Małgorzata Korzeniowska née Wiweger1†
Katarzyna Chabros2†Weronika Rzepnikowska2
Andrzej Kochański2
Dagmara Kabzińska2*- 1Laboratory of Protein Engineering, Mossakowski Medical Research Institute, Polish Academy of Sciences, Warsaw, Poland
- 2Department of Neuromuscular Disorders, Mossakowski Medical Research Institute, Polish Academy of Sciences, Warsaw, Poland
Charcot–Marie–Tooth (CMT) disease is one of the most common inherited neuromuscular disorders, characterized by progressive peripheral nerve degeneration, muscle weakness, and sensory loss. To date, no effective therapy has been developed for CMT. The extreme genetic heterogeneity of CMT, encompassing mutations in more than 50 genes and the involvement of diverse pathological mechanisms, continues to pose significant challenges for disease modeling and therapeutic development. To address these challenges and interrogate specific hypotheses with greater experimental control, researchers have increasingly turned to alternative model organisms that offer genetic tractability and in vivo functional readouts. Zebrafish models have been employed to study hallmark features of CMT, including motor deficits, sensory dysfunction, skeletal abnormalities, and auditory neuropathy. Through the use of forward and reverse genetic screening approaches, as well as transgenic lines, zebrafish have yielded some interesting insights into the functional roles of specific genes implicated in CMT and the effects of pathogenic mutations. Moreover, zebrafish serve as a versatile platform for evaluating potential therapeutic interventions, including pharmacological compounds and gene therapy strategies. This review underscores the value of zebrafish as a robust model for advancing our understanding of CMT pathophysiology. It also addresses the ongoing challenges in genetic diagnosis and highlights the therapeutic potential of this model in guiding future treatments for CMT.
1 Introduction
Inherited peripheral neuropathies represent a broad, heterogeneous group of genetic disorders. They include hereditary sensory-motor conditions, also known as Charcot-Marie-Tooth diseases (CMT), first described by Charcot (1886), distal hereditary motor neuropathies (dHMN), hereditary sensory autonomic neuropathies (HSAN), and hereditary neuropathy with pressure palsies (HNPP). These conditions share partial phenotypic and genetic overlap. Therefore, for the sake of clarity and consistency, the term “CMT” is used throughout this text to encompass the entire spectrum of hereditary motor, sensory, and sensorimotor neuropathies. CMT diseases are the most common inherited neurological condition, with an estimated global prevalence of 1 in 2,500 individuals, but there is substantial variation in prevalence across different regions (Barreto et al., 2016; Skre, 1974).
Clinical manifestations of CMT are highly variable, but typically include muscle weakness and atrophy, typically beginning in the distal muscles of the feet and hands and progressing proximally. Patients often present with foot drop, steppage gait, and decreased or absent deep tendon reflexes. Sensory deficits, particularly affecting pain and temperature perception, as well as proprioception, are also typical and contribute to gait instability and balance difficulties. In some cases, autonomic symptoms such as orthostatic hypotension, bladder dysfunction, and sweating abnormalities may occur. Skeletal abnormalities, including pes cavus (high-arched feet), hammer toes, hand deformities, and scoliosis, frequently arise due to muscle imbalance and weakness (Cortese et al., 2019; Laurá et al., 2019). In recessive forms of the disease, onset typically occurs during the first decade of life. In contrast, dominant forms most commonly manifest in the third or fourth decade; however, cases with very late onset, even in the seventh decade, have also been reported. The age of symptom onset is also influenced by the specific gene involved, the type of mutation (e.g., missense, deletions, insertions, nonsense mutations), and the location of the mutation within the protein.
Charcot-Marie-Tooth is classified according to inheritance patterns and the predominant type of nerve pathology. The major subtypes include CMT Type 1 (CMT1), CMT Type 2 (CMT2), Intermediate CMT (DI-CMT), CMT Type 4 (CMT4), and X-linked CMT (CMTX). Sensory neuropathies are divided into eight types and sixteen subtypes from HSANI to HSANVIII, while motor neuropathies encompass twenty-four to over thirty types of dHMN depending on the classification adopted (Bird, 1993; Pisciotta and Shy, 2023; Schwartzlow and Kazamel, 2019; Tazir and Nouioua, 2024). CMT1, a demyelinating form, is typically caused by mutations in genes encoding myelin proteins such as PMP22, MPZ, and GJB1. This subtype is characterized by slowed nerve conduction velocities due to myelin sheath abnormalities. CMT2, an axonal form, results from mutations in genes involved in axonal structure and function, such as MFN2, RAB7, HSPB1, and presents with normal or mildly reduced nerve conduction velocities but marked axonal degeneration. Intermediate CMT exhibits characteristics of both demyelination and axonal loss and is often associated with mutations in DNM2 and YARS. CMT4 comprises autosomal recessive forms involving various genes and clinical phenotypes. X-linked CMT, primarily caused by mutations in GJB1 encoding connexin 32, typically affects males more severely.
CMT diagnosis involves a comprehensive clinical assessment, family history evaluation, electrophysiological studies, and genetic testing. Neurological examination is critical to delineate patterns of weakness, atrophy, and sensory loss. Family history can provide essential clues regarding inheritance. Electrophysiological studies, including nerve conduction velocity and electromyography, help distinguish between demyelinating and axonal forms. Genetic testing using next-generation sequencing panels or whole-exome sequencing confirms the diagnosis and facilitates genetic counseling.
Genetic characterization of hereditary neuropathies began in the late 20th century. However, the term “CMT genes” is variably defined, with classifications encompassing approximately 50 to over 150 genes, often including other syndromes in which neuropathy is a major component of the phenotype. In pure forms of CMT, the number of associated genes is estimated to be between 50 and 60. However, when broader phenotypes are considered, such as genetic syndromes in which neuropathy is part of the clinical presentation, the number of implicated genes increases to approximately 150. This broad inclusion underscores the considerable genetic heterogeneity of the disorder. In classic sensory-motor neuropathy, over 50 genes were described. A small number of mutations, such as those in PMP22, MPZ, MFN2, and GJB1, account for over 90% of diagnosed cases (Murphy et al., 2012), while others, like GDAP1, are rare and often family-specific (Kabzińska et al., 2022). A common cause of CMT is a 1.4 Mb duplication on chromosome 17 (Lupski et al., 1991; Raeymaekers et al., 1992). Currently, more than 30 genes are associated with motor neuropathies, among them some genes were identified as capable of causing both pure motor neuropathy and classic CMT, such as HSPB1, HSPB8, SORD, and DNAJB2 (Tazir and Nouioua, 2024). Similarly, 15 genes of sensory neuropathy have been described, like SPTLC1, ATL1, NTRK1, and SCN9A (Schwartzlow and Kazamel, 2019). The number of identifiable genes has progressively increased with advancements in next-generation sequencing (NGS)-based diagnostic technologies. Non-Mendelian inheritance patterns, including multilocus and oligogenic inheritance, have also been proposed (Bis-Brewer et al., 2020), and some mutations can exhibit both dominant and recessive inheritance (Rzepnikowska and Kochański, 2018). The molecular diagnosis is further complicated by weak-effect sequence variants, structural mutations (Cutrupi et al., 2018; Gonzaga-Jauregui et al., 2015), and the ambiguous pathogenicity of specific genetic alterations.
Variants are classified into five categories: benign, likely benign, variant of uncertain significance (VUS), likely pathogenic, and pathogenic based on ACMG guidelines (Richards et al., 2015). VUS remain particularly problematic in poorly characterized genes such as WARS1, SARS1, and RAB40B (Favalli et al., 2021). Conflicting variant interpretations further complicate diagnostics; for example, GJB1 shows a 7.3% conflict rate. In GARS1, 49% of variants are VUS and only 8% are classified as pathogenic. Similarly, DNM2 mutations, linked to both myopathy and intermediate CMT, include 43.5% VUS and just 4.6% pathogenic variants (Koutsopoulos et al., 2011). For MFN2, implicated in CMT2A, less than 20% of variants are pathogenic, with over 50% remaining as VUS (Beręsewicz et al., 2018; Züchner et al., 2006). The inconsistency of bioinformatics tools used for pathogenicity prediction underscores the urgent need for improved variant interpretation methods. Most CMT-associated variants have not been functionally validated, as such analyses often lie outside the scope of routine diagnostics. Despite technological advances, only about 50% of CMT cases are genetically diagnosed (Drew et al., 2015; Schabhüttl et al., 2014), with even lower diagnostic yields in HMN and HSAN subgroups (Cortese et al., 2019).
Currently, there is no cure for CMT, and applied therapies focus on symptomatic treatment, maintaining mobility, and improving quality of life. Physical and occupational therapy, alongside assistive devices and customized exercise programs, can help preserve muscle function. Orthopedic interventions, including surgical correction of deformities and orthotic support, aid mobility and pain management. Medications such as gabapentin, pregabalin, and NSAIDs are used to treat neuropathic pain. Genetic counseling provides essential guidance on inheritance, recurrence risks, and reproductive options.
Ongoing advancements in molecular biology and genetics offer hope for targeted therapies.
2 New therapeutical approaches for CMT diseases
Numerous novel therapeutic strategies have been proposed (Okamoto and Takashima, 2023; Pisciotta et al., 2021; Stavrou et al., 2021), offering hope for the development of effective treatments. Several compounds have undergone clinical testing. Among those demonstrating acceptable safety profiles but limited or inconclusive efficacy are PXT3003 and epalrestat. PXT3003 is being developed for the treatment of CMT1A, the most prevalent CMT subtype, caused by a PMP22 gene duplication. It is a combination of baclofen, naltrexone, and sorbitol, three drugs approved for other indications, formulated as an oral solution. In preclinical studies, PXT3003 modestly reduced PMP22 expression, enhanced myelination, increased the number and normalized the size of functional neuromuscular junctions (NMJs), and generally improved the clinical phenotype in CMT1A transgenic rat models (Chumakov et al., 2014; Prukop et al., 2020). A Phase II clinical trial (NCT01401257) provided preliminary evidence of PXT3003’s efficacy and safety in CMT1A patients (Attarian et al., 2012). In the Phase III trial (NCT02579759), the high-dose group demonstrated statistically significant improvement in the primary endpoint. However, concerns regarding the stability of the high-concentration formulation emerged (Attarian et al., 2021), prompting the initiation of a new clinical trial in 2021 (NCT04762758).
Applied Therapeutics has developed a next-generation aldose reductase inhibitor (ARI), AT-007 (govorestat), which effectively inhibits the conversion of glucose to sorbitol. Preliminary results from the INSPIRE clinical trial (NCT05397665) in Sorbitol Dehydrogenase (SORD) Deficiency using AT-007 demonstrated a significant reduction in sorbitol levels in patients (averaging 52%) compared to the placebo group and a statistically significant correlation between sorbitol level, the pre-specified CMT-FOM composite clinical endpoint, and the CMT Health Index (De Grado et al., 2025; GlobeNewswire, 2024; Zhu et al., 2023). SORD encodes sorbitol dehydrogenase, the second enzyme in the polyol pathway, where glucose is first converted into sorbitol by aldose reductase and then into fructose by SORD. Loss-of-function mutations in SORD lead to sorbitol accumulation in cells and plasma (Cortese et al., 2020). Another drug, epalrestat, an aldose reductase inhibitor, blocks the conversion of glucose to sorbitol and has significantly reduced sorbitol levels in fibroblasts derived from SORD-CMT patients (Cortese et al., 2020). Epalrestat is indicated primarily for the management of diabetes-related complications, particularly diabetic peripheral neuropathy. While it does not exert direct neurodegenerative effects, its ability to mitigate hyperglycemia-induced neuronal injury allows for indirect neuroprotection and preservation of peripheral nerve function. The therapeutic effect of epalrestat is based on the inhibition of aldose reductase. Under hyperglycemic conditions, excessive intracellular accumulation of toxic sorbitol in neuronal tissue contributes to osmotic stress, oxidative damage, and subsequent cellular dysfunction (Li et al., 2016; Ran et al., 2024). A similar effect has been observed in animal models as well as in patients with CMT caused by mutations in the SORD gene. By reducing sorbitol levels, epalrestat may attenuate or delay the progression of neuropathy and associated nerve cell damage (Prukop et al., 2020). It is currently approved in several countries for treating diabetic complications and has demonstrated a favorable safety profile (Grewal et al., 2016). A clinical trial evaluating epalrestat’s safety and efficacy for SORD CMT2 was registered in 2023, although recruitment has not yet commenced (NCT05777226).
Gene therapy is among the most actively pursued therapeutic approaches for genetic disorders, including CMT. It encompasses techniques aimed at suppressing disease phenotypes by replacing, modifying, silencing, or repairing defective genetic material in patient cells. Tailored strategies may be required depending on the underlying genetic mechanism. For loss-of-function mutations, gene replacement is typically indicated, whereas dominant-negative or toxic gain-of-function mutations may benefit from gene silencing, editing, or dosage reduction (Stavrou et al., 2023). The majority of gene therapies for CMT are still in the preclinical stage of development (Stavrou et al., 2023). One therapy that has advanced further is VM202, a non-viral, intramuscularly delivered synthetic cDNA hybrid encoding human hepatocyte growth factor (HPHGF). This therapy aims to stimulate nerve regeneration (Ko et al., 2018). A Phase I/IIa clinical trial (NCT05361031) evaluated its safety and tolerability of in patients with CMT1A caused by PMP22 duplication.
A separate investigational approach involves neurotrophin-3 (NT-3), a neurotrophic factor essential for Schwann cell survival and nerve regeneration (Sahenk and Ozes, 2020). Although a Phase I/IIa trial was initiated for CMT1A patients, it is currently suspended due to vector production issues (NCT03520751). In parallel, another early-stage clinical trial is underway to deliver a functional IGHMBP2 gene for treating IGHMBP2-related neuropathies, including CMT2S (NCT05152823).
Another promising avenue involves the use of stem cell-based therapies. Mesenchymal stem cells (MSCs) offer neuroprotective effects and promote regeneration by secreting antioxidant, antiapoptotic, and immunomodulatory molecules. They have shown efficacy in remyelination processes (Yousefi et al., 2019). A completed Phase I study (NCT05333406) assessed the safety and dosing of a single intravenous administration of allogeneic umbilical cord-derived MSCs (EN001) in nine CMT1A patients, with no serious adverse reactions reported. As a follow-up, a clinical trial was registered for CMT1E (caused by point mutations in PMP22) (NCT06218134).
Currently, recruitment is ongoing for a Phase I trial of CLZ-2002 in CMT1 patients. This trial will evaluate the safety and tolerability of intramuscular injections of allogeneic MSC-derived neuronal regeneration-promoting cells (Schwann cell-like cells) (NCT05947578).
3 Advantages and limitations of models used in CMT research
Animal and cellular models have provided crucial insights into human disease mechanisms and therapeutic development, including for genetic disorders such as CMT. Numerous rodent models of CMT have been successfully developed and extensively characterized (Bosco et al., 2021; Juneja et al., 2019). An additional valuable mammalian model includes dogs, in which spontaneous mutations have led to naturally occurring inherited neuropathies that resemble human CMT. Such neuropathies have been identified in at least 22 dog breeds (Granger, 2011). Dogs offer several advantages as disease models, including larger body size, longer lifespan, and greater physiological similarity to humans compared to rodents (Drögemüller et al., 2010). Moreover, as companion animals, they share environmental exposures with humans, adding ecological relevance to disease studies (Skedsmo et al., 2019). Despite these benefits, mammalian models are typically expensive and time-consuming to maintain, and their use raises ethical concerns. Therefore, alternative systems for CMT modeling that adhere to the 3Rs: principle Replacement (whenever possible to use other methods and models to replace the mammals), Reduction (to use the minimal number of animals that is needed to obtain statistically valid results), and Refinement (to minimize animal’s burden during experiment) should be employed whenever feasible.
Beyond animal models, several cellular systems have been established to study CMT pathogenesis. Although yeast models have significant limitations, including a lack of neuronal complexity, absence of genes involved in myelination, and inability to simulate interactions between different cell types, they remain useful for investigating basic cellular mechanisms, screening potential therapeutic compounds (Binięda et al., 2021; Qiu et al., 2023), and identifying candidate targets for intervention (Rzepnikowska et al., 2020a; Rzepnikowska et al., 2020b; Rzepnikowska et al., 2022). Organoids derived from human induced pluripotent stem cells (iPSCs) offer another advanced model system, capable of mimicking complex cellular environments. CMT1A-specific iPSC-derived organoids containing neurons, Schwann cells, muscle cells, endothelial, and glial cells have been developed (Van Lent et al., 2022). These models enable the study of axonal myelination and intercellular interactions. However, a significant limitation is the absence of directional cell growth, which contrasts with the in vivo development of the peripheral nervous system. Consequently, organoids may not be suitable for neuromuscular junction (NMJ)-focused studies (Van Lent et al., 2022). iPSCs are widely employed in disease modeling due to their human origin, high differentiation potential, and accessibility from skin fibroblasts or blood cells. Both patient-derived and genetically engineered iPSC-derived motor neurons serve as relevant tissue models for investigating disease mechanisms and identifying candidate therapies (Feliciano et al., 2021; Perez-Siles et al., 2020; Saporta et al., 2015; Van Lent et al., 2021). Nevertheless, traditional 2D and 3D cultures cannot replicate the full cellular complexity of the peripheral nervous system, limiting their utility, particularly for modeling demyelinating CMT types. While Schwann cells have been generated from human iPSCs (Liu et al., 2012), they - like primary human Schwann cells - have failed to robustly myelinate iPSC-derived neurons in vitro. Notably, myelination has been observed in co-cultures involving iPSC-derived neurons and rat-derived myelinating Schwann cells (Clark et al., 2017).
More complex yet scalable models include the nematode Caenorhabditis elegans and the fruit fly Drosophila melanogaster, both of which are advantageous for high-throughput screening and functional genetic studies. These invertebrates have been used to assess behavioral, cellular, and molecular effects of CMT-related mutations (Cortese et al., 2020; El Fissi et al., 2018; Kitani-Morii and Noto, 2020; Lin et al., 2022; López Del Amo et al., 2015; Brozkova et al., 2015; Soh et al., 2020). However, a significant limitation of these organisms is the absence of Schwann cells and myelinated axons, making them unsuitable for modeling demyelinating forms of CMT (Chung et al., 2020). In contrast, fish models such as zebrafish overcome all these limitations.
4 The zebrafish model of CMT
An ideal model organism for studying neuropathies should have a well-characterized and accessible nervous system, a conserved neuromuscular architecture, and the ability to replicate key aspects of human pathology, including axonal degeneration, demyelination, and neuromuscular dysfunction. Despite notable differences in structure, complexity, and remarkable regenerative capacity, zebrafish fulfill these criteria (Figure 1). It shares significant anatomical and functional similarities with humans in their neuromuscular systems. Both species have a central nervous system (CNS) comprising the brain and spinal cord, and a peripheral nervous system (PNS) consisting of sensory and motor neurons responsible for regulating crucial processes, such as the strength of muscle contractions, which are impaired in CMT (Babin et al., 2014; Singh and Patten, 2022). Although the zebrafish PNS has fewer types of sensory neurons and a less complex branching pattern in the peripheral nerves compared to humans or other mammalian models of CMT, it performs similar functions. At early stages of development, the zebrafish PNS is highly accessible for live imaging, making it a valuable research tool (Chia et al., 2022; Xiao et al., 2015). In zebrafish, peripheral axons are myelinated, though the myelin sheets are thinner and begin forming only after functional axons are established, typically starting at 3–5 days post fertilization (dpf) (D’Rozario et al., 2017). Similarly, as in humans, zebrafish myotomes derived from somites contain three distinct types of muscle fibers (slow, fast, and intermediate). These fibers are organized into repeating units called myomeres, which are divided by a connective tissue (myoseptum) into structural and functional units. However, unlike mammals, zebrafish slow and fast muscles are spatially segregated - slow muscle fibers are located on the superficial (outer) layer of the myotome, while fast muscle fibers occupy the deeper (inner) layers (Daya et al., 2020). This spatial organization provides a unique opportunity to investigate how specific motor neurons target different muscle fiber types, how these connections are affected by neuromuscular disorders like CMT, and how fiber-typespecific deficits contribute to motor dysfunction. Additionally, this segregation simplifies the assessment of fiber-type-specific regeneration or degeneration in response to nerve or muscle damage, enhancing the zebrafish’s utility as a model organism for neuromuscular research.
Figure 1. The usefulness of zebrafish in modeling hereditary neuropathies. Zebrafish serve as a valuable model organism for studying a wide range of human disorders, including hereditary neuropathies. The illustration highlights various human physiological systems and the corresponding CMT disease symptoms that can be effectively modeled using zebrafish.
In addition to their utility in studying neuromuscular connections, zebrafish also provide valuable insights into secondary complications associated with CMT, including skeletal abnormalities. Due to the aquatic environment and the buoyancy it provides, the zebrafish skeleton is not subjected to the same gravitational loading experienced by humans. Nevertheless, zebrafish can develop different axial deformities, including age- or disease-related spine deformities and idiopathic scoliosis (Boswell and Ciruna, 2017). Furthermore, zebrafish can be used to study defects in bone mineralization, vertebral segmentation, or skeletal growth, providing insights into the genetic and molecular mechanisms underlying these conditions (Carnovali et al., 2019; Marí-Beffa et al., 2021; Van Hul et al., 2020).
Hearing deficits in CMT, often linked to auditory neuropathy, can also be effectively modeled in zebrafish (Bever and Fekete, 2002; Vona et al., 2020). Although zebrafish lack a cochlea, which limits their ability to replicate the complex auditory processes seen in humans, their inner ear and lateral line system share structural and functional similarities with mammalian auditory systems, including conserved stereocilia architecture, synaptic mechanisms, and neuronal connectivity (Kindt and Sheets, 2018; Whitfield, 2002). Moreover, the lateral line system is externally accessible and exhibits robust hair cell regeneration, providing a unique platform for studying mechanisms of auditory damage and repair (Hardy et al., 2021). Ototoxic stress can be induced using drugs or environmental stimuli (Domarecka et al., 2020), enabling studies of the cellular and molecular responses to such stressors, facilitating the identification and evaluation of potential therapeutic targets.
The cardiac system in zebrafish also offers valuable insights into CMT-related complications, such as arrhythmias and conduction disturbances in association with peripheral muscle atrophy. Although zebrafish hearts have only a single atrium and ventricle, they share key physiological properties, including similar heart rate and action potential duration, conserved ion channels, conduction pathways, and autonomic regulation of heart function (Tesoriero et al., 2023). Zebrafish are particularly well-suited for real-time imaging of cardiac activity, making them a powerful tool for studying heart function. Additionally, zebrafish models enable the investigation of the role of the autonomic nervous system in regulating heart rate and rhythm, which is often disrupted in CMT (Pedroni et al., 2024).
Some authors suggest that neurogenic bladder disorders that result from peripheral neuropathy, which disrupts the normal communication between the bladder and the nervous system, are associated with CMT. The presence of the urinary bladder has been confirmed in some teleost fish, though its existence in zebrafish was previously questioned. Recent findings by the Catto group demonstrated that the zebrafish urinary bladder is present in adult zebrafish (Jubber et al., 2023) but in contrast to the multi-layered human urothelium, zebrafish urinary bladder is lined by epithelium composed of one or two cell layers, expressing proteins characteristic of both superficial (uroplakins) and basal (Cytokeratin 5 and CD44) layers of human urothelium. Using fluorescent dye, Jubber et al. (2023) showed that the urine accumulates in the zebrafish urinary bladder and is intermittently released via a distinct urethra. While the responses of the urinary bladder to various stimuli have been described in the Atlantic cod (Nilsson, 1970), similar studies in zebrafish are still lacking.
Sweating abnormalities can significantly impact the quality of life in CMT patients. Although fish lack sweat glands and are therefore not suitable for studying sweating dysfunctions in the traditional sense, zebrafish provide a valuable model for assessing autonomic dysfunctions, such as impairments in temperature regulation and sympathetic nervous system function. For example, zebrafish can be tracked as they navigate through a thermal gradient to select their preferred environmental temperature, thereby achieving temperature homeostasis (López-Olmeda and Sánchez-Vázquez, 2011; Palieri et al., 2024). In this way, zebrafish offer key insights into thermal regulation and its impact on broader physiological processes. However, these studies have not yet been conducted in the context of CMT.
To elucidate the molecular and cellular mechanisms underlying CMT in zebrafish and explore potential therapeutic strategies, a variety of experimental approaches can be employed. For example, mitochondrial function, axonal transport, and myelination can be chemically modulated (Azevedo et al., 2020; Toni et al., 2023). In most cases, substances are directly added to the fish water, making this type of experiment straightforward and highly efficient in terms of time, cost, and labor. This approach is particularly advantageous for zebrafish embryos and larvae, which are typically maintained in Petri dishes or multi-well plates, containing relatively small volumes of liquid, thereby enabling effective use of limited quantities of test substances. The availability of diverse transgenic zebrafish lines further amplifies the utility of the zebrafish model by enabling the visualization of cellular events at high resolution. For example, Tg(hb9:MTS-Kaede) line was used to visualize mitochondrial dynamics in motor neurons and the effects of CMT2A-causing mutations on mitochondrial movement (Bergamin et al., 2016), and with Tg(TagRFP-caax), it was possible to assess the effects of CMT2b-associated alterations on long projection sensory neurons (Ponomareva et al., 2016). Transgenic lines, like Tg(mbp:nfsB-egfp) in which bacterial nitroreductase enzyme (NTR) converting metronidazole into a cytotoxic compound is driven under oligodendrocyte-specific promoter, can be used, for e.g., selective and reversible ablation of oligodendrocytes and subsequent demyelination upon treatment with metronidazole (Chung et al., 2013). As zebrafish have an amazing regeneration capacity, once metronidazole is withdrawn, this transgenic system offers the possibility to study remyelination.
Reverse genetic screens were also effective and facilitated efficient and rapid investigation across various genetic backgrounds, allowing for the precise identification of the roles of different genes and modifiers. Among the methods used to study gene function in model organisms, siRNA-mediated knockdown is generally not effective in zebrafish. In contrast, morpholino oligomers (MOs), which typically are ∼ 25-nucleotide molecules designed to block translation or alter splicing by binding to target mRNAs/pre-mRNAs can be used to create morphants – zebrafish embryos and larvae with robust but transient gene knockdown (Stainier et al., 2017; Vettori et al., 2011). Although the use of MOs can be advantageous when studying early development in hypomorphic conditions, however, in other cases, the incomplete knockdown and off-target effects findings should be validated with methods complementary to MO. Since the CRISPR/Cas9 technology has revolutionized genome editing, both transient genetic modifications (crispants) and stable edits via non-homologous end joining (NHEJ) can be created with high efficiency, whereas homology-directed repair (HDR), a key genome editing mechanism in mammalian models like mice, remains far less efficient in zebrafish compared to NHEJ. In addition to morphants and crispants, dominant-negative effects of different genes or their modulators can also be assessed by injecting DNA, RNA, or proteins into one-cell zebrafish embryos and observing their impact on developing embryos (Hong et al., 2016; Mullen et al., 2021).
Dozens of CMT and neuropathy-related genes have been studied in zebrafish (Table 1), one of which is the RAB7 gene. The zebrafish Rab7a shares 97.6% amino acid identity with the human RAB7 protein, with 100% identity at the residues affected in the human disease, specifically L129F, K157N, N161T, and V162M. To study the role of rab7a in the axon growth and guidance defects during sensory neuron development Ponomareva and colleagues (Ponomareva et al., 2016) created constructs in which mutated rab7a was placed under control of cis-regulatory elements from the neurogenin 1 gene, driving expression to Rohon-Beard (RB) spinal sensory neurons. Transient expression was obtained by injecting constructs into one-cell stage embryos, allowing the first analysis already at 23 hours post fertilization (hpf), when the RB neurons start to develop. Using the same constructs and Tol2 transposase stable transgenic lines: Tg(-3.1ngn1:GFP-Rab7), Tg(-3.1ngn1:GFP-Rab7L129F), and Tg(-3.1ngn1:GFP-Rab7K157N), with CMT2b Rab7 mutations in spinal sensory neurons only were generated (Ponomareva et al., 2016). Using those tools, the authors demonstrated that, as in patients, mutations in rab7a caused neurodevelopmental defects. Moreover, reduced axon growth and branching most likely resulted from the expression of a constitutively active form of Rab7a. Tol2 is still used as an efficient tool for random integration of larger DNA fragments into the zebrafish genome, and humanized zebrafish transgenic lines like the Tg (DNM2WT-EGFP), which was created to study subcellular localization of DNM2-EGFP in skeletal muscle cells, is an example of this application (Zhao et al., 2019).
Table 1. Charcot–Marie–Tooth-related genes investigated in zebrafish.
Compared to other vertebrate models, zebrafish, with its easily available large clutches of embryos, offer unique advantages for cost-effective forward genetic screens that allow identification of new genes involved in certain processes. For example, the Tablot group conducted a genetic screen to identify genes that are critical for the development of myelinated axons in zebrafish (Pogoda et al., 2006). In their study, the authors utilized homozygous mutants from the F3 generation, generated through premeiotic mutagenesis with the chemical mutagen ENU (N-ethyl-N-nitrosourea). Their approach involved analyzing the expression of myelin basic protein (mbp) – a robust marker of myelinating glia in the CNS and PNS. By screening 1859 clutches of F3 larvae from 504 F2 families, they identified 13 mutations affecting 10 genes that are essential for myelinated axon development. Of these mutations, st23 mapping in the linkage group 23 was pointed out as a novel gene which is likely to be a good model of CMT2 axonal peripheral neuropathies. Later, the Talbot group showed that st43 mutation affects kinesin motor protein (kif1b), a gene which required to localize myelin mRNA to oligodendrocyte processes, ensuring proper myelin sheath formation around axons, and preventing the ectopic production of myelin-like membrane (Lyons et al., 2009). Although this study did not ultimately identify a new gene, the identification of kif1b in a forward genetic screen demonstrated the model’s relevance for this type of studies. Another member of kinesin proteins – KIF5A, which has two semi-orthologs in zebrafish - kif5Aa and kif5Ab, also sheds light on Kinesin complexity in CMT and reveals determinants of specific Kif5A functions in mitochondrial transport, adaptor binding, and axonal maintenance (Campbell et al., 2014). Similar to SPG10 patients, zebrafish kif5Aasa7168 mutant display striking motor dysfunction. Campbell and co-authors showed that the peripheral sensory axons from the kif5Aa mutant lack mitochondria and degenerate. Moreover, concurrent loss of the kinesin-3, kif1b, or its adaptor kbp, exacerbates axonal degeneration via a non-mitochondrial cargo common to Kif5Aa (Campbell and Marlow, 2013). The example also shows that gene duplication, which in CMT related genes is twice higher than the average for the genome (Kozol et al., 2016), does not discredit the usefulness of the model. Instead, it underscores the model’s capacity to account for genetic variations and complexities, which can be essential for understanding and addressing CMT.
The zebrafish model not only enables the exploration of the functions of genes already associated with CMT but also serves as a crucial tool for investigating the effects of new variants. For example, a zebrafish mutant carrying a rare missense variant in neuregulin 1 (nrg1), provided initial evidences supporting the pathogenicity of a homozygous NRG1 variant identified in a patient with sensory and motor deficits consistent with mixed axonal and demyelinating peripheral neuropathy may cause peripheral neuropathy. These findings suggest that NRG1 should be further investigated in families with peripheral neuropathy of unknown cause (Lysko et al., 2022). The absence of the desired mutation in the zebrafish genome is not a limiting factor. Three CMT-associated substitutions (V155G, Y330C, R137Q) in the cytoplasmic histidyl-tRNA synthetase (hars1) on neurite outgrowth and peripheral nervous system development were also studied in the zebrafish model by injecting Y330C and V155G variants of human HARS1 mRNA (Mullen et al., 2021). Hong et al. (2016) using similar approach, showed that Y223H DGAT2 induced an axonal defect in the peripheral nervous system of zebrafish and Talbot group after identifying a rare R > Q missense variant in NRG1 used zebrafish model to provide evidence indicating that partial loss of NRG1 function indeed may cause peripheral neuropathy in humans (Lysko et al., 2022). By modeling variants of unknown significance, researchers can determine their functional impact, offering valuable information for both clinical interpretation and therapeutic development.
Motor behavior, muscle morphology, and motor neuron in fish over-expressing the G537C mutation in the PH domain of human DYNAMIN-2 were also reflected in human CMT (Bragato et al., 2016). Notably, zebrafish can be used to uncover even more complex scenarios. Holloway and coauthors reported a story of a child with leukemia and no family history of neuropathy who developed severe chemotherapy-induced peripheral neuropathy after vincristine treatment (Holloway et al., 2016). The child was found to have a novel loss-of-function mutation in GARS, suspected of predisposing a patient to severe CIPN. The authors successfully modeled the impact of the mutation in morphant and mRNA-injected zebrafish and obtained a similar phenotype as in the patient, both prior to and after the chemotherapy. Moreover, some of the vincristine-induced neurotoxicity and axonal defects were elevated when fish were co-administered with microtubule stabilizing drug paclitaxel (vincristine is a microtubule-destabilizing drug (Holloway et al., 2016). These findings highlight the potential of zebrafish models for studying disease mechanisms and identifying therapeutic strategies, emphasizing the value of drug combination approaches in mitigating chemotherapy-induced side effects.
5 Conclusion
CMT are complex diseases that require a multidisciplinary diagnostic and therapeutic approach. Ongoing research and close collaboration among geneticists, neurologists, and other healthcare professionals are essential for advancing the understanding and treatment of these challenging neuropathies. Various model organisms are used in CMT research, each offering distinct advantages. Among D. melanogaster, C. elegans, and mouse, zebrafish stand out as a particularly valuable laboratory animal due to their unique advantages (Figure 2).
Figure 2. Comparative advantages and disadvantages of animal models in the study of human diseases. This figure presents a side-by-side comparison of commonly used animal models fruit fly, worm, zebrafish, and rodent, in biomedical research. The comparison helps illustrate the strengths and tradeoffs associated with each organism in the context of disease modeling.
Zebrafish embryos provide a cost-effective and scalable platform for early-stage drug discovery and preclinical testing. Zebrafish can be employed to evaluate compounds or therapies that target or mitigate the effects of genetic variants. Importantly, the zebrafish model not only enables the exploration of genes already associated with CMT but also serves as a crucial tool for investigating the effects of novel or rare genetic variants. Because zebrafish muscles, nerves, visual system, auditory system, cardiac structures, and skeletal components develop rapidly and become functional within 120 hpf, they are particularly well-suited for the rapid assessment of motor deficits, as well as visual, auditory, cardiac, and skeletal abnormalities. In contrast to time- and cost-effective experiments conducted on zebrafish larvae up to 5 dpf, studies of late-onset forms of CMT in adult zebrafish are more demanding but remain valuable. Adult models enable the assessment of disease progression and delayed responses to genetic or pharmacological interventions, thereby significantly advancing our understanding of CMT pathophysiology and therapeutic development. However, their advantage over mammalian models at this stage becomes limited.
It should be noted that gene duplication and the high rate of polymorphism, both common in zebrafish, can complicate genetic analyses. Furthermore, inconsistent nomenclature of some ohnologs and their orthologs continues to cause confusion in comparative genetics and disease modeling (Gasanov et al., 2021). Despite these challenges, the continued application of zebrafish models is expected to substantially contribute to the development of novel therapeutic strategies for disorders within the CMT disease spectrum. A variety of tools – including transgenic lines, antibodies, and dyes are already available for studying CMT; examples are listed in Table 2. Additional resources can be found in an expertly curated and cross-referenced zebrafish research database of the Zebrafish Information Network (ZFIN)1.
Table 2. The list of transgenic lines and antibodies, and dyes used to study Charcot–Marie–Tooth in the zebrafish model.
Author contributions
MK: Writing – original draft, Writing – review and editing. KC: Writing – original draft, Writing – review and editing, Funding acquisition. WR: Writing – original draft, Writing – review and editing. AK: Writing – original draft, Writing – review and editing. DK: Visualization, Writing – original draft, Writing – review and editing.
Funding
The author(s) declare that financial support was received for the research and/or publication of this article. This work was supported by the Mossakowski Medical Research Institute, Polish Academy of Sciences (IMDiK PAN).
Acknowledgments
We would like to acknowledge the support of the National Science Centre, Poland (grant number 2024/08/X/NZ4/00499, awarded to KC). Although this review was not directly funded by the grant, it serves as a conceptual cornerstone for the subsequent research supported by NCN.
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
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The authors declare that Genereative AI was used in the creation of this manuscript. During the preparation of this manuscript, the authors used OpenAI’s ChatGPT [version (e.g., GPT-4)] for assistance in improving the English language of this manuscript. The authors have reviewed and edited the output and take full responsibility for the content of this publication.
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Footnotes
References
Aizawa, H., Bianco, I. H., Hamaoka, T., Miyashita, T., Uemura, O., Concha, M. L., et al. (2005). Laterotopic representation of left-right information onto the dorso-ventral axis of a zebrafish midbrain target nucleus. Curr. Biol. 15, 238–243. doi: 10.1016/j.cub.2005.01.014
Antonellis, A., Dennis, M. Y., Burzynski, G., Huynh, J., Maduro, V., Hodonsky, C. J., et al. (2010). A rare Myelin Protein Zero (MPZ) variant alters enhancer activity in vitro and in vivo. PLoS One 5:e14346. doi: 10.1371/journal.pone.0014346
Asakawa, K., Suster, M. L., Mizusawa, K., Nagayoshi, S., Kotani, T., Urasaki, A., et al. (2008). Genetic dissection of neural circuits by Tol2 transposon-mediated Gal4 gene and enhancer trapping in zebrafish. Proc. Natl. Acad. Sci U S A. 105, 1255–1260. doi: 10.1073/pnas.0704963105
Attarian, S., Vallat, J. M., Magy, L., Funalot, B., Gonnaud, P. M., Lacour, A., et al. (2012). An exploratory randomised double-blind and placebo-controlled phase 2 study of a combination of baclofen, naltrexone and sorbitol (PXT3003) in patients with charcot-marie-tooth disease type 1A. Orphanet J. Rare Dis. 9:199. doi: 10.1186/s13023-014-0199-0
Attarian, S., Young, P., Brannagan, T. H., Adams, D., Van Damme, P., Thomas, F. P., et al. (2021). A double-blind, placebo-controlled, randomized trial of PXT3003 for the treatment of charcot-marie-tooth type 1A. Orphanet J. Rare Dis. 16:433. doi: 10.1186/s13023-021-02040-8
Azevedo, R. D. S., Falcão, K. V. G., Amaral, I. P. G., Leite, A. C. R., and Bezerra, R. S. (2020). Mitochondria as targets for toxicity and metabolism research using zebrafish. Biochim Biophys. Acta Gen. Subj. 1864:129634. doi: 10.1016/j.bbagen.2020.129634
Babin, P. J., Goizet, C., and Raldúa, D. (2014). Zebrafish models of human motor neuron diseases: Advantages and limitations. Prog. Neurobiol. 118, 36–58. doi: 10.1016/j.pneurobio.2014.03.001
Bao, W., Wang, X., Luo, L., and Ni, R. (2021). The lysosomal storage disorder due to fig4a mutation causes robust liver vacuolation in zebrafish. Zebrafish 18, 175–183. doi: 10.1089/zeb.2020.1911
Barreto, L. C., Oliveira, F. S., Nunes, P. S., de França Costa, I. M., Garcez, C. A., Goes, G. M., et al. (2016). Epidemiologic study of charcot-marie-tooth disease: A systematic review. Neuroepidemiology 46, 157–165. doi: 10.1159/000443706
Bercier, V., Brustein, E., Liao, M., Dion, P. A., Lafrenière, R. G., Rouleau, G. A., et al. (2013). WNK1/HSN2 mutation in human peripheral neuropathy deregulates KCC2 expression and posterior lateral line development in zebrafish (Danio rerio). PLoS Genet. 9:e1003124. doi: 10.1371/journal.pgen.1003124
Bercier, V., Hubbard, J. M., Fidelin, K., Duroure, K., Auer, T. O., Revenu, C., et al. (2019). Dynactin1 depletion leads to neuromuscular synapse instability and functional abnormalities. Mol. Neurodegener. 14:27. doi: 10.1186/s13024-019-0327-3
Beręsewicz, M., Charzewski, Ł, Krzyśko, K. A., Kochański, A., and Zabłocka, B. (2018). Molecular modelling of mitofusin 2 for a prediction for Charcot-Marie-Tooth 2A clinical severity. Sci. Rep. 8:16900. doi: 10.1038/s41598-018-35133-9
Bergamin, G., Cieri, D., Vazza, G., Argenton, F., and Mostacciuolo, M. L. (2016). Zebrafish Tg(hb9:mts-kaede): A new in vivo tool for studying the axonal movement of mitochondria. Biochim. Biophys. Acta 1860, 1247–1255. doi: 10.1016/j.bbagen.2016.03.007
Bever, M. M., and Fekete, D. M. (2002). Atlas of the developing inner ear in zebrafish. Dev. Dyn. 223, 536–543. doi: 10.1002/dvdy.10062
Binięda, K., Rzepnikowska, W., Kolakowski, D., Kaminska, J., Szczepankiewicz, A. A., Nieznańska, H., et al. (2021). Mutations in GDAP1 influence structure and function of the trans-golgi network. Int. J. Mol. Sci. 22:914. doi: 10.3390/ijms22020914
Bird, T. D. (1993). “Charcot-Marie-Tooth hereditary neuropathy overview,” in GeneReviews§, eds M. P. Adam, J. Feldman, G. M. Mirzaa, R. A. Pagon, S. E. Wallace, and A. Amemiya (Seattle (WA): University of Washington).
Bis-Brewer, D. M., Gan-Or, Z., Sleiman, P., Hakonarson, H., Fazal, S., et al. (2020). Assessing non-mendelian inheritance in inherited axonopathies. Genet. Med. 22, 2114–2119. doi: 10.1038/s41436-020-0924-0
Bögershausen, N., Krawczyk, H. E., Jamra, R. A., Lin, S. J., Yigit, G., Hüning, I., et al. (2022). WARS1 and SARS1: Two tRNA synthetases implicated in autosomal recessive microcephaly. Hum. Mutat. 43, 1454–1471. doi: 10.1002/humu.24430
Bosco, L., Falzone, Y. M., and Previtali, S. C. (2021). Animal models as a tool to design therapeutical strategies for CMT-like hereditary neuropathies. Brain Sci. 11:1237. doi: 10.3390/brainsci11091237
Boswell, C. W., and Ciruna, B. (2017). Understanding idiopathic scoliosis: A new zebrafish school of thought. Trends Genet. 33, 183–196. doi: 10.1016/j.tig.2017.01.001
Bragato, C., Gaudenzi, G., Blasevich, F., Pavesi, G., Maggi, L., Giunta, M., et al. (2016). Zebrafish as a model to investigate dynamin 2-Related diseases. Sci. Rep. 6:20466. doi: 10.1038/srep20466
Brockmann, S. J., Freischmidt, A., Oeckl, P., Müller, K., Ponna, S. K., Helferich, A. M., et al. (2018). CHCHD10 mutations p.R15L and p.G66V cause motoneuron disease by haploinsufficiency. Hum. Mol. Genet. 27, 706–715. doi: 10.1093/hmg/ddx436
Brozkova, D. S., Deconinck, T., Griffin, L. B., Ferbert, A., Haberlova, J., Mazanec, R., et al. (2015). Loss of function mutations in HARS cause a spectrum of inherited peripheral neuropathies. Brain 138(Pt 8), 2161–2172. doi: 10.1093/brain/awv158
Butterfield, R. J., Stevenson, T. J., Xing, L., Newcomb, T. M., Nelson, B., Zeng, W., et al. (2014). Congenital lethal motor neuron disease with a novel defect in ribosome biogenesis. Neurology 82, 1322–1330. doi: 10.1212/WNL.0000000000000305
Campbell, P. D., and Marlow, F. L. (2013). Temporal and tissue specific gene expression patterns of the zebrafish kinesin-1 heavy chain family, kif5s, during development. Gene Exp. Patt. 13, 271–279. doi: 10.1016/j.gep.2013.05.002
Campbell, P. D., Shen, K., Sapio, M. R., Glenn, T. D., Talbot, W. S., and Marlow, F. L. (2014). Unique function of Kinesin Kif5A in localization of mitochondria in axons. J. Neurosci. 34, 14717–14732. doi: 10.1523/JNEUROSCI.2770-14.2014
Carney, T. J., Dutton, K. A., Greenhill, E., Delfino-Machín, M., Dufourcq, P., Blader, P., et al. (2006). A direct role for Sox10 in specification of neural crest-derived sensory neurons. Development 133, 4619–4630. doi: 10.1242/dev.02668
Carnovali, M., Banfi, G., and Mariotti, M. (2019). Zebrafish models of human skeletal disorders: Embryo and adult swimming together. Biomed. Res. Int. 2019:1253710. doi: 10.1155/2019/1253710
Carrasco Apolinario, M. E., Umeda, R., Teranishi, H., Shan, M., Phurpa, Sebastian, W. A., et al. (2023). Behavioral and neurological effects of Vrk1 deficiency in zebrafish. Biochem. Biophys. Res. Commun. 675, 10–18. doi: 10.1016/j.bbrc.2023.07.005
Castanon, I., Hannich, J. T., Abrami, L., Huber, F., Dubois, M., Müller, M., et al. (2020). Wnt-controlled sphingolipids modulate anthrax toxin receptor palmitoylation to regulate oriented mitosis in zebrafish. Nat. Commun. 11:3317. doi: 10.1038/s41467-020-17196-3
Chaouch, A., Porcelli, V., Cox, D., Edvardson, S., Scarcia, P., De Grassi, A., et al. (2014). Mutations in the mitochondrial citrate carrier SLC25A1 are associated with impaired neuromuscular transmission. J. Neuromuscul. Dis. 1, 75–90. doi: 10.3233/JND-140021
Charcot, J. M. M. P. (1886). Sur une forme particulière d’atrophie musculaire progressive souvent familiale débutant par les pieds et les jambes et atteignant plus tard les mains. Rev. Med. 6, 96–138.
Chen, J., Wu, X., Yao, L., Yan, L., Zhang, L., Qiu, J., et al. (2017). Impairment of cargo transportation caused by gbf1 mutation disrupts vascular integrity and causes hemorrhage in zebrafish embryos. J. Biol. Chem. 292, 2315–2327. doi: 10.1074/jbc.M116.767608
Chen, K., Tian, J., Wang, J., Jia, Z., Zhang, Q., Huang, W., et al. (2021). Lipopolysaccharide-induced TNFα factor (LITAF) promotes inflammatory responses and activates apoptosis in zebrafish Danio rerio. Gene 780:145487. doi: 10.1016/j.gene.2021.145487
Chen, X., Liu, F., Chen, K., Wang, Y., Yin, A., Kang, X., et al. (2022). TFG mutation induces haploinsufficiency and drives axonal Charcot-Marie-Tooth disease by causing neurite degeneration. CNS Neurosci. Ther. 28, 2076–2089. doi: 10.1111/cns.13943
Chia, K., Klingseisen, A., Sieger, D., and Priller, J. (2022). Zebrafish as a model organism for neurodegenerative disease. Front. Mol. Neurosci. 15:940484. doi: 10.3389/fnmol.2022.940484
Chumakov, I., Milet, A., Cholet, N., Primas, G., Boucard, A., Pereira, Y., et al. (2014). Polytherapy with a combination of three repurposed drugs (PXT3003) down-regulates Pmp22 over-expression and improves myelination, axonal and functional parameters in models of CMT1A neuropathy. Orphanet J. Rare Dis. 9:201. doi: 10.1186/s13023-014-0201-x
Chung, A. Y., Kim, P. S., Kim, S., Kim, E., Kim, D., Jeong, I., et al. (2013). Generation of demyelination models by targeted ablation of oligodendrocytes in the zebrafish CNS. Mol. Cells 36, 82–87. doi: 10.1007/s10059-013-0087-9
Chung, K. W., Kim, J. S., and Lee, K. S. (2020). A database of caenorhabditis elegans locomotion and body posture phenotypes for the peripheral neuropathy model. Mol. Cells 43, 880–888. doi: 10.14348/molcells.2020.0178
Clark, A. J., Kaller, M. S., Galino, J., Willison, H. J., Rinaldi, S., and Bennett, D. L. H. (2017). Co-cultures with stem cell-derived human sensory neurons reveal regulators of peripheral myelination. Brain 140, 898–913. doi: 10.1093/brain/awx012
Cortese, A., Wilcox, J. E., Polke, J. M., Poh, R., Skorupinska, M., Rossor, A. M., et al. (2019). Targeted nextgeneration sequencing panels in the diagnosis of Charcot-Marie-Tooth disease. Davis, CA: Dryad.
Cortese, A., Zhu, Y., Rebelo, A. P., Negri, S., Courel, S., Abreu, L., et al. (2020). Biallelic mutations in SORD cause a common and potentially treatable hereditary neuropathy with implications for diabetes. Nat. Genet. 52, 473–481. doi: 10.1038/s41588-020-0615-4
Cutrupi, A. N., Brewer, M. H., Nicholson, G. A., and Kennerson, M. L. (2018). Structural variations causing inherited peripheral neuropathies: A paradigm for understanding genomic organization, chromatin interactions, and gene dysregulation. Mol. Genet. Genom. Med. 6, 422–433. doi: 10.1002/mgg3.390
Davison, J. M., Akitake, C. M., Goll, M. G., Rhee, J. M., Gosse, N., Baier, H., et al. (2007). Transactivation from Gal4-VP16 transgenic insertions for tissue-specific cell labeling and ablation in zebrafish. Dev. Biol. 304, 811–824. doi: 10.1016/j.ydbio.2007.01.033
Daya, A., Donaka, R., and Karasik, D. (2020). Zebrafish models of sarcopenia. Dis. Model Mech. 13:dmm042689. doi: 10.1242/dmm.042689
De Grado, A., Serio, M., Saveri, P., Pisciotta, C., and Pareyson, D. (2025). Charcot-Marie-Tooth disease: A review of clinical developments and its management - What’s new in 2025? Exp. Rev. Neurother. 25, 427–442. doi: 10.1080/14737175.2025.2470980
de la Cruz, C. C., Der-Avakian, A., Spyropoulos, D. D., Tieu, D. D., and Carpenter, E. M. (1999). Targeted disruption of Hoxd9 and Hoxd10 alters locomotor behavior, vertebral identity, and peripheral nervous system development. Dev. Biol. 216, 595–610. doi: 10.1006/dbio.1999.9528
Demy, D. L., Campanari, M. L., Munoz-Ruiz, R., Durham, H. D., Gentil, B. J., and Kabashi, E. (2020). Functional characterization of neurofilament light splicing and misbalance in zebrafish. Cells 9:1238. doi: 10.3390/cells9051238
Domarecka, E., Skarzynska, M., Szczepek, A. J., and Hatzopoulos, S. (2020). Use of zebrafish larvae lateral line to study protection against cisplatin-induced ototoxicity: A scoping review. Int. J. Immunopathol. Pharmacol. 34:2058738420959554. doi: 10.1177/2058738420959554
Drew, A. P., Zhu, D., Kidambi, A., Ly, C., Tey, S., Brewer, M. H., et al. (2015). Improved inherited peripheral neuropathy genetic diagnosis by whole-exome sequencing. Mol. Genet. Genom. Med. 3, 143–154. doi: 10.1002/mgg3.126
Drögemüller, C., Becker, D., Kessler, B., Kemter, E., Tetens, J., Jurina, K., et al. (2010). A deletion in the N-myc downstream regulated gene 1 (NDRG1) gene in Greyhounds with polyneuropathy. PLoS One 5:e11258. doi: 10.1371/journal.pone.0011258
D’Rozario, M., Monk, K. R., and Petersen, S. C. (2017). Analysis of myelinated axon formation in zebrafish. Methods Cell Biol. 138, 383–414. doi: 10.1016/bs.mcb.2016.08.001
Dubińska-Magiera, M., Niedbalska-Tarnowska, J., Migocka-Patrzałek, M., Posyniak, E., and Daczewska, M. (2020). Characterization of Hspb8 in zebrafish. Cells 9:1562. doi: 10.3390/cells9061562
Dutton, K., Abbas, L., Spencer, J., Brannon, C., Mowbray, C., Nikaido, M., et al. (2009). A zebrafish model for Waardenburg syndrome type IV reveals diverse roles for Sox10 in the otic vesicle. Dis. Model Mech. 2, 68–83. doi: 10.1242/dmm.001164
Eijkenboom, I., Sopacua, M., Otten, A. B. C., Gerrits, M. M., Hoeijmakers, J. G. J., Waxman, S. G., et al. (2019a). Expression of pathogenic SCN9A mutations in the zebrafish: A model to study small-fiber neuropathy. Exp. Neurol. 311, 257–264. doi: 10.1016/j.expneurol.2018.10.008
Eijkenboom, I., Vanoevelen, J. M., Hoeijmakers, J. G. J., Wijnen, I., Gerards, M., Faber, C. G., et al. (2019b). A zebrafish model to study small-fiber neuropathy reveals a potential role for GDAP1. Mitochondrion 47, 273–281. doi: 10.1016/j.mito.2019.01.002
El Fissi, N., Rojo, M., Aouane, A., Karatas, E., Poliacikova, G., David, C., et al. (2018). Mitofusin gain and loss of function drive pathogenesis in Drosophila models of CMT2A neuropathy. EMBO Rep. 19:e45241. doi: 10.15252/embr.201745241
Fassier, C., Hutt, J. A., Scholpp, S., Lumsden, A., Giros, B., Nothias, F., et al. (2010). Zebrafish atlastin controls motility and spinal motor axon architecture via inhibition of the BMP pathway. Nat. Neurosci. 13, 1380–1387. doi: 10.1038/nn.2662
Favalli, V., Tini, G., Bonetti, E., Vozza, G., Guida, A., Gandini, S., et al. (2021). Machine learning-based reclassification of germline variants of unknown significance: The RENOVO algorithm. Am. J. Hum. Genet. 108, 682–695. doi: 10.1016/j.ajhg.2021.03.010
Feliciano, C. M., Wu, K., Watry, H. L., Marley, C. B. E., Ramadoss, G. N., Ghanim, H. Y., et al. (2021). Allele-Specific gene editing rescues pathology in a human model of charcot-marie-tooth disease type 2E. Front. Cell Dev. Biol. 9:723023. doi: 10.3389/fcell.2021.723023
Gasanov, E. V., Jędrychowska, J., Kuźnicki, J., and Korzh, V. (2021). Evolutionary context can clarify gene names: Teleosts as a case study. Bioessays 43:e2000258. doi: 10.1002/bies.202000258
Gibbs, E. M., Davidson, A. E., Telfer, W. R., Feldman, E. L., and Dowling, J. J. (2014). The myopathy-causing mutation DNM2-S619L leads to defective tubulation in vitro and in developing zebrafish. Dis. Model Mech. 7, 157–161. doi: 10.1242/dmm.012286
Gibbs, E. M., Davidson, A. E., Trickey-Glassman, A., Backus, C., Hong, Y., Sakowski, S. A., et al. (2013). Two dynamin-2 genes are required for normal zebrafish development. PLoS One 8:e55888. doi: 10.1371/journal.pone.0055888
GlobeNewswire (2024). Applied therapeutics announces positive results from 12-month interim analysis of govorestat (AT-007) in the ongoing inspire phase 3 trial in sorbitol dehydrogenase (SORD) deficiency. Available online at: https://ir.appliedtherapeutics.com/newsreleases/news-release-details/applied-therapeutics-announces-positive-results-12-monthinterim (accessed February 15, 2024).
Gonzaga-Jauregui, C., Harel, T., Gambin, T., Kousi, M., Griffin, L. B., Francescatto, L., et al. (2015). Exome sequence analysis suggests that genetic burden contributes to phenotypic variability and complex neuropathy. Cell Rep. 12, 1169–1183. doi: 10.1016/j.celrep.2015.07.023
Granger, N. (2011). Canine inherited motor and sensory neuropathies: An updated classification in 22 breeds and comparison to Charcot-Marie-Tooth disease. Vet. J. 188, 274–285. doi: 10.1016/j.tvjl.2010.06.003
Grewal, A. S., Bhardwaj, S., Pandita, D., Lather, V., and Sekhon, B. S. (2016). Updates on aldose reductase inhibitors for management of diabetic complications and non-diabetic diseases. Mini. Rev. Med. Chem. 16, 120–162. doi: 10.2174/1389557515666150909143737
Guo, Y., Chen, Y., Yang, M., Xu, X., Lin, Z., Ma, J., et al. (2020). A rare KIF1A missense mutation enhances synaptic function and increases seizure activity. Front. Genet. 11:61. doi: 10.3389/fgene.2020.00061
Hardy, K., Amariutei, A. E., De Faveri, F., Hendry, A., Marcotti, W., and Ceriani, F. (2021). The journal of physiology functional development and regeneration of hair cells in the zebrafish lateral line. J. Physiol. 599, 3913–3936. doi: 10.1113/JP281522
Ho, R. K., and Kane, D. A. (1990). Cell-autonomous action of zebrafish spt-1 mutation in specific mesodermal precursors. Nature 348, 728–730. doi: 10.1038/348728a0
Holloway, M. P., DeNardo, B. D., Phornphutkul, C., Nguyen, K., Davis, C., Jackson, C., et al. (2016). An asymptomatic mutation complicating severe chemotherapy-induced peripheral neuropathy (CIPN): A case for personalised medicine and a zebrafish model of CIPN. NPJ Genom. Med. 1:16016. doi: 10.1038/npjgenmed.2016.16
Hong, Y. B., Kang, J., Kim, J. H., Lee, J., Kwak, G., Hyun, Y. S., et al. (2016). DGAT2 mutation in a family with autosomal-dominant early-onset axonal charcot-marie-tooth disease. Hum. Mutat. 37, 473–480. doi: 10.1002/humu.22959
Jin, B., Xie, L., Zhan, D., Zhou, L., Feng, Z., He, J., et al. (2022). Nrf2 dictates the neuronal survival and differentiation of embryonic zebrafish harboring compromised alanyl-tRNA synthetase. Development 149:dev200342. doi: 10.1242/dev.200342
Jones, E. A., Brewer, M. H., Srinivasan, R., Krueger, C., Sun, G., Charney, K. N., et al. (2012). Distal enhancers upstream of the Charcot-Marie-Tooth type 1A disease gene PMP22. Hum. Mol. Genet. 21, 1581–1591. doi: 10.1093/hmg/ddr595
Jubber, I., Morhardt, D. R., Griffin, J., Cumberbatch, M. G., Glover, M., Zheng, Y., et al. (2023). Analysis of the distal urinary tract in larval and adult zebrafish reveals homology to the human system. Dis. Model Mech. 16:dmm050110. doi: 10.1242/dmm.050110
Juneja, M., Burns, J., Saporta, M. A., and Timmerman, V. (2019). Challenges in modelling the Charcot-Marie-Tooth neuropathies for therapy development. J. Neurol. Neurosurg. Psychiatry 90, 58–67. doi: 10.1136/jnnp-2018-318834
Jung, S. H., Kim, S., Chung, A. Y., Kim, H. T., So, J. H., Ryu, J., et al. (2010). Visualization of myelination in GFP-transgenic zebrafish. Dev. Dyn. 239, 592–597. doi: 10.1002/dvdy.22166
Kabzińska, D., Chabros, K., Kamińska, J., and Kochański, A. (2022). The GDAP1 p.Glu222Lys variant-weak pathogenic effect, cumulative effect of weak sequence variants, or synergy of both factors? Genes (Basel) 13:1546. doi: 10.3390/genes13091546
Kindt, K. S., and Sheets, L. (2018). Transmission disrupted: Modeling auditory synaptopathy in zebrafish. Front. Cell Dev. Biol. 6:114. doi: 10.3389/fcell.2018.00114
Kitani-Morii, F., and Noto, Y. I. (2020). Recent advances in Drosophila models of charcot-marie-tooth disease. Int. J. Mol. Sci. 21:7419. doi: 10.3390/ijms21197419
Ko, K. R., Lee, J., Lee, D., Nho, B., and Kim, S. (2018). Hepatocyte Growth Factor (HGF) promotes peripheral nerve regeneration by activating repair schwann cells. Sci. Rep. 8:8316. doi: 10.1038/s41598-018-26704-x
Koutsopoulos, O. S., Koch, C., Tosch, V., Böhm, J., North, K. N., and Laporte, J. (2011). Mild functional differences of dynamin 2 mutations associated to centronuclear myopathy and Charcot-Marie Tooth peripheral neuropathy. PLoS One 6:e27498. doi: 10.1371/journal.pone.0027498
Kozol, R. A., Abrams, A. J., James, D. M., Buglo, E., Yan, Q., and Dallman, J. E. (2016). Function over form: Modeling groups of inherited neurological conditions in zebrafish. Front. Mol. Neurosci. 9:55. doi: 10.3389/fnmol.2016.00055
Laurá, M., Pipis, M., Rossor, A. M., and Reilly, M. M. (2019). Charcot-Marie-Tooth disease and related disorders: An evolving landscape. Curr. Opin. Neurol. 32, 641–650. doi: 10.1097/WCO.0000000000000735
Li, Q. R., Wang, Z., Zhou, W., Fan, S. R., Ma, R., Xue, L., et al. (2016). Epalrestat protects against diabetic peripheral neuropathy by alleviating oxidative stress and inhibiting polyol pathway. Neural Regen. Res. 11, 345–351. doi: 10.4103/1673-5374.177745
Lin, S. J., Vona, B., Barbalho, P. G., Kaiyrzhanov, R., Maroofian, R., Petree, C., et al. (2021). Biallelic variants in KARS1 are associated with neurodevelopmental disorders and hearing loss recapitulated by the knockout zebrafish. Genet. Med. 23, 1933–1943. doi: 10.1038/s41436-021-01239-1
Lin, S. J., Vona, B., Porter, H. M., Izadi, M., Huang, K., Lacassie, Y., et al. (2022). Biallelic variants in WARS1 cause a highly variable neurodevelopmental syndrome and implicate a critical exon for normal auditory function. Hum. Mutat. 43, 1472–1489. doi: 10.1002/humu.24435
Lindzon, J., List, M., Geissah, S., Ariaz, A., Zhao, M., and Dowling, J. J. (2025). Characterization of a novel zebrafish model of MTMR5-associated Charcot-Marie-Tooth disease type 4B3. Brain Commun. 7:fcaf077. doi: 10.1093/braincomms/fcaf077
Liu, Q., Spusta, S. C., Mi, R., Lassiter, R. N., Stark, M. R., Höke, A., et al. (2012). Human neural crest stem cells derived from human ESCs and induced pluripotent stem cells: Induction, maintenance, and differentiation into functional schwann cells. Stem Cells Transl. Med. 1, 266–278. doi: 10.5966/sctm.2011-0042
Liu, T. Y., Chen, Y. C., Jong, Y. J., Tsai, H. J., Lee, C. C., Chang, Y. S., et al. (2017). Muscle developmental defects in heterogeneous nuclear ribonucleoprotein A1 knockout mice. Open Biol. 7:160303. doi: 10.1098/rsob.160303
López Del Amo, V., Seco-Cervera, M., García-Giménez, J. L., Whitworth, A. J., Pallardó, F. V., and Galindo, M. I. (2015). Mitochondrial defects and neuromuscular degeneration caused by altered expression of Drosophila Gdap1: Implications for the Charcot-Marie-Tooth neuropathy. Hum. Mol. Genet. 24, 21–36. doi: 10.1093/hmg/ddu416
López-Olmeda, J. F., and Sánchez-Vázquez, F. J. (2011). Thermal biology of zebrafish (Danio rerio). J. Therm. Biol. 36, 91–104. doi: 10.1016/j.jtherbio.2010.12.005
Lupski, J. R., de Oca-Luna, R. M., Slaugenhaupt, S., Pentao, L., Guzzetta, V., Trask, B. J., et al. (1991). DNA duplication associated with Charcot-Marie-Tooth disease type 1A. Cell 66, 219–232. doi: 10.1016/0092-8674(91)90613-4
Lyons, D. A., Naylor, S. G., Scholze, A., and Talbot, W. S. (2009). Kif1b is essential for mRNA localization in oligodendrocytes and development of myelinated axons. Nat. Genet. 41, 854–858. doi: 10.1038/ng.376
Lysko, D. E., Meireles, A. M., Folland, C., McNamara, E., Laing, N. G., Lamont, P. J., et al. (2022). Partial loss-of-function variant in neuregulin 1 identified in family with heritable peripheral neuropathy. Hum. Mutat. 43, 1216–1223. doi: 10.1002/humu.24393
Malissovas, N., Griffin, L. B., Antonellis, A., and Beis, D. (2016). Dimerization is required for GARS-mediated neurotoxicity in dominant CMT disease. Hum. Mol. Genet. 25, 1528–1542. doi: 10.1093/hmg/ddw031
Mao, L., Bryantsev, A. L., Chechenova, M. B., and Shelden, E. A. (2005). Cloning, characterization, and heat stress-induced redistribution of a protein homologous to human hsp27 in the zebrafish Danio rerio. Exp. Cell Res. 306, 230–241. doi: 10.1016/j.yexcr.2005.02.007
Marí-Beffa, M., Mesa-Román, A. B., and Duran, I. (2021). Zebrafish models for human skeletal disorders. Front. Genet. 12:675331. doi: 10.3389/fgene.2021.675331
Middleton, R. C., and Shelden, E. A. (2013). Small heat shock protein HSPB1 regulates growth of embryonic zebrafish craniofacial muscles. Exp. Cell Res. 319, 860–874. doi: 10.1016/j.yexcr.2013.01.002
Mullen, P., Abbott, J. A., Wellman, T., Aktar, M., Fjeld, C., Demeler, B., et al. (2021). Neuropathy-associated histidyl-tRNA synthetase variants attenuate protein synthesis in vitro and disrupt axon outgrowth in developing zebrafish. FEBS J. 288, 142–159. doi: 10.1111/febs.15449
Murphy, S. M., Laura, M., Fawcett, K., Pandraud, A., Liu, Y. T., Davidson, G. L., et al. (2012). Charcot-Marie-Tooth disease: Frequency of genetic subtypes and guidelines for genetic testing. J. Neurol. Neurosurg. Psychiatry 83, 706–710. doi: 10.1136/jnnp-2012-302451
Naef, V., Meschini, M. C., Tessa, A., Morani, F., Corsinovi, D., Ogi, A., et al. (2023). Converging Role for REEP1/SPG31 in Oxidative Stress. Int. J. Mol. Sci. 24:3527. doi: 10.3390/ijms24043527
Nilsson, S. (1970). Excitatory and inhibitory innervation of the urinary bladder and gonads of a teleost, Gadus morhua. Comp. Gen. Pharmacol. 1, 23–28. doi: 10.1016/0010-4035(70)90004-2
Okamoto, Y., and Takashima, H. (2023). The current state of charcot-marie-tooth disease treatment. Genes (Basel) 14:1391. doi: 10.3390/genes14071391
Pagnamenta, A. T., Kaiyrzhanov, R., Zou, Y., Da’as, S. I., Maroofian, R., Donkervoort, S., et al. (2021). An ancestral 10-bp repeat expansion in VWA1 causes recessive hereditary motor neuropathy. Brain 144, 584–600. doi: 10.1093/brain/awaa420
Palieri, V., Paoli, E., Wu, Y. K., Haesemeyer, M., Grunwald Kadow, I. C., and Portugues, R. (2024). The preoptic area and dorsal habenula jointly support homeostatic navigation in larval zebrafish. Curr. Biol. 34, 489–504.e7. doi: 10.1016/j.cub.2023.12.030
Pedroni, A., Yilmaz, E., Del Vecchio, L., Bhattarai, P., Vidal, I. T., Dai, Y. E., et al. (2024). Decoding the molecular, cellular, and functional heterogeneity of zebrafish intracardiac nervous system. Nat. Commun. 15:10483. doi: 10.1038/s41467-024-54830-w
Pei, W., Xu, L., Varshney, G. K., Carrington, B., Bishop, K., Jones, M., et al. (2016). Additive reductions in zebrafish PRPS1 activity result in a spectrum of deficiencies modeling several human PRPS1-associated diseases. Sci. Rep. 6:29946. doi: 10.1038/srep29946
Perez-Siles, G., Cutrupi, A., Ellis, M., Screnci, R., Mao, D., Uesugi, M., et al. (2020). Energy metabolism and mitochondrial defects in X-linked Charcot-Marie-Tooth (CMTX6) iPSC-derived motor neurons with the p.R158H PDK3 mutation. Sci. Rep. 10:9262. doi: 10.1038/s41598-020-66266-5
Petel Légaré, V., Rampal, C. J., Gurberg, T. J. N., Aaltonen, M. J., Janer, A., Zinman, L., et al. (2023). Loss of mitochondrial Chchd10 or Chchd2 in zebrafish leads to an ALS-like phenotype and complex I deficiency independent of the mitochondrial integrated stress response. Dev. Neurobiol. 83, 54–69. doi: 10.1002/dneu.22909
Pisciotta, C., and Shy, M. E. (2023). Hereditary neuropathy. Handb. Clin. Neurol. 195, 609–617. doi: 10.1016/B978-0-323-98818-6.00009-1
Pisciotta, C., Saveri, P., and Pareyson, D. (2021). Updated review of therapeutic strategies for Charcot-Marie-Tooth disease and related neuropathies. Exp. Rev. Neurother. 21, 701–713. doi: 10.1080/14737175.2021.1935242
Pogoda, H. M., Sternheim, N., Lyons, D. A., Diamond, B., Hawkins, T. A., Woods, I. G., et al. (2006). A genetic screen identifies genes essential for development of myelinated axons in zebrafish. Dev. Biol. 298, 118–131. doi: 10.1016/j.ydbio.2006.06.021
Ponomareva, O. Y., Eliceiri, K. W., and Halloran, M. C. (2016). Charcot-Marie-Tooth 2b associated Rab7 mutations cause axon growth and guidance defects during vertebrate sensory neuron development. Neural Dev. 11:2. doi: 10.1186/s13064-016-0058-x
Preston, M. A., Finseth, L. T., Bourne, J. N., and Macklin, W. B. (2019). A novel myelin protein zero transgenic zebrafish designed for rapid readout of in vivo myelination. Glia 67, 650–667. doi: 10.1002/glia.23559
Prukop, T., Wernick, S., Boussicault, L., Ewers, D., Jäger, K., Adam, J., et al. (2020). Synergistic PXT3003 therapy uncouples neuromuscular function from dysmyelination in male Charcot-Marie-Tooth disease type 1A (CMT1A) rats. J. Neurosci. Res. 98, 1933–1952. doi: 10.1002/jnr.24679
Qiu, Y., Kenana, R., Beharry, A., Wilhelm, S. D. P., Hsu, S. Y., Siu, V. M., et al. (2023). Histidine supplementation can escalate or rescue HARS deficiency in a Charcot-Marie-Tooth disease model. Hum. Mol. Genet. 32, 810–824. doi: 10.1093/hmg/ddac239
Raeymaekers, P., Timmerman, V., Nelis, E., Van Hul, W., De Jonghe, P., Martin, J. J., et al. (1992). Estimation of the size of the chromosome 17p11.2 duplication in Charcot-Marie-Tooth neuropathy type 1a (CMT1a). HMSN collaborative research group. J. Med. Genet. 29, 5–11. doi: 10.1136/jmg.29.1.5
Ramesh, T., Lyon, A. N., Pineda, R. H., Wang, C., Janssen, P. M., Canan, B. D., et al. (2010). A genetic model of amyotrophic lateral sclerosis in zebrafish displays phenotypic hallmarks of motoneuron disease. Dis. Model Mech. 3, 652–662. doi: 10.1242/dmm.005538
Ran, G. L., Li, Y. P., Lu, L. C., and Lan, S. H. (2024). Disease-modifying therapies for diabetic peripheral neuropathy: A systematic review and meta-analysis of randomized controlled trials. J. Diab. Complicat. 38:108691. doi: 10.1016/j.jdiacomp.2024.108691
Richards, S., Aziz, N., Bale, S., Bick, D., Das, S., Gastier-Foster, J., et al. (2015). Standards and guidelines for the interpretation of sequence variants: A joint consensus recommendation of the American college of medical genetics and genomics and the association for molecular pathology. Genet. Med. 17, 405–424. doi: 10.1038/gim.2015.30
Rzepnikowska, W., and Kochański, A. (2018). A role for the GDAP1 gene in the molecular pathogenesis of Charcot-Marie-Tooth disease. Acta Neurobiol. Exp. (Wars) 78, 1–13.
Rzepnikowska, W., Kaminska, J., and Kochański, A. (2022). Validation of the pathogenic effect of IGHMBP2 gene mutations based on yeast s. cerevisiae model. Int. J. Mol. Sci. 23:9913. doi: 10.3390/ijms23179913
Rzepnikowska, W., Kaminska, J., Kabzińska, D., Binięda, K., and Kochański, A. (2020a). A yeast-based model for hereditary motor and sensory neuropathies: A simple system for complex. Heterogeneous Diseases. Int. J. Mol. Sci. 21:4277. doi: 10.3390/ijms21124277
Rzepnikowska, W., Kaminska, J., Kabzińska, D., and Kochański, A. (2020b). Pathogenic effect of GDAP1 gene mutations in a yeast model. Genes (Basel) 11:310. doi: 10.3390/genes11030310
Sahenk, Z., and Ozes, B. (2020). Gene therapy to promote regeneration in Charcot-Marie-Tooth disease. Brain Res. 1727:146533. doi: 10.1016/j.brainres.2019.146533
Saporta, M. A., Dang, V., Volfson, D., Zou, B., Xie, X. S., Adebola, A., et al. (2015). Axonal Charcot-Marie-Tooth disease patient-derived motor neurons demonstrate disease-specific phenotypes including abnormal electrophysiological properties. Exp. Neurol. 263, 190–199. doi: 10.1016/j.expneurol.2014.10.005
Sato, T., Takahoko, M., and Okamoto, H. (2006). HuC:kaede, a useful tool to label neural morphologies in networks in vivo. Genesis 44, 136–142. doi: 10.1002/gene.20196
Saxena, A., Peng, B. N., and Bronner, M. E. (2013). Sox10-dependent neural crest origin of olfactory microvillous neurons in zebrafish. Elife 2:e00336. doi: 10.7554/eLife.00336
Schabhüttl, M., Wieland, T., Senderek, J., Baets, J., Timmerman, V., De Jonghe, P., et al. (2014). Whole-exome sequencing in patients with inherited neuropathies: Outcome and challenges. J. Neurol. 261, 970–982. doi: 10.1007/s00415-014-7289-8
Schonkeren, S. L., Massen, M., van der Horst, R., Koch, A., Vaes, N., and Melotte, V. (2019). Nervous NDRGs: The N-myc downstream-regulated gene family in the central and peripheral nervous system. Neurogenetics 20, 173–186. doi: 10.1007/s10048-019-00587-0
Schwartzlow, C., and Kazamel, M. (2019). Hereditary sensory and autonomic neuropathies: Adding more to the classification. Curr. Neurol. Neurosci. Rep. 19:52. doi: 10.1007/s11910-019-0974-3
Shrimpton, A. E., Levinsohn, E. M., Yozawitz, J. M., Packard, D. S., Cady, R. B., Middleton, F. A., et al. (2004). A HOX gene mutation in a family with isolated congenital vertical talus and Charcot-Marie-Tooth disease. Am. J. Hum. Genet. 75, 92–96. doi: 10.1086/422015
Silbernagel, N., Walecki, M., Schäfer, M. K., Kessler, M., Zobeiri, M., Rinné, S., et al. (2018). The VAMP-associated protein VAPB is required for cardiac and neuronal pacemaker channel function. FASEB J. 32, 6159–6173. doi: 10.1096/fj.201800246R
Singh, J., and Patten, S. A. (2022). Modeling neuromuscular diseases in zebrafish. Front. Mol. Neurosci. 15:1054573. doi: 10.3389/fnmol.2022.1054573
Skedsmo, F. S., Tranulis, M. A., Espenes, A., Prydz, K., Matiasek, K., Gunnes, G., et al. (2019). Cell and context-dependent sorting of neuropathy-associated protein NDRG1 - insights from canine tissues and primary Schwann cell cultures. BMC Vet. Res. 15:121. doi: 10.1186/s12917-019-1872-2
Skre, H. (1974). Genetic and clinical aspects of Charcot-Marie-Tooth’s disease. Clin. Genet. 6, 98–118. doi: 10.1111/j.1399-0004.1974.tb00638.x
Soh, M. S., Cheng, X., Vijayaraghavan, T., Vernon, A., Liu, J., and Neumann, B. (2020). Disruption of genes associated with Charcot-Marie-Tooth type 2 lead to common behavioural, cellular and molecular defects in Caenorhabditis elegans. PLoS One 15:e0231600. doi: 10.1371/journal.pone.0231600
Son, W., Jeong, H. S., Nam, D. E., Lee, A. J., Nam, S. H., Lee, J. E., et al. (2023). Peripheral neuropathy and decreased locomotion of a RAB40B mutation in human and model animals. Exp. Neurobiol. 32, 410–422. doi: 10.5607/en23027
Stainier, D. Y. R., Raz, E., Lawson, N. D., Ekker, S. C., Burdine, R. D., Eisen, J. S., et al. (2017). Guidelines for morpholino use in zebrafish. PLoS Genet. 13:e1007000. doi: 10.1371/journal.pgen.1007000
Stavrou, M., Kagiava, A., Sargiannidou, I., Georgiou, E., and Kleopa, K. A. (2023). Charcot-Marie-Tooth neuropathies: Current gene therapy advances and the route toward translation. J. Peripher. Nerv. Syst. 28, 150–168. doi: 10.1111/jns.12543
Stavrou, M., Sargiannidou, I., Georgiou, E., Kagiava, A., and Kleopa, K. A. (2021). Emerging therapies for charcot-marie-tooth inherited neuropathies. Int. J. Mol. Sci. 22:6048. doi: 10.3390/ijms22116048
Tazir, M., and Nouioua, S. (2024). Distal hereditary motor neuropathies. Rev. Neurol. 180, 1031–1036. doi: 10.1016/j.neurol.2023.09.005
Tesoriero, C., Greco, F., Cannone, E., Ghirotto, F., Facchinello, N., Schiavone, M., et al. (2023). Modeling human muscular dystrophies in zebrafish: Mutant lines, transgenic fluorescent biosensors, and phenotyping assays. Int. J. Mol. Sci. 24:8314. doi: 10.3390/ijms24098314
Toni, M., Arena, C., Cioni, C., and Tedeschi, G. (2023). Temperature- and chemical-induced neurotoxicity in zebrafish. Front. Physiol. 14:1276941. doi: 10.3389/fphys.2023.1276941
Van Hul, W., Winkler, C., Laurent, M. R., Tonelli, F., Willem Bek, J., Besio, R., et al. (2020). Zebrafish: A resourceful vertebrate model to investigate skeletal disorders. Front. Endocrinol. 11:489. doi: 10.3389/fendo.2020.00489
Van Lent, J., Vendredy, L., Adriaenssens, E., Da Silva Authier, T., Asselbergh, B., Kaji, M., et al. (2022). Downregulation of PMP22 ameliorates myelin defects in iPSC-derived human organoid cultures of CMT1A. Brain 146, 2885–2896. doi: 10.1093/brain/awac475
Van Lent, J., Verstraelen, P., Asselbergh, B., Adriaenssens, E., Mateiu, L., Verbist, C., et al. (2021). Induced pluripotent stem cell-derived motor neurons of CMT type 2 patients reveal progressive mitochondrial dysfunction. Brain 144, 2471–2485. doi: 10.1093/brain/awab226
Vettori, A., Bergamin, G., Moro, E., Vazza, G., Polo, G., Tiso, N., et al. (2011). Developmental defects and neuromuscular alterations due to mitofusin 2 gene (MFN2) silencing in zebrafish: A new model for Charcot-Marie-Tooth type 2A neuropathy. Neuromuscul. Disord. 21, 58–67. doi: 10.1016/j.nmd.2010.09.002
Vona, B., Doll, J., Hofrichter, M. A. H., Haaf, T., and Varshney, G. K. (2020). Small fish, big prospects: Using zebrafish to unravel the mechanisms of hereditary hearing loss. Hear. Res. 397:107906. doi: 10.1016/j.heares.2020.107906
Waldron, A. L., Cahan, S. H., Francklyn, C. S., and Ebert, A. M. (2017). A single Danio rerio hars gene encodes both cytoplasmic and mitochondrial histidyl-tRNA synthetases. PLoS One 12:e0185317. doi: 10.1371/journal.pone.0185317
Weterman, M. A. J., Kuo, M., Kenter, S. B., Gordillo, S., Karjosukarso, D. W., Takase, R., et al. (2018). Hypermorphic and hypomorphic AARS alleles in patients with CMT2N expand clinical and molecular heterogeneities. Hum. Mol. Genet. 27, 4036–4050. doi: 10.1093/hmg/ddy290
Weterman, M. A., Sorrentino, V., Kasher, P. R., Jakobs, M. E., van Engelen, B. G., Fluiter, K., et al. (2012). A frameshift mutation in LRSAM1 is responsible for a dominant hereditary polyneuropathy. Hum. Mol. Genet. 21, 358–370. doi: 10.1093/hmg/ddr471
Whitfield, T. T. (2002). Zebrafish as a model for hearing and deafness. J. Neurobiol. 53, 157–171. doi: 10.1002/neu.10123
Won, S. Y., Kwon, S., Jeong, H. S., Chung, K. W., Choi, B. O., Chang, J. W., et al. (2020). Fibulin 5, a human Wharton’s jelly-derived mesenchymal stem cells-secreted paracrine factor, attenuates peripheral nervous system myelination defects through the Integrin-RAC1 signaling axis. Stem Cells 38, 1578–1593. doi: 10.1002/stem.3287
Xiao, Y., Faucherre, A., Pola-Morell, L., Heddleston, J. M., Liu, T. L., Chew, T. L., et al. (2015). High-resolution live imaging reveals axon-glia interactions during peripheral nerve injury and repair in zebrafish. Dis. Model Mech. 8, 553–564. doi: 10.1242/dmm.018184
Yousefi, F., Lavi Arab, F., Nikkhah, K., Amiri, H., and Mahmoudi, M. (2019). Novel approaches using mesenchymal stem cells for curing peripheral nerve injuries. Life Sci. 221, 99–108. doi: 10.1016/j.lfs.2019.01.052
Zhao, M., Smith, L., Volpatti, J., Fabian, L., and Dowling, J. J. (2019). Insights into wild-type dynamin 2 and the consequences of DNM2 mutations from transgenic zebrafish. Hum. Mol. Genet. 28, 4186–4196. doi: 10.1093/hmg/ddz260
Zhou, W., Hsu, A. Y., Wang, Y., Syahirah, R., Wang, T., Jeffries, J., et al. (2020). Mitofusin 2 regulates neutrophil adhesive migration and the actin cytoskeleton. J. Cell Sci. 133:jcs248880. doi: 10.1242/jcs.248880
Zhu, Y., Lobato, A. G., Rebelo, A. P., Canic, T., Ortiz-Vega, N., Tao, X., et al. (2023). Sorbitol reduction via govorestat ameliorates synaptic dysfunction and neurodegeneration in sorbitol dehydrogenase deficiency. JCI Insight 8:e164954. doi: 10.1172/jci.insight.164954
Keywords: Charcot-Marie-Tooth disease, CMT, neuropathy, animal models, zebrafish
Citation: Korzeniowska née Wiweger M, Chabros K, Rzepnikowska W, Kochański A and Kabzińska D (2025) Modeling of Charcot-Marie-Tooth disease in zebrafish. Front. Mol. Neurosci. 18:1641793. doi: 10.3389/fnmol.2025.1641793
Received: 05 June 2025; Accepted: 11 July 2025;
Published: 04 August 2025.
Edited by:
Stephan C. F. Neuhauss, University of Zurich, SwitzerlandReviewed by:
Matthias Carl, University of Trento, ItalyMatthias Gesemann, University of Zurich, Switzerland
Copyright © 2025 Korzeniowska née Wiweger, Chabros, Rzepnikowska, Kochański and Kabzińska. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Dagmara Kabzińska, ZGFna2FiQGltZGlrLnBhbi5wbA==
†These authors have contributed equally to this work and share first authorship