatin [ cis -Diammine(1,1-cyclobutanedicarboxylato)platinum(II)] is a Cisplatin analogue frequently used as first line treatment for advanced ovarian carcinoma [ 103 ]. It is characterized by a lower neurotoxic profile, although at high doses (1600 mg/m 2 ) it can induce a neuropathy very similar to that observed after Cisplatin treatment, with painful paresthesia and sensory ataxia [ 104 ].

11. Oxaliplatin

A one-off case is represented by Oxaliplatin [ trans - R , R -cyclohexane-1,2-diamineoxalatoplatinum(II)], a 3rd generation drug used as first line treatment for colorectal cancer [ 100 ]. Oxaliplatin is characterized by a unique neurotoxic profile, since, besides the classical platinum-dependent chronic neurotoxicity, generally arising at a cumulative dose above 550 mg/m 2 , it also shows an acute toxicity, that typically develops within the first 2 days of treatment and manifests with cold-induced distal dysesthesia and paresthesia [ 100 ]. The chronic neuropathy is very similar to that induced by Cisplatin, being a neuronopathy with anterograde axonal transport impairment secondary to neuronal damage [ 105 ]. The acute neurotoxicity seems to mainly affect the axons, by the effect of oxalate salt on the Na + channels present along the axons, thus determining axonal conductivity impairment [ 8 , 106 ]. Moreover, a study have also evidenced an important vacuolization of axonal mitochondrion and a marked reduction of intraepidermal nerve fiber density after Oxaliplatin exposure [ 107 , 108 ]. By the use of neurography, a molecular imaging technology, Schellingerhout et al. [ 109 ] have demonstrated that the Oxaliplatin-dependent neurotoxicity determines an impairment of the retrograde axonal transport involved in neurotrophin transport [ 109 ].

12. Thalidomide

The sunrise of Thalidomide story, a glutamic acid derivative, became sadly and worldwide known when, in the 1950s, it was prescribed as mild sedative and antiemetic to pregnant women, having no knowledge of the teratogenic and anti-abortive side effects, with the result of a number of children affected by serious malformations, and the consequent withdrawal of the drug from the market in the 1960s [ 110 ]. Afterwards, further studies of Thalidomide have allowed to reintroduce it in clinic for the therapy of peculiar dermatological diseases, such as leprosy, and recently the use of Thalidomide has been extended to the treatment of Multiple Myeloma, due to its anti-angiogenic and anti-inflammatory actions [ 111 , 112 ].

After the use of Thalidomide, with typical doses ranging from 50 mg to 300 mg/day, the onset of a dose-dependent peripheral sensory neuropathy has been reported, which symmetrically begins from the distal extremities (hands and feet), and which clinically manifests with paresthesia of the extremities [ 113 , 114 , 115 ]. Further studies confirmed that the sensory neuropathy is length-dependent [ 20 , 116 ]. Many studies have tried to understand the molecular mechanisms at the basis of neuropathy induction, and to identify the target of this side action. Concerning the molecular mechanisms many hypotheses have been formulated, strictly linked to the mechanisms of Thalidomide antiproliferative action [ 114 ]. The reduction of blood supply to the axons due to anti-angiogenic activity could determine their degeneration with greater damage of longest neurites. In addition, some authors focus their attention on the regulatory role of Thalidomide on cytokines signaling pathway, through the inhibition of Nuclear Factor κB (NF-κB), which has a pivotal role in Nerve Growth Factor (NGF) signaling pathway [ 114 ]. This neurotrophin is essential for neuronal survival and arborization, and therefore Thalidomide inhibitive action on NF-κB could interfere with NGF activity, thus leading to neuronal and axonal degeneration.

Thalidomide direct toxic action on neurons rather than on axons, as hypothesized by some authors, casts doubts on the main target of Thalidomide neurotoxicity [ 9 ]. The answer to this question is not purely academic, but very important when thinking about neuroprotective strategies. In fact, if the axon is the drug target, it is possible to hypothesize about reparative/regenerative therapy, but if the damage occurs on DRG neurons, it can easily be irreversible, and therefore a preventive therapy should be applied. Although in vitro only the neuronal degeneration has been observed, by a clinical point of view the axonal damage seems to be more relevant [ 17 ].

Since Thalidomide induces a dose-dependent neuropathy, the clinical recommendation is to use the lowest effective dose, with a careful monitoring of electrophysiological parameters which represent the first neuropathic sign, such as the reduction of SNAPs amplitude, to evaluate the drug dose reduction or even the treatment temporary or permanent withdrawal. In addition, preventive strategies such as anti-oxidant or growth factors administration are currently under evaluation [ 117 , 118 ], as well as the use of derivatives with a very low neurotoxic profile, such as Lenalidomide and Pomalidomide [ 113 ].

13. Bortezomib

Bortezomib, a boronic acid dipeptide, represents the first-in-class member of the proteasome inhibitors family, and it currently is the gold standard treatment for multiple myeloma [ 114 ]. As other antineoplastic drugs, also Bortezomib induces a dose-dependent peripheral neuropathy and patients develop a length-dependent, sensory and painful neuropathy [ 114 , 119 ].

Different mechanisms have been proposed to explain the neurotoxic effect, mainly ascribable to Bortezomib antiproliferative action, which is the inhibition of proteasome complex, the system by which the cell recognizes and eliminates the ubiquitinated proteins, aimed to be destroyed [ 119 ]. The inhibition of such a system induces the apoptotic death pathway cells, but it does not spare the neuronal cells. Protein aggregate accumulation can have a neurotoxic effect directly on soma neurons, but it can also block the correct turnover of axonal proteins, thus leading to axonal transport impairment and then to neuronal death. Anyway, the target of Bortezomib-induced peripheral neuropathy seems to be DRG neurons rather than axons, thus characterizing as a neuronopathy, even if sometimes the damage of Schwann cells has also been reported [ 120 ]. Some authors have reported the increased expression of TNF-alpha in DRG after Bortezomib administration, thus explaining the neurotoxic action [ 119 ]. Other authors have demonstrated a possible role of the immune system in neuropathy onset [ 121 , 122 ], but such a role has been recently questioned [ 123 ]. Undoubtedly Bortezomib treatment also reduces the plasmatic level of some neurotrophins, in particular of BDNF [ 124 , 125 ], with a consequent alteration in neuronal survival. However, Nasu and co. have demonstrated an alteration of the excitatory membrane potentials after Bortezomib administration, and therefore of the correct axonal functionality, thus focusing the attention on the axons [ 126 ]. In this way it is important to report that a direct action of Bortezomib on microtubules and on tubulin besides proteasome inhibition has been described [ 10 , 127 ] without evidences of a survival reduction of neither neurons nor Schwann cells [ 10 , 127 , 128 ]. It is possible that both aspects, axonopathy and neuronopathy, coexist, but probably in the clinical setting the axonopathy symptoms prevail.

Currently there are some analogs of Bortezomib under evaluation, such as Calfizomib and Marazomib, with a very mild profile of neurotoxicity [ 129 , 130 ].

14. Conclusions

Despite different mechanisms of action, the various chemotherapeutic agents share some toxic effect on axonal transport, which has the induction of a dose-limiting peripheral neuropathy as a consequence. Several neuroprotective therapies have already been tried, but with limited effectiveness [ 100 , 118 , 125 ]. The knowledge of the mechanisms underlying such neurotoxic effect has a great importance in terms of neuroprotective strategies against CIPN, since it could be hypothesized as an ad hoc choice of the best preventive/therapeutic approach, based on the drug and on its specific alterations to axonal transport. From this perspective, preventive strategies could partner the action of chemotherapeutic drugs targeting the DRG, while regenerative/reparative approaches could also follow the anticancer treatments. Since the onset of the peripheral neuropathy is often the main dose-limiting side effect of chemotherapy, the chance to limit it represents an huge goal to maximize the antineoplastic drug effect (and patient’s quality of life), without the need to reduce their dose or even to stop the therapy. The knowledge of the mechanisms underlying such neurotoxic effect could be helpful to identify neuroprotective strategies, in order to treat or even to prevent the neuropathy onset, and in this way to maximize the antineoplastic drug effect.

Conflicts of Interest

The authors declare no conflict of interest.

References

1. Morfini G.A. Burns M.R. Stenoien D.L. Brady S.T. Basic Neurochemistry (Eighth Edition) Principles of Molecular, Cellular, and Medical Neurobiology Brady S.T. Siegel G.J. Albers R.W. Price D.L. Elsevier Amsterdam, Netherlands 2012 146 164

2. Gunawardena S. Goldstein L.S. Cargo-carrying motor vehicles on the neuronal highway: Transport pathways and degenerative disease J. Neurobiol. 2004 58 258 271 10.1002/neu.10319 14704957

3. Thaler C.D. Haimo L.T. Microtubules and microtubule motors: Mechanisms of regulation Int. Rev. Cytol. 1996 164 269 327 8575892 10.1016/s0074-7696(08)62388-8

4. Derry W.B. Wilson L. Jordan M.A. Low potency of taxol at microtubule minus ends: Implications for its antimitotic and therapeutic mechanism Cancer Res. 1998 58 1177 1184 9515803

5. LaPointe N.E. Morfini G. Brady S.T. Feinstein S.C. Wilson L. Jordan M.A. Effects of eribulin, vincristine, paclitaxel and ixabepilone on fast axonal transport and kinesin-1 driven microtubule gliding: Implications for chemotherapy-induced peripheral neuropathy Neurotoxicology 2013 37 231 239 10.1016/j.neuro.2013.05.008 23711742 PMC4169189

6. Lobert S. Vulevic B. Correia J.J. Interaction of vinca alkaloids with tubulin: A comparison of vinblastine, vincristine, and vinorelbine Biochemistry 1996 35 6806 6814 10.1021/bi953037i 8639632

7. Russell J.W. Windebank A.J. McNiven M.A. Brat D.J. Brimijoin W.S. Effect of cisplatin and ACTH4-9 on neural transport in cisplatin induced neurotoxicity Brain Res. 1995 676 258 267 10.1016/0006-8993(95)00100-5 7613995

8. Adelsberger H. Quasthoff S. Grosskreutz J. Lepier A. Eckel F. Lersch C. The chemotherapeutic oxaliplatin alters voltage-gated Na + channel kinetics on rat sensory neurons Eur. J. Pharmacol. 2000 406 25 32 10.1016/S0014-2999(00)00667-1 11011028

9. Giannini F. Volpi N. Rossi S. Passero S. Fimiani M. Cerase A. Thalidomide-induced neuropathy: A ganglionopathy? Neurology 2003 60 877 878 10.1212/01.WNL.0000049462.03800.B1 12629253

10. Staff N.P. Podratz J.L. Grassner L. Bader M. Paz J. Knight A.M. Loprinzi C.L. Trushina E. Windebank A.J. Bortezomib alters microtubule polymerization and axonal transport in rat dorsal root ganglion neurons Neurotoxicology 2013 39 124 131 10.1016/j.neuro.2013.09.001 24035926 PMC3844018

11. Stenoien D.L. Brady S.T. Slow Axonal Transport Basic Neurochemistry: Molecular, Cellular and Medical Aspects 6th ed. Siegel G.J. Agranoff B.W. Albers R.W. Fisher S.K. Uhler M.D. Lippincott-Raven Philadelphia, PA, USA 1999

12. Cashman C.R. Höke A. Mechanisms of distal axonal degeneration in peripheral neuropathies Neurosci. Lett. 2015 596 33 50 10.1016/j.neulet.2015.01.048 25617478 PMC4428955

13. Cavaletti G. Marmiroli P. Chemotherapy-induced peripheral neurotoxicity Nat. Rev. Neurol. 2010 6 657 666 10.1038/nrneurol.2010.160 21060341

14. Ferrier J. Pereira V. Busserolles J. Authier N. Balayssac D. Emerging trends in understanding chemotherapy-induced peripheral neuropathy Curr. Pain Headache Rep. 2013 17 364 10.1007/s11916-013-0364-5 23996720

15. Grisold W. Cavaletti G. Windebank A.J. Peripheral neuropathies from chemotherapeutics and targeted agents: Diagnosis, treatment, and prevention Neuro Oncol. 2012 14 iv45 iv54 10.1093/neuonc/nos203 23095830 PMC3480245

16. Jaggi A.S. Singh N. Mechanisms in cancer-chemotherapeutic drugs-induced peripheral neuropathy Toxicology 2012 291 1 9 10.1016/j.tox.2011.10.019 22079234

17. Argyriou A.A. Bruna J. Marmiroli P. Cavaletti G. Chemotherapy-induced peripheral neurotoxicity (CIPN): An update Crit. Rev. Oncol. Hematol. 2012 82 51 77 10.1016/j.critrevonc.2011.04.012 21908200

18. Carozzi V.A. Canta A. Chiorazzi A. Chemotherapy-induced peripheral neuropathy: What do we know about mechanisms? Neurosci. Lett. 2015 596 90 107 10.1016/j.neulet.2014.10.014 25459280

19. Balayssac D. Ferrier J. Descoeur J. Ling B. Pezet D. Eschalier A. Authier N. Chemotherapy-induced peripheral neuropathies: From clinical relevance to preclinical evidence Expert Opin. Drug Saf. 2011 10 407 417 10.1517/14740338.2011.543417 21210753

20. Han Y. Smith M.T. Pathobiology of cancer chemotherapy-induced peripheral neuropathy (CIPN) Front. Pharmacol. 2013 4 156 10.3389/fphar.2013.00156 24385965 PMC3866393

21. Cavaletti G. Cavalletti E. Oggioni N. Sottani C. Minoia C. D’Incalci M. Zucchetti M. Marmiroli P. Tredici G. Distribution of paclitaxel within the nervous system of the rat after repeated intravenous administration Neurotoxicology 2000 21 389 393 10.1046/j.1529-8027.2001.01008-5.x 10894128

22. Windebank A.J. Grisold W. Chemotherapy-induced neuropathy J. Peripher. Nerv. Syst. 2008 13 27 46 10.1111/j.1529-8027.2008.00156.x 18346229

23. Uwah A.N. Ackler J. Leighton J.C. Jr. Pomerantz S. Tester W. The effect of diabetes on oxaliplatin-induced peripheral neuropathy Clin. Colorectal Cancer 2012 11 275 279 10.1016/j.clcc.2012.05.002 22682776

24. Cavaletti G. Alberti P. Marmiroli P. Chemotherapy-induced peripheral neurotoxicity in the era of pharmacogenomics Lancet Oncol. 2011 12 1151 1161 10.1016/S1470-2045(11)70131-0 21719347

25. Park S.B. Kwok J.B. Loy C.T. Friedlander M.L. Lin C.S. Krishnan A.V. Lewis C.R. Kiernan M.C. Paclitaxel-induced neuropathy: Potential association of MAPT and GSK3B genotypes BMC Cancer 2014 14 993 10.1186/1471-2407-14-993 25535399 PMC4364586

26. Avan A. Postma T.J. Ceresa C. Avan A. Cavaletti G. Giovannetti E. Peters G.J. Platinum-induced neurotoxicity and preventive strategies: Past, present, and future Oncologist 2015 20 411 432 10.1634/theoncologist.2014-0044 25765877 PMC4391771

27. Johnson C. Pankratz V.S. Velazquez A.I. Aakre J.A. Loprinzi C.L. Staff N.P. Windebank A.J. Yang P.J. Candidate pathway-based genetic association study of platinum and platinum-taxane related toxicity in a cohort of primary lung cancer patients Neurol. Sci. 2015 349 124 128 10.1016/j.jns.2014.12.041 25586538 PMC4334320

28. Albers J.W. Chaudhry V. Cavaletti G. Donehower R.C. Interventions for preventing neuropathy caused by cisplatin and related compounds Cochrane Database Syst. Rev. 2014 3 CD005228 24687190 10.1002/14651858.CD005228.pub4 PMC10891440

29. Morris P.G. Fornier M.N. Microtubule active agents: Beyond the taxane frontier Clin. Cancer Res. 2008 14 7167 7172 10.1158/1078-0432.CCR-08-0169 19010832

30. Tofthagen C. McAllister R.D. Visovsky C. Peripheral neuropathy caused by Paclitaxel and docetaxel: An evaluation and comparison of symptoms J. Adv. Pract. Oncol. 2013 4 204 215 25032002 PMC4093436

31. Rowinsky E.K. Chaudhry V. Cornblath D.R. Donehower R.C. Neurotoxicity of Taxol J. Natl. Cancer Inst. Monogr. 1993 15 107 115 7912516

32. Hilkens P.H. Verweij J. Vecht C.J. Stoter G. van den Bent M.J. Clinical characteristics of severe peripheral neuropathy induced by docetaxel (Taxotere) Ann. Oncol. 1997 8 187 190 10.1023/A:1008245400251 9093729

33. Caplow M. Zeeberg B. Dynamic properties of microtubules at steady state in the presence of taxol Eur. J. Biochem. 1982 127 319 324 10.1111/j.1432-1033.1982.tb06873.x 6128228

34. Xiao H. Wang H. Zhang X. Tu Z. Bulinski C. Khrapunovich-Baine M. Hogue Angeletti R. Horwitz S.B. Structural evidence for cooperative microtubule stabilization by Taxol and the endogenous dynamics regulator MAP4 ACS Chem. Biol. 2012 7 744 752 10.1021/cb200403x 22270553 PMC3804043

35. Wade R.H. On and around microtubules: An overview Mol. Biotechnol. 2009 43 177 191 10.1007/s12033-009-9193-5 19565362

36. Khrapunovich-Baine M. Menon V. Yang C.P. Northcote P.T. Miller J.H. Angeletti R.H. Fiser A. Horwitz S.B. Xiao H. Hallmarks of molecular action of microtubule stabilizing agents: Effects of epothilone B, ixabepilone, peloruside A, and laulimalide on microtubule conformation J. Biol. Chem. 2011 286 11765 11778 10.1074/jbc.M110.162214 21245138 PMC3064228

37. Andreu J.M. Díaz J.F. Gil R. de Pereda J.M. García de Lacoba M. Peyrot V. Briand C. Towns-Andrews E. Bordas J. Solution structure of Taxotere-induced microtubules to 3-nm resolution. The change in protofilament number is linked to the binding of the taxol side chain J. Biol. Chem. 1994 269 31785 31792 7989352

38. Díaz J.F. Valpuesta J.M. Chacón P. Diakun G. Andreu J.M. Changes in microtubule protofilament number induced by Taxol binding to an easily accessible site. Internal microtubule dynamics J. Biol. Chem. 1998 273 33803 33810 10.1074/jbc.273.50.33803 9837970

39. Scuteri A. Nicolini G. Miloso M. Bossi M. Cavaletti G. Windebank A.J. Tredici G. Paclitaxel toxicity in post-mitotic dorsal root ganglion (DRG) cells Anticancer Res. 2006 26 1065 1070 16619507

40. Theiss C. Meller K. Taxol impairs anterograde axonal transport of microinjected horseradish peroxidase in dorsal root ganglia neurons in vitro Cell Tissue Res. 2000 299 213 224 10.1007/s004410050019 10741462

41. Nakata T. Yorifuji H. Morphological evidence of the inhibitory effect of taxol on the fast axonal transport Neurosci. Res. 1999 35 113 122 10.1016/S0168-0102(99)00074-7 10616915

42. Goshima Y. Usui H. Shiozawa T. Hida T. Kuraoka S. Takeshita S. Yamashita N. Ichikawa Y. Kamiya Y. Gotoh T. Computational analysis of the effects of antineoplastic agents on axonal transport J. Pharmacol. Sci. 2010 114 168 179 10.1254/jphs.09352FP 20859062

43. Shemesh O.A. Spira M.E. Paclitaxel induces axonal microtubules polar reconfiguration and impaired organelle transport: Implications for the pathogenesis of paclitaxel-induced polyneuropathy Acta Neuropathol. 2010 119 235 248 10.1007/s00401-009-0586-0 19727778

44. Shemesh O.A. Erez H. Ginzburg I. Spira M.E. Tau-induced traffic jams reflect organelles accumulation at points of microtubule polar mismatching Traffic 2008 9 458 471 10.1111/j.1600-0854.2007.00695.x 18182010

45. Welte M.A. Bidirectional transport along microtubules Curr. Biol. 2004 14 R525 R537 10.1016/j.cub.2004.06.045 15242636

46. Gindhart J.G. Towards an understanding of kinesin-1 dependent transport pathways through the study of protein-protein interactions Brief Funct. Genomic Proteomic 2006 5 74 86 10.1093/bfgp/ell002 16769683

47. Akhmanova A. Hammer J.A. 3rd Linking molecular motors to membrane cargo Curr. Opin. Cell Biol. 2010 22 479 487 10.1016/j.ceb.2010.04.008 20466533 PMC3393125

48. Morfini G.A. Burns M. Binder L.I. Kanaan N.M. LaPointe N. Bosco D.A. Brown R.H. Jr. Brown H. Tiwari A. Hayward L. Axonal transport defects in neurodegenerative diseases J. Neurosci. 2009 29 12776 12786 10.1523/JNEUROSCI.3463-09.2009 19828789 PMC2801051

49. Blagosklonny M.V. Schulte T. Nguyen P. Trepel J. Neckers L.M. Taxol-induced apoptosis and phosphorylation of Bcl-2 protein involves c-Raf-1 and represents a novel c-Raf-1 signal transduction pathway Cancer Res. 1996 56 1851 1854 8620503

50. Figueroa-Masot X.A. Hetman M. Higgins M.J. Kokot N. Xia Z. Taxol induces apoptosis in cortical neurons by a mechanism independent of Bcl-2 phosphorylation J. Neurosci. 2001 21 4657 4667 11425893 10.1523/JNEUROSCI.21-13-04657.2001 PMC6762365

51. Nicolini G. Rigolio R. Scuteri A. Miloso M. Saccomanno D. Cavaletti G. Tredici G. Effect of trans-resveratrol on signal transduction pathways involved in paclitaxel-induced apoptosis in human neuroblastoma SH-SY5Y cells Neurochem. Int. 2003 42 419 429 10.1016/S0197-0186(02)00132-8 12510025

52. Morfini G. Pigino G. Szebenyi G. You Y. Pollema S. Brady S.T. JNK mediates pathogenic effects of polyglutamine-expanded androgen receptor on fast axonal transport Nat. Neurosci. 2006 9 907 916 10.1038/nn1717 16751763

53. DeBerg H.A. Blehm B.H. Sheung J. Thompson A.R. Bookwalter C.S. Torabi S.F. Schroer T.A. Berger C.L. Lu Y. Trybus K.M. Motor domain phosphorylation modulates kinesin-1 transport J. Biol. Chem. 2013 288 32612 32621 10.1074/jbc.M113.515510 24072715 PMC3820893

54. Hammond J.W. Huang C.F. Kaech S. Jacobson C. Banker G. Verhey K.J. Posttranslational modifications of tubulin and the polarized transport of kinesin-1 in neurons Mol. Biol. Cell 2010 21 572 583 10.1091/mbc.E09-01-0044 20032309 PMC2820422

55. Feinstein S.C. Wilson L. Inability of tau to properly regulate neuronal microtubule dynamics: A loss-of-function mechanism by which tau might mediate neuronal cell death Biochim. Biophys. Acta 2005 1739 268 279 10.1016/j.bbadis.2004.07.002 15615645

56. Choi M.C. Raviv U. Miller H.P. Gaylord M.R. Kiris E. Ventimiglia D. Needleman D.J. Kim M.W. Wilson L. Feinstein S.C. Human microtubule-associated-protein tau regulates the number of protofilaments in microtubules: A synchrotron x-ray scattering study Biophys. J. 2009 97 519 527 10.1016/j.bpj.2009.04.047 19619466 PMC2711315

57. Spittaels K. van den Haute C. van Dorpe J. Bruynseels K. Vandezande K. Laenen I. Geerts H. Mercken M. Sciot R. van Lommel A. Prominent axonopathy in the brain and spinal cord of transgenic mice overexpressing four-repeat human tau protein Am. J. Pathol. 1999 155 2153 2165 10.1016/S0002-9440(10)65533-2 10595944 PMC1866931

58. Stamer K. Vogel R. Thies E. Mandelkow E. Mandelkow E.M. Tau blocks traffic of organelles, neurofilaments, and APP vesicles in neurons and enhances oxidative stress J. Cell Biol. 2002 156 1051 1063 10.1083/jcb.200108057 11901170 PMC2173473

59. Mandelkow E.M. Stamer K. Vogel R. Thies E. Mandelkow E. Clogging of axons by tau, inhibition of axonal traffic and starvation of synapses Neurobiol. Aging 2003 24 1079 1085 10.1016/j.neurobiolaging.2003.04.007 14643379

60. Nogales E. Wolf S.G. Khan I.A. Ludueña R.F. Downing K.H. Structure of tubulin at 6.5 A and location of the taxol-binding site Nature 1995 375 424 427 10.1038/375424a0 7760939

61. Kar S. Fan J. Smith M.J. Goedert M. Amos L.A. Repeat motifs of tau bind to the insides of microtubules in the absence of taxol EMBO J. 2003 22 70 77 10.1093/emboj/cdg001 12505985 PMC140040

62. Samsonov A. Yu J.Z. Rasenick M. Popov S.V. Tau interaction with microtubules in vivo J. Cell Sci. 2004 117 Pt 25 6129 6141 10.1242/jcs.01531 15564376

63. Black M.M. Taxol interferes with the interaction of microtubule-associated proteins with microtubules in cultured neurons J. Neurosci. 1987 7 3695 3702 2890724 10.1523/JNEUROSCI.07-11-03695.1987 PMC6569023

64. DeAngelis L.M. Gnecco C. Taylor L. Warrell R.P. Jr. Evolution of neuropathy and myopathy during intensive vincristine/corticosteroid chemotherapy for non-Hodgkin’s lymphoma Cancer 1991 67 2241 2246 10.1002/1097-0142(19910501)67:9<2241::AID-CNCR2820670905>3.0.CO;2-A 1707337

65. Cassady J.R. Tonnesen G.L. Wolfe L.C. Sallan S.E. Augmentation of vincristine neurotoxicity by irradiation of peripheral nerves Cancer Treat. Rep. 1980 64 963 965 6256075

66. Desai Z.R. van den Berg H.W. Bridges J.M. Shanks R.G. Can severe vincristine neurotoxicity be prevented? Cancer Chemother. Pharmacol. 1982 8 211 214 10.1007/BF00255486 6125275

67. Bradley W.G. Lassman L.P. Pearce G.W. Walton J.N. The neuromyopathy of vincristine in man. Clinical, electrophysiological and pathological studies J. Neurol. Sci. 1970 10 107 131 10.1016/0022-510X(70)90013-4 4314181

68. Pachman D.R. Barton D.L. Watson J.C. Loprinzi C.L. Chemotherapy-induced peripheral neuropathy: Prevention and treatment Clin. Pharmacol. Ther. 2011 90 377 387 10.1038/clpt.2011.115 21814197

69. Xiao W.H. Zheng H. Zheng F.Y. Nuydens R. Meert T.F. Bennett G.J. Mitochondrial abnormality in sensory, but not motor, axons in paclitaxel-evoked painful peripheral neuropathy in the rat Neuroscience 2011 199 461 469 10.1016/j.neuroscience.2011.10.010 22037390 PMC3237950

70. Tanner K.D. Levine J.D. Topp K.S. Microtubule disorientation and axonal swelling in unmyelinated sensory axons during vincristine-induced painful neuropathy in rat J. Comp. Neurol. 1998 395 481 492 10.1002/(SICI)1096-9861(19980615)395:4<481::AID-CNE5>3.0.CO;2-Y 9619501

71. Topp K.S. Tanner K.D. Levine J.D. Damage to the cytoskeleton of large diameter sensory neurons and myelinated axons in vincristine-induced painful peripheral neuropathy in the rat J. Comp. Neurol. 2000 424 563 576 10.1002/1096-9861(20000904)424:4<563::AID-CNE1>3.0.CO;2-U 10931481

72. Tanner K.D. Reichling D.B. Levine J.D. Nociceptor hyper-responsiveness during vincristine-induced painful peripheral neuropathy in the rat J. Neurosci. 1998 18 6480 6491 9698336 10.1523/JNEUROSCI.18-16-06480.1998 PMC6793188

73. Pace A. Bove L. Nisticò C. Ranuzzi M. Innocenti P. Pietrangeli A. Terzoli E. Jandolo B. Vinorelbine neurotoxicity: Clinical and neurophysiological findings in 23 patients J. Neurol. Neurosurg. Psychiatry 1996 61 409 411 10.1136/jnnp.61.4.409 8890782 PMC486585

74. Said R. Tsimberidou A.M. Pharmacokinetic evaluation of vincristine for the treatment of lymphoid malignancies Expert Opin. Drug Metab. Toxicol. 2014 10 483 494 10.1517/17425255.2014.885016 24512004

75. Jordan M.A. Kamath K. Manna T. Okouneva T. Miller H.P. Davis C. Littlefield B.A. Wilson L. The primary antimitotic mechanism of action of the synthetic halichondrin E7389 is suppression of microtubule growth Mol. Cancer Ther. 2005 4 1086 1095 10.1158/1535-7163.MCT-04-0345 16020666

76. Jordan M.A. Wilson L. Microtubules as a target for anticancer drugs Nat. Rev. Cancer 2004 4 253 265 10.1038/nrc1317 15057285

77. Smith J.A. Wilson L. Azarenko O. Zhu X. Lewis B.M. Littlefield B.A. Jordan M.A. Eribulin binds at microtubule ends to a single site on tubulin to suppress dynamic instability Biochemistry 2010 49 1331 1337 10.1021/bi901810u 20030375 PMC2846717

78. Lee J.J. Swain S.M. Development of novel chemotherapeutic agents to evade the mechanisms of multidrug resistance (MDR) Semin. Oncol. 2005 32 6 Suppl. 7 S22 S26 10.1053/j.seminoncol.2005.09.013 16360719

79. Lee F.Y. Borzilleri R. Fairchild C.R. Kamath A. Smykla R. Kramer R. Vite G. Preclinical discovery of ixabepilone, a highly active antineoplastic agent Cancer Chemother. Pharmacol. 2008 63 157 166 10.1007/s00280-008-0724-8 18347795

80. Hunt J.T. Discovery of ixabepilone Mol. Cancer Ther. 2009 8 275 281 10.1158/1535-7163.MCT-08-0999 19174552

81. Steinberg M. Ixabepilone: A novel microtubule inhibitor for the treatment of locally advanced or metastatic breast cancer Clin. Ther. 2008 30 1590 1617 10.1016/j.clinthera.2008.09.015 18840366

82. Canta A. Chiorazzi A. Cavaletti G. Tubulin: A target for antineoplastic drug into the cancer cells but also in the peripheral nervous system Curr. Med. Chem. 2009 16 1315 1324 10.2174/092986709787846488 19355888

83. Argyriou A.A. Marmiroli P. Cavaletti G. Kalofonos H.P. Epothilone-induced peripheral neuropathy: A review of current knowledge J. Pain Symptom Manag. 2011 42 931 940 10.1016/j.jpainsymman.2011.02.022 21621965

84. Klar U. Buchmann B. Schwede W. Skuballa W. Hoffmann J. Lichtner R.B. Total synthesis and antitumor activity of ZK-EPO: The first fully synthetic epothilone in clinical development Angew. Chem. Int. Ed. Engl. 2006 45 7942 7948 10.1002/anie.200602785 17006870

85. Hoffmann J. Vitale I. Buchmann B. Galluzzi L. Schwede W. Senovilla L. Skuballa W. Vivet S. Lichtner R.B. Vicencio J.M. Improved cellular pharmacokinetics and pharmacodynamics underlie the wide anticancer activity of sagopilone Cancer Res. 2008 68 5301 5308 10.1158/0008-5472.CAN-08-0237 18593931

86. Hoffmann J. Fichtner I. Lemm M. Lienau P. Hess-Stumpp H. Rotgeri A. Hofmann B. Klar U. Sagopilone crosses the blood-brain barrier in vivo to inhibit brain tumor growth and metastases Neuro Oncol. 2009 11 158 166 10.1215/15228517-2008-072 18780814 PMC2718987

87. Freedman R.A. Bullitt E. Sun L. Gelman R. Harris G. Ligibel J.A. Krop I.E. Partridge A.H. Eisenberg E. Winer E.P. A phase II study of sagopilone (ZK 219477; ZK-EPO) in patients with breast cancer and brain metastases Clin. Breast Cancer 2011 11 376 383 10.1016/j.clbc.2011.03.024 21697017 PMC3773692

88. Stupp R. Tosoni A. Bromberg J.E. Hau P. Campone M. Gijtenbeek J. Frenay M. Breimer L. Wiesinger H. Allgeier A. Sagopilone (ZK-EPO, ZK 219477) for recurrent glioblastoma. A phase II multicenter trial by the European Organisation for Research and Treatment of Cancer (EORTC) Brain Tumor Group Ann. Oncol. 2011 22 2144 2149 10.1093/annonc/mdq729 21321091 PMC3164435

89. Perez E.A. Microtubule inhibitors: Differentiating tubulin-inhibiting agents based on mechanisms of action, clinical activity, and resistance Mol. Cancer Ther. 2009 8 2086 2095 10.1158/1535-7163.MCT-09-0366 19671735

90. Meurer-Grob P. Kasparian J. Wade R.H. Microtubule structure at improved resolution Biochemistry 2001 40 8000 8008 10.1021/bi010343p 11434769

91. Twelves C. Cortes J. Vahdat L. Olivo M. He Y. Kaufman P.A. Awada A. Efficacy of eribulin in women with metastatic breast cancer: A pooled analysis of two phase 3 studies Breast Cancer Res. Treat 2014 148 553 561 Erratum in: Breast Cancer Res. Treat . 2015 , 149 , 313 10.1007/s10549-014-3245-7 25381136 PMC4243003

92. Vahdat L.T. Pruitt B. Fabian C.J. Rivera R.R. Smith D.A. Tan-Chiu E. Wright J. Tan A.R. Dacosta N.A. Chuang E. Phase II study of eribulin mesylate, a halichondrin B analog, in patients with metastatic breast cancer previously treated with an anthracycline and a taxane J. Clin. Oncol. 2009 27 2954 2961 10.1200/JCO.2008.17.7618 19349550

93. Hensley M.L. Kravetz S. Jia X. Iasonos A. Tew W. Pereira L. Sabbatini P. Whalen C. Aghajanian C.A. Zarwan C. Eribulin mesylate (halichondrin B analog E7389) in platinum-resistant and platinum-sensitive ovarian cancer: A 2-cohort, phase 2 study Cancer 2012 118 2403 2410 10.1002/cncr.26569 21935916 PMC3245795

94. Seletsky B.M. Wang Y. Hawkins L.D. Palme M.H. Habgood G.J. DiPietro L.V. Towle M.J. Salvato K.A. Wels B.F. Aalfs K.K. Structurally simplified macrolactone analogues of halichondrin B Bioorg. Med. Chem. Lett. 2004 14 5547 5550 10.1016/j.bmcl.2004.08.068 15482921

95. Towle M.J. Salvato K.A. Budrow J. Wels B.F. Kuznetsov G. Aalfs K.K. Welsh S. Zheng W. Seletsky B.M. Palme M.H. In vitro and in vivo anticancer activities of synthetic macrocyclic ketone analogues of halichondrin B Cancer Res. 2001 61 1013 1021 11221827

96. Towle M.J. Salvato K.A. Wels B.F. Aalfs K.K. Zheng W. Seletsky B.M. Zhu X. Lewis B.M. Kishi Y. Yu M.J. Eribulin induces irreversible mitotic blockade: Implications of cell-based pharmacodynamics for in vivo efficacy under intermittent dosing conditions Cancer Res. 2011 71 496 505 10.1158/0008-5472.CAN-10-1874 21127197

97. Cortes J. Montero A.J. Glück S. Eribulin mesylate, a novel microtubule inhibitor in the treatment of breast cancer Cancer Treat. Rev. 2012 38 143 151 10.1016/j.ctrv.2011.03.006 21550727

98. Okouneva T. Azarenko O. Wilson L. Littlefield B.A. Jordan M.A. Inhibition of centromere dynamics by eribulin (E7389) during mitotic metaphase Mol. Cancer Ther. 2008 7 2003 2011 10.1158/1535-7163.MCT-08-0095 18645010 PMC2562299

99. Screnci D. McKeage M.J. Platinum neurotoxicity: Clinical profiles, experimental models and neuroprotective approaches J. Inorg. Biochem. 1999 77 105 110 10.1016/S0162-0134(99)00135-X 10626361

100. Park S.B. Krishnan A.V. Lin C.S. Goldstein D. Friedlander M. Kiernan M.C. Mechanisms underlying chemotherapy-induced neurotoxicity and the potential for neuroprotective strategies Curr. Med. Chem. 2008 15 3081 3094 10.2174/092986708786848569 19075655

101. Hansen S.W. Helweg-Larsen S. Trojaborg W. Long-term neurotoxicity in patients treated with cisplatin, vinblastine, and bleomycin for metastatic germ cell cancer J. Clin. Oncol. 1989 7 1457 1461 2476531 10.1200/JCO.1989.7.10.1457

102. Gill J.S. Windebank A.J. Cisplatin-induced apoptosis in rat dorsal root ganglion neurons is associated with attempted entry into the cell cycle J. Clin. Investig. 1998 101 2842 2850 10.1172/JCI1130 9637718 PMC508875

103. Boere I.A. van der Burg M.E. Review of dose-intense platinum and/or paclitaxel containing chemotherapy in advanced and recurrent epithelial ovarian cancer Curr. Pharm. Des. 2012 18 3741 3753 10.2174/138161212802002634 22591417

104. Cavaletti G. Bogliun G. Zincone A. Marzorati L. Melzi P. Frattola L. Marzola M. Bonazzi C. Cantù M.G. Chiari S. Neuro- and ototoxicity of high-dose carboplatin treatment in poor prognosis ovarian cancer patients Anticancer Res. 1998 18 3797 3802 9854499

105. Gamelin E. Gamelin L. Bossi L. Quasthoff S. Clinical aspects and molecular basis of oxaliplatin neurotoxicity: Current management and development of preventive measures Semin. Oncol. 2002 29 5 Suppl. 15 21 33 10.1053/sonc.2002.35525 12422305

106. Grolleau F. Gamelin L. Boisdron-Celle M. Lapied B. Pelhate M. Gamelin E. A possible explanation for a neurotoxic effect of the anticancer agent oxaliplatin on neuronal voltage-gated sodium channels J. Neurophysiol. 2001 85 2293 2297 11353042 10.1152/jn.2001.85.5.2293

107. Xiao W.H. Zheng H. Bennett G.J. Characterization of oxaliplatin-induced chronic painful peripheral neuropathy in the rat and comparison with the neuropathy induced by paclitaxel Neuroscience 2012 203 194 206 10.1016/j.neuroscience.2011.12.023 22200546 PMC3273641

108. Zheng H. Xiao W.H. Bennett G.J. Functional deficits in peripheral nerve mitochondria in rats with paclitaxel- and oxaliplatin-evoked painful peripheral neuropathy Exp. Neurol. 2011 232 154 161 10.1016/j.expneurol.2011.08.016 21907196 PMC3202047

109. Schellingerhout D. LeRoux L.G. Hobbs B.P. Bredow S. Impairment of retrograde neuronal transport in oxaliplatin-induced neuropathy demonstrated by molecular imaging PLoS ONE 2012 7 e45776 10.1371/journal.pone.0045776 23029238 PMC3447809

110. Von Moos R. Stolz R. Cerny T. Gillessen S. Thalidomide: From tragedy to promise Swiss Med. Wkly. 2003 133 77 87 12644961 10.4414/smw.2003.09947

111. Singhal S. Mehta J. Thalidomide in cancer Biomed. Pharmacother. 2002 56 4 12 10.1016/S0753-3322(01)00146-9 11905508

112. Calabrese L. Fleischer A.B. Thalidomide: Current and potential clinical applications Am. J. Med. 2000 108 487 495 10.1016/S0002-9343(99)00408-8 10781782

113. Patel U.H. Mir M.A. Sivik J.K. Raheja D. Pandey M.K. Talamo G. Central neurotoxicity of immunomodulatory drugs in multiple myeloma Hematol. Rep. 2015 7 5704 10.4081/hr.2015.5704 25852850 PMC4378207

114. Mohty B. El-Cheikh J. Yakoub-Agha I. Moreau P. Harousseau J.L. Mohty M. Peripheral neuropathy and new treatments for multiple myeloma: Background and practical recommendations Haematologica 2010 95 311 319 10.3324/haematol.2009.012674 20139393 PMC2817035

115. Briani C. Zara G. Rondinone R. Della Libera S. Ermani M. Ruggero S. Ghirardello A. Zampieri S. Doria A. Thalidomide neurotoxicity: Prospective study in patients with lupus erythematosus Neurology 2004 62 2288 2290 10.1212/01.WNL.0000130499.91775.2C 15210897

116. Chaudhry V. Cornblath D.R. Corse A. Freimer M. Simmons-O’Brien E. Vogelsang G. Thalidomide-induced neuropathy Neurology 2002 59 1872 1875 10.1212/01.WNL.0000037480.59194.85 12499476

117. Areti A. Yerra V.G. Naidu V. Kumar A. Oxidative stress and nerve damage: Role in chemotherapy induced peripheral neuropathy Redox Biol. 2014 2 289 295 10.1016/j.redox.2014.01.006 24494204 PMC3909836

118. Park S.B. Goldstein D. Krishnan A.V. Lin C.S. Friedlander M.L. Cassidy J. Koltzenburg M. Kiernan M.C. Chemotherapy-induced peripheral neurotoxicity: A critical analysis CA Cancer J. Clin. 2013 63 419 437 10.3322/caac.21204 24590861

119. Alé A. Bruna J. Navarro X. Udina E. Neurotoxicity induced by antineoplastic proteasome inhibitors Neurotoxicology 2014 43 28 35 10.1016/j.neuro.2014.02.001 24525285

120. Cavaletti G. Gilardini A. Canta A. Rigamonti L. Rodriguez-Menendez V. Ceresa C. Marmiroli P. Bossi M. Oggioni N. D’Incalci M. Bortezomib-induced peripheral neurotoxicity: A neurophysiological and pathological study in the rat Exp. Neurol. 2007 204 317 325 10.1016/j.expneurol.2006.11.010 17214983

121. Saifee T.A. Elliott K.J. Rabin N. Yong K.L. D’Sa S. Brandner S. Lunn M.P. Blake J. Reilly M.M. Bortezomib-induced inflammatory neuropathy J. Peripher. Nerv. Syst. 2010 15 366 368 Erratum in: J. Peripher. Nerv. Syst. 2011 , 16 , 74 10.1111/j.1529-8027.2010.00287.x 21199108

122. Ravaglia S. Corso A. Piccolo G. Lozza A. Alfonsi E. Mangiacavalli S. Varettoni M. Zappasodi P. Moglia A. Lazzarino M. Immune-mediated neuropathies in myeloma patients treated with bortezomib Clin. Neurophysiol. 2008 119 2507 2512 10.1016/j.clinph.2008.08.007 18829381

123. Carozzi V.A. Renn C.L. Bardini M. Fazio G. Chiorazzi A. Meregalli C. Oggioni N. Shanks K. Quartu M. Serra M.P. Bortezomib-induced painful peripheral neuropathy: An electrophysiological, behavioral, morphological and mechanistic study in the mouse PLoS ONE 2013 8 e72995 10.1371/journal.pone.0072995 24069168 PMC3772181

124. Azoulay D. Lavie D. Horowitz N. Suriu C. Gatt M.E. Akria L. Perlman R. Braester A. Ben-Yehuda D. Bortezomib-induced peripheral neuropathy is related to altered levels of brain-derived neurotrophic factor in the peripheral blood of patients with multiple myeloma Br. J. Haematol. 2014 164 454 456 10.1111/bjh.12624 24164472

125. Argyriou A.A. Iconomou G. Kalofonos H.P. Bortezomib-induced peripheral neuropathy in multiple myeloma: A comprehensive review of the literature Blood 2008 112 1593 1599 10.1182/blood-2008-04-149385 18574024

126. Nasu S. Misawa S. Nakaseko C. Shibuya K. Isose S. Sekiguchi Y. Mitsuma S. Ohmori S. Iwai Y. Beppu M. Bortezomib-induced neuropathy: Axonal membrane depolarization precedes development of neuropathy Clin. Neurophysiol. 2014 125 381 387 10.1016/j.clinph.2013.07.014 23973385

127. Meregalli C. Canta A. Carozzi V.A. Chiorazzi A. Oggioni N. Gilardini A. Ceresa C. Avezza F. Crippa L. Marmiroli P. Bortezomib-induced painful neuropathy in rats: A behavioral, neurophysiological and pathological study in rats Eur. J. Pain 2010 14 343 350 10.1016/j.ejpain.2009.07.001 19695912

128. Meregalli C. Chiorazzi A. Carozzi V.A. Canta A. Sala B. Colombo M. Oggioni N. Ceresa C. Foudah D. La Russa F. Evaluation of tubulin polymerization and chronic inhibition of proteasome as citotoxicity mechanisms in bortezomib-induced peripheral neuropathy Cell Cycle 2014 13 612 621 10.4161/cc.27476 24335344

129. Siegel D. Martin T. Nooka A. Harvey R.D. Vij R. Niesvizky R. Badros A.Z. Jagannath S. McCulloch L. Rajangam K. Integrated safety profile of single-agent carfilzomib: Experience from 526 patients enrolled in 4 phase II clinical studies Haematologica 2013 98 1753 1761 10.3324/haematol.2013.089334 23935022 PMC3815177

130. Thompson J.L. Carfilzomib: A second-generation proteasome inhibitor for the treatment of relapsed and refractory multiple myeloma Ann. Pharmacother. 2013 47 56 62 10.1345/aph.1R561 23300152