Gene therapy approaches targeting Schwann cells for demyelinating neuropathies

Journal Pre-proofs Review Gene therapy approaches targeting Schwann cells for demyelinating neuropathies Irene Sargiannidou, Alexia Kagiava, Kleopas A. Kleopa PII: DOI: Reference:

S0006-8993(19)30626-2 https://doi.org/10.1016/j.brainres.2019.146572 BRES 146572

To appear in:

Brain Research

Received Date: Revised Date: Accepted Date:

15 July 2019 12 November 2019 26 November 2019

Please cite this article as: I. Sargiannidou, A. Kagiava, K.A. Kleopa, Gene therapy approaches targeting Schwann cells for demyelinating neuropathies, Brain Research (2019), doi: https://doi.org/10.1016/j.brainres.2019.146572

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Special Issue of Brain Research devoted to Charcot-Marie-Tooth disease

Gene therapy approaches targeting Schwann cells for demyelinating neuropathies

Irene Sargiannidou,a Alexia Kagiava, a Kleopas A. Kleopa a,b,*

aNeuroscience

Laboratory and bNeurology Clinics, The Cyprus Institute of Neurology and

Genetics and Cyprus School of Molecular Medicine, Nicosia, Cyprus

Words in Abstract: 235 Words total: 5729 Table: 1 Figure: 1 Number of References: 148 *Correspondence:

Prof. Kleopas A. Kleopa, MD The Cyprus Institute of Neurology and Genetics 6 International Airport Avenue, P.O. Box 23462, 1683, Nicosia, CYPRUS +357 22 358600 +357 22 392786 [email protected]

Acknowledgements: Work in the author’s laboratory related to this topic has been funded by the Muscular Dystrophy Association (MDA grants 277250, 480030 and 603003), by the Charcot-Marie-Tooth Association, and by AFM-Telethon (Grant 19719).

Highlights: 

Demyelinating CMT results from mutations in Schwann cell genes



Intrathecal vector delivery could provide a translatable access to PNS



Schwann cell-targeted gene expression using myelin-specific promoters



Encouraging gene replacement studies in CMT1X and CMT4C models

1

Abstract Charcot-Marie-Tooth disease (CMT) encompasses numerous genetically heterogeneous inherited neuropathies, which together are one of the commonest neurogenetic disorders. Axonal CMT types result from mutations in neuronally expressed genes, whereas demyelinating CMT forms mostly result from mutations in genes expressed by myelinating Schwann cells. The demyelinating forms are the most common, and may be caused by dominant mutations and gene dosage effects (as in CMT1), as well as by recessive mutations and loss of function mechanisms (as in CMT4). The discovery of causative genes and increasing insights into molecular mechanisms through the study of experimental disease models has provided the basis for the development of gene therapy approaches. For demyelinating CMT, gene silencing or gene replacement strategies need to be targeted to Schwann cells. Progress in gene replacement for two different CMT forms, including CMT1X caused by GJB1 gene mutations, and CMT4C, caused by SH3TC2 gene mutations, has been made through the use of a myelin-specific promoter to restrict expression in Schwann cells, and by lumbar intrathecal delivery of lentiviral viral vectors to achieve more widespread biodistribution in the peripheral nervous system. This review summarizes the molecular-genetic mechanisms of selected demyelinating CMT neuropathies and the progress made so far, as well as the remaining challenges in the path towards a gene therapy to treat these disorders through the use of optimal gene therapy tools such as using clinically translatable delivery methods and adeno-associated viral (AAV) vectors.

Keywords: Charcot-Marie-Tooth disease, Schwann cells, axons, myelin, viral vectors, gene replacement, gene silencing, gene editing

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1. Introduction Non-syndromic inherited peripheral neuropathies, collectively known as Charcot-Marie-Tooth (CMT) disease, include numerous types and when combined, they represent one of the most common neurogenetic disorders, with a prevalence of 1:2500 worldwide (Baets et al., 2014; Kleopa and Scherer, 2002). CMT neuropathies are genetically heterogeneous, associated with mutations in over 100 different genes, affecting diverse cellular functions. While mutations in the same gene may be associated with variable phenotypes, in most cases mutations in genes expressed by neurons result in axonal neuropathy, whereas mutations in genes expressed in myelinating Schwann cells lead to demyelinating types of neuropathy through cell autonomous effects (Rossor et al., 2013; Scherer and Wrabetz, 2008; Scherer et al., 2015; Suter and Scherer, 2003). Updated and detailed listing of CMT genes and associated mutations with phenotypes can be found in the Online Mendelian Inheritance in Man

(OMIM;

http://www.ncbi.nlm.nih.gov/omim),

in

the

Gene

Reviews

(http://www.ncbi.nlm.nih.gov/books/NBK1358/), and in the inherited neuropathy variant browser (http://hihg.med.miami.edu/code/http/cmt/public_html/index.html#/). Despite the increasing understanding of the complex genetic basis and diverse cellular mechanisms underpinning CMT neuropathies, there is currently no effective treatment for any of the CMT forms. Only symptomatic and supportive therapy can be offered to patients. Thus, there is a great need for new treatment strategies for CMT. In the last two decades there has been an effort to develop gene therapies for the treatment of CMT, with several preclinical studies providing proof-of-concept that certain therapeutic approaches have the potential to reach clinical translation. Recent advances in the development of gene therapy for other neuromuscular diseases, notably clinical trials for giant axonal neuropathy (GAN) and spinal muscular atrophy (SMA) have provided further impetus towards gene therapy studies for CMT neuropathies as well. Furthermore, recent preclinical studies of gene therapy for certain axonal CMT neuropathies have shown success (Morelli et al., 2019; Shababi et al., 2016). While these gene therapy approaches hold promise for the future to treat diseases of the central and peripheral nervous system (PNS), multiple challenges remain to be overcome along this effort. In this review we will discuss recent advances towards cell-targeted gene therapy specifically for representative demyelinating CMT forms caused by genes expressed in Schwann cells, and will highlight further steps needed for the development of translatable therapeutic approaches. 2. Molecular mechanisms in demyelinating Charcot-Marie-Tooth (CMT) neuropathies Several types of autosomal dominant, X-linked, and autosomal recessive types of demyelinating neuropathies have been identified (Bird, 1993; Fridman et al., 2015; Murphy 3

et al., 2012; Saporta et al., 2011; Scherer et al., 2015) (Table 1). Dominantly inherited types (CMT1) are the most common. They were initially classified based on upper limb motor nerve conduction velocities (NCVs) below 38 m/s, and segmental demyelination and remyelination with onion bulb formation in nerve biopsies (Scherer et al., 2015; Shy et al., 2005). CMT1X and some cases of CMT1B show mixed axonal and demyelinating features with intermediate slowing of NCVs. Recessive demyelinating CMT forms grouped under the name CMT4 are much rarer than dominant ones even within multi-center international databases (Fridman et al., 2015). Most of the genes involved in demyelinating CMT forms, regardless of the mode of inheritance, are expressed predominantly or exclusively by myelinating Schwann cells. They serve a variety of cellular functions, including myelin formation and metabolism, signaling, endosomal sorting, gene expression regulation, mitochondrial dynamics, stability of the cytoskeleton and basal membrane adhesion. Experimental models have provided insights into the cellular mechanisms in demyelinating CMT neuropathies indicating that mutations cause either a gain-of-function in CMT1 or a loss-of-function in CMT1X and CMT4 patients (Azzedine et al., 1993; Rotthier et al., 2012; Sargiannidou et al., 2009; Scherer and Wrabetz, 2008). Below we focus on clinical features and cellular mechanisms of two representative forms of demyelinating CMT neuropathies, CMT1X and CMT4C, as a basis for developing gene therapy approaches. 2.1.

X-linked CMT type 1 (CMT1X) and Connexin32 mutations

The X-linked CMT type 1 (CMT1X) is the second most common CMT form worldwide and results from mutations in the GJB1 gene, which encodes the gap junction (GJ) protein Cx32 (Bergoffen et al., 1993; Kleopa and Scherer, 2006). Cx32 is expressed by myelinating Schwann cells and other cell types (Chandross et al., 1996; Ressot and Bruzzone, 2000; Scherer et al., 1995). GJ channels are formed by two hexamers (hemichannels) of individual connexin molecules arranged around a central pore. Cx32 GJ channels are localized in noncompact myelin areas including the paranodal myelin loops and Schmidt-Lantermann incisures (Scherer et al., 1995). Here, they play an important role in preserving the homeostasis of the myelin sheath and the axon, transferring ions, metabolites and second messengers (Bortolozzi, 2018; Bruzzone et al., 1996; Kleopa et al., 2012; Meier et al., 2004). The clinical phenotype of CMT1X includes peripheral neuropathy usually as the only clinical manifestation, with progressive weakness and atrophy starting in distal leg muscles, difficulty running and frequently sprained ankles, with onset by 10 years of age or earlier in most affected males (Birouk et al., 1998; Hahn et al., 2000). The disease is slowly progressive 4

causing weakness of foreleg muscles, foot drop, foot deformities, hand muscle weakness, and distal sensory loss with sometimes painful paresthesias. Heterozygous females with CMT1X may be asymptomatic or they may develop milder clinical manifestations at an older age (Jerath et al., 2016). Intermediate slowing (30-40 m/s) of motor NCVs and progressive loss

of

motor

units

due

to

length-dependent

axonal

degeneration

are

typical

electrophysiological features (Hahn et al., 1990). Nerve biopsies show mixed axonal and demyelinating abnormalities (Hahn et al., 2001; Hattori et al., 2003) with thin myelin sheaths and loss of large myelinated fibers replaced by regenerating axon clusters (Kleopa et al., 2006). Transient CNS manifestations have been described in some, mostly younger CMT1X patients (Al-Mateen et al., 2014; Sargiannidou et al., 2015b), and consist of mild chronic or more dramatic but transient encephalopathy syndromes typically triggered by conditions of metabolic stress (Kleopa, 2011; Paulson et al., 2002; Taylor et al., 2003). More recent studies of large CMT1X cohorts indicate that the overall frequency of CNS phenotypes is less than 10% among patients with different coding and non-coding GJB1 mutations (Vivekanandam et al., 2018; Yuan et al., 2018). More than 400 GJB1 mutations have been reported to date occurring throughout the open reading frame (ORF), many of which in more than one families, including: 498 missense (71%); 3 stop-lost; 49 Inframe INDELs (7%); 25 Stop-Gained (4%); and 122 Frameshift INDELs (17%) (http://hihg.med.miami.edu/code/http/cmt/public_html/index.html#/). Several mutations have also been discovered in non-coding GJB1 regions, including the promoter and 3’-end region (Murphy et al., 2011; Tomaselli et al., 2017). Frameshift, premature stop, and non-coding mutations are likely to cause complete loss of protein synthesis or rapid degradation, precluding any dominant-negative effects. Several missense and in-frame mutations expressed in vitro showed intracellular retention (Omori et al., 1996; Yoshimura et al., 1998; Yum et al., 2002) in the ER and/or Golgi (Deschênes et al., 1997; Kleopa et al., 2002; Martin et al., 2000; Oh et al., 1997; Yum et al., 2002) with failure to form functional GJ channels. Retained Cx32 mutants do not cause aggregates and are efficiently degraded via lysosomal and proteasomal pathways (Kleopa et al., 2002; VanSlyke et al., 2000). Some mutants however exerted dominant-negative effects on co-expressed wild type (WT) Cx32 (Kyriakoudi et al., 2017; Omori et al., 1996). Other mutants reached the cell membrane and formed functional channels with altered biophysical characteristics (Oh et al., 1997). Cx32 knockout (KO) mice with deletion of the Gjb1 gene develop a progressive, predominantly motor demyelinating peripheral neuropathy beginning at about three months 5

of age with reduced sciatic motor NCV and motor amplitude (Anzini et al., 1997; Scherer et al., 1998). Early axonal pathology evident in dysregulated neurofilament phosphorylation and slowing of axonal transport has been shown to even precede demyelination in Cx32 KO mice (Vavlitou et al., 2010), in keeping with early axonal abnormalities described in nerve biopsies from CMT1X patients as well (Hahn et al., 2000; Hahn et al., 2001; Hattori et al., 2003). Furthermore, increased inflammatory responses have been demonstrated in the PNS of Cx32 KO mice with dysregulated cytokine colony stimulating factor-1 (CSF-1) and increased macrophage infiltrates that may drive further nerve pathology (Groh et al., 2010; Groh et al., 2016; Kobsar et al., 2002; Kobsar et al., 2003). Thus, loss of Cx32 GJ channels in non-compact myelin appears to affect the homeostasis of both myelinating Schwann cells and the axon simultaneously (Kleopa, 2011). Expression of WT human Cx32 protein driven by the rat myelin protein zero (Mpz/P0) promoter prevented demyelination in Cx32 KO mice (Scherer et al., 2005), confirming that loss of Schwann cell autonomous expression of Cx32 is sufficient to cause CMT1X pathology. Transgenic mice expressing CMT1X mutations showed no detectable Cx32 protein in the 175fs mutant line (Abel et al., 1999), while R142W, T55I, R75W and N175D transgenic mice showed retention of the mutant protein in the perinuclear region, similar to in vitro expression pattern, and developed a demyelinating neuropathy similar to Cx32 KO mice (Jeng et al., 2006; Kagiava et al., 2018; Sargiannidou et al., 2009). In the presence of the Golgi-retained R142W, R75W and N175D mutants (but not of the ER-retained T55I mutant), there was reduced expression of the endogenous mouse WT Cx32, indicating that Golgi-retained mutants may have dominant-negative effects on WT Cx32. This is not clinically relevant for CMT1X patients expressing only one GJB1 allele in each cell, but must be considered when planning a gene addition therapy. None of the mutants expressed in vivo had any other toxic or dominant effects on other co-expressed connexins in Schwann cells or oligodendrocytes (Jeng et al., 2006; Sargiannidou et al., 2009). The C-terminus mutants C280G and S281X were properly localized and prevented demyelination in Cx32 KO mice, so that it remains unclear how they cause neuropathy in humans (Huang et al., 2005). Overall, results from animal models of CMT1X (Sargiannidou et al., 2009; Scherer et al., 1998; Scherer et al., 2005) have demonstrated that in most cases the neuropathy results from cell-autonomous loss of function of Cx32 in Schwann cells. In addition, recent studies of large CMT1X patient cohorts with different GJB1 mutations showed that disability increases with age, and is comparable with that observed in patients with a complete GJB1 deletion (Panosyan et al., 2017; Shy et al., 2007). Likewise, the degree of pathology in 6

CMT1X nerve biopsies is not associated with particular GJB1 mutations (Hahn et al., 2000; Hattori et al., 2003). Taken together, these findings suggest that most GJB1 mutations cause loss of Cx32 function. Therefore, cell-targeted gene replacement strategies delivering the normal GJB1 gene may have the potential to treat the disease. 2.2. Charcot-Marie-Tooth type 4C (CMT4C) disease CMT4C is an autosomal recessive inherited neuropathy that appears to be the most prevalent among the overall rare recessive demyelinating CMT4 forms, being responsible for almost half of all CMT4 cases (Azzedine et al., 1993; Fridman et al., 2015; Piscosquito et al., 2016). Patients with CMT4C usually present in the first decade of life with foot deformities and scoliosis, weakness, areflexia and sensory loss (Azzedine et al., 2006; Gabreels-Festen et al., 1999; Kessali et al., 1997). Cranial nerve involvement with hearing impairment, trigeminal neuralgia, slow pupillary light reflexes, and lingual fasciculation are common and phenotypic variations in patients with identical mutations have been described (Colomer et al., 2006; Gooding et al., 2005; Kontogeorgiou et al., 2019; Piscosquito et al., 2016; Varley et al., 2015). Electrophysiological studies in CMT4C patients confirm the demyelinating process with mean median motor NCV at 22.6 m/s. Nerve biopsy findings are characterized by an increase of basal membranes around myelinated, demyelinated, and unmyelinated axons, relatively few onion bulbs, and, most typically, large cytoplasmic extensions of Schwann cells (Gabreels-Festen et al., 1999; Kessali et al., 1997; Senderek et al., 2003; Yger et al., 2012). Linkage analysis studies and homozygosity mapping (LeGuern et al., 1996) led to the discovery of the disease locus on chromosome 5q32 and subsequently to the initial discovery of 11 different mutations in the SH3TC2 gene, mostly truncating but also missense (Senderek et al., 2003). At least 100 different SH3TC2 mutations have been described to date. Certain mutations are more common among certain ethnic groups (Claramunt et al., 2007; Lassuthova et al., 2011) with likely founder effects (Gooding et al., 2005). The full transcript cDNA length measures 3864 bp. SH3TC2 encodes a protein of 1,288 aa containing two Src homology 3 (SH3) and 10 tetratricopeptide repeat (TPR) domains sharing no overall significant similarity to any other human protein with known function. The presence of SH3 and TPR domains suggests that SH3TC2 could act as a scaffold protein (Senderek et al., 2003). SH3TC2 is well conserved among vertebrate species, whereas no non-vertebrate orthologs were identified. SH3TC2 is present in several components of the endocytic pathway including early and late endosomes, and clathrin-coated vesicles close to the trans-Golgi network and in the plasma membrane. This localization is altered in CMT4C

7

(Lupo et al., 2009) with mistargeting of SH3TC2 away from the recycling endosome (Roberts et al., 2010). The Sh3tc2-/- mouse model of CMT4C develops an early onset but progressive peripheral neuropathy with slowing of motor and sensory NCVs and early onset hypomyelination (Arnaud et al., 2009; Gouttenoire et al., 2013). There is increasing myelin pathology from 2 and until 12 months of age. Murine Sh3tc2 is specifically expressed in Schwann cells and is localized to the plasma membrane and to the perinuclear endocytic recycling compartment, suggesting a possible function in myelination and/or in regions of axoglial interactions. Ultrastructural analysis of myelin in the peripheral nerve of mutant mice showed abnormal organization of the node of Ranvier, a phenotype that was confirmed in nerve biopsies from CMT4C patients. These findings suggested a role for SH3TC2 not only in myelination but also in the formation and integrity of the node of Ranvier (Arnaud et al., 2009). Although axonal degeneration was not observed in this model, altered expression of extracellular matrix proteins important for neuromuscular junction (NMJ) integrity and post-synaptic fragmentation of NMJs has been found at an older age (Cipriani et al., 2018). Thus, the Sh3tc2-/- mouse recapitulates all major features of CMT4C disease and provides a relevant model to test therapies. Furthermore, both the inheritance pattern, as well as the insights obtained from in vitro and in vivo disease models indicate that a Schwann cell autonomous loss of SH3TC2 function is the cause of neuropathy in CMT4C. 3. Design of Schwann cell-targeted gene delivery approach As outlined above, most demyelinating forms for CMT result from mutations in genes expressed either exclusively, or predominantly in myelinating Schwann cells. Even when they are also expressed in other tissues, such as the CNS (Cx32 and SH3TC2), disease manifestations originate mainly from their loss of function in Schwann cells. Therefore, targeting genetic therapies to Schwann cells appears to be the optimal strategy in order to minimize off-target toxicity and unpredictable effects in other cell types from ectopic gene expression. Cell targeted expression can be achieved either by using viral vectors with selective tropism for a specific cell type, or by restricting expression through the use of cellspecific promoters to drive gene expression. 3.1. Viral vectors to target Schwann cells for gene therapy The most commonly used viral vectors for gene therapy applications are lentiviral and adeno-associated viral (AAV) vectors. Lentiviral vectors offer several advantages, as they have a large capacity for transgene transfer close to 8kb, they provide a stable and longlasting gene expression through integration into the host genome, and cause no significant 8

immunogenicity (Consiglio et al., 2001; Lattanzi et al., 2010; Lundberg et al., 2008; Naldini et al., 1996). Moreover, they have a demonstrated high tropism for post-mitotic cells including Schwann cells, both in vitro (Hoyng et al., 2015) and in vivo (Kagiava et al., 2016; Sargiannidou et al., 2015a). Therefore, many of the studies described below were based on lentiviral vector gene delivery (Kagiava et al., 2016; Sargiannidou et al., 2015a; Schiza et al., 2019). However, relatively low expression levels achieved resulted in loss of therapeutic effect in certain CMT1X models expressing interfering Cx32 mutants (Kagiava et al., 2018). Most importantly, safety concerns stemming from the random integration into the host genome and even minimal risk of insertional mutagenesis (Hacein-Bey-Abina et al., 2003; Hacein-Bey-Abina et al., 2008; McCormack and Rabbitts, 2004; Somia and Verma, 2000) limit the potential for widely using lentiviral vectors in direct in vivo gene delivery for human trials. AAV vectors have emerged as valuable tools for gene therapy for neurological disorders because of extensive biodistribution with in vivo delivery, lack of integration into the host genome, and stable expression (Choi et al., 2006; Gao et al., 2002; Powell et al., 2015). Therefore, they are currently the vector of choice for direct in vivo clinical applications (Kaplitt et al., 1994). Moreover, AAV vectors could provide higher expression levels (Foust et al., 2009; Gurda et al., 2016; Tanguy et al., 2015), also when compared to lentiviral vectors (Doherty et al., 2011; Powell et al., 2015), while offering the advantage of improved safety with direct in vivo delivery because of their low immunogenicity (Calcedo and Wilson, 2013; Lentz et al., 2012). However, AAV serotypes may show variable tropism for different target cells. For applications to treat disorders with cell autonomous mechanisms, including the demyelinating CMT neuropathies, high tropism for Schwann cells is needed. Most AAV used are pseudotyped in preclinical and clinical gene therapy assays. Different AAV serotypes are used for targeted gene delivery in the nervous system but have been mainly tested in the CNS and their relative tropism for Schwann cells in vivo has not been studied. AAV1/2, 9, and rh10 are the most common serotypes used for gene therapy while new serotypes are introduced like the AAV/DJ, DJ8 or AAV.PHP and even some pseudotyped viruses such as the AAV2/rh10. Recent studies show that AAV9 can easily pass the blood brain barrier (ΒΒΒ) with a wide distribution in the CNS showing high tropism for neurons and astrocytes (Foust et al., 2009; Jackson et al., 2015), while intrathecal delivery can evade the effects of possible neutralizing antibodies (Gray et al., 2013). Moreover, AAV9 is the most commonly used AAV vector in clinical trials (below).

9

Although in vitro effectiveness of AAV vectors does not fully reflect their in vivo effects, Schwann cell specificity of different AAV serotypes has been tested in vitro in cultured rat and human Schwann cells. These experiments indicated that different AAV serotypes may show variation in their tropism between Schwann cells from different species (Bai et al., 2019; Hoyng et al., 2015). Thus, findings in rodent Schwann cells may need to be replicated in human Schwann cells to confirm adequate tropism. However, testing AAV tropism in cell cultures may not be significant as the cells are not differentiated but dividing without control and lack the structure and differentiation of the nerve tissues. Furthermore, model and species-restricted tropism patterns have been described for AAV serotypes also in vivo (Homs et al., 2011; Hordeaux et al., 2018). It remains to be shown whether AAV-mediated gene therapies can adequately target human Schwann cells to treat demyelinating CMT neuropathies. 3.2. Schwann cell-specific promoters Ubiquitous promoters that are commonly used in gene therapy studies may show variable expression patterns, as well as targeting of expression to specific cell populations (Gray et al., 2011). These promoters have not been studied specifically for Schwann cell expression. Alternatively, cell specific promoters, typically of myelin-related genes, may be used to drive gene expression selectively in myelinating cells. Owing to common transcription binding elements, several promoters can drive expression in myelinating cells both in the CNS and PNS. The full length myelin basic protein (MBP) promoter can drive expression in both cell types while shorter elements express only in oligodendrocytes (Farhadi et al., 2003; Forghani et al., 2001). This has been applied to drive viral vector expression in oligodendrocytes with the shorter 1.3-1.9 kb MBP promoter elements (Georgiou et al., 2017; von Jonquieres et al., 2013). However, using the full-length MBP promoter in viral vectors to target Schwann cells is not feasible. Different segments (2.2, 1.5 and 0.3 kb) of the myelin associated glycoprotein (MAG) promoter (von Bartheld et al., 1996; von Jonquieres et al., 2016) have been tested with an AAV vector and all showed strong expression in oligodendrocytes. Although likely, expression in Schwann cells has not been examined with these promoter elements. The 2,3-cyclic nucleotide 3-phosphodiesterase (CNP) (Kagiava et al., 2014; Sargiannidou et al., 2009) and the proteolipid protein (PLP) (Schiza et al., 2015) promoters have been shown to express transgenically in both cell types, while the CNP promoter was also applied in a lentiviral vector to express in oligodendrocytes (Kagiava et al., 2014). Their large size however precludes application in relevant viral vectors. The promoter which has a relatively short length allowing packaging into viral vectors, as well as high selectivity for Schwann cells is the myelin-specific myelin protein zero (Mpz) promoter (Messing et al., 1992). This promoter was shown to drive transgenically high-level 10

expression of Cx32 in Schwann cells of Cx32 KO mice, leading to a complete rescue of the phenotype in this model of CMT1X (Scherer et al., 2005). Furthermore, it was shown both in vitro and in vivo to drive viral vector expression specifically in Schwann cells. Long lasting expression of transgenes in Schwann cells was achieved after intraneural (Sargiannidou et al., 2015a), and intrathecal injection (Kagiava et al., 2016; Schiza et al., 2019) when packaged into a lentiviral vector. Since lentiviral vectors can carry plasmids up to 8 kb, a full-length promoter along with a larger gene can be easily inserted in the vector. However, this is not the case for the AAV vectors which can carry only 4.7 kb (Wu et al., 2010), having a limitation for packaging large transgenes. Thus, for gene therapy applications using AAV vectors the development of shorter cell-specific promoters may be needed in order to treat some of the demyelinating CMT forms. 4. Gene delivery approaches to target Schwann cells and the PNS Delivery of genes to the PNS remains a challenge, since peripheral nerves are not easily accessible to systemically delivered viral vectors owing to the presence of the blood-nerve barrier. In order to develop gene therapy approaches for demyelinating CMT neuropathies, the gene of interest will have to be delivered to Schwann cells since the majority of genes causing demyelinating CMT have cell-autonomous effects (Rossor et al., 2013; Scherer and Wrabetz, 2008; Scherer et al., 2015; Suter and Scherer, 2003). Various gene delivery approaches directly to the PNS have been tried for targeting either Schwann cells or neurons in a variety of preclinical studies. Intravenous, intramuscular, intrathecal, or intraneural injections and even injections into the dorsal root ganglia (DRG) have been used for the delivery of plasmids and different viral vectors. Intraneural injection directly into the sciatic nerve has been used successfully for gene delivery to Schwann cells using AAV and lentiviral vectors (Glatzel et al., 2000; Gonzalez et al., 2014; Homs et al., 2011; Sargiannidou et al., 2015a; Van Hameren et al., 2018). Intraneural gene therapy resulted in stable expression in over 50% of myelinating Schwann cells along the entire length of the sciatic nerve, and improved nerve pathology in a mouse model of X-linked CMT (Sargiannidou et al., 2015a). However, gene expression was only detected in the injected nerves. The utility of this approach for clinical translation is limited by the invasiveness of the technique and the fact that multiple nerves will need to be injected to achieve a functional therapeutic result. Likewise, direct injection of viral vectors into the DRGs resulted in gene expression in up to 30% of DRG neurons (Pleticha et al., 2014). This

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approach may be useful for predominantly sensory neuropathies. However, the potential for clinical translation is limited due to safety concerns in humans. Intramuscular injections have been used to indirectly deliver therapeutic and neuroprotective genes to the peripheral nerves. The rationale for intramuscular delivery is that vectors or the trophic factors they express can be taken up by the supplying peripheral nerve through the neuromuscular junction. This approach has shown some promise after intramuscular delivery of a viral vector expressing neurotrophin-3 (NT-3) in a model of CMT1A (Sahenk et al., 2014; Yalvac et al., 2018). Although this approach is clinically more feasible (Hislop et al., 2014), injections in multiple muscles will still be required to obtain a more widespread therapeutic effect. However, intravenous administration of vectors using muscle-specific promoters may counteract this problem. Gene delivery through a lumbar intrathecal injection appears to be a more clinically translatable approach to target the PNS. The rationale for this route is that the cerebrospinal fluid (CSF) within the intrathecal space is in continuity with the endoneurial fluid reaching the peripheral nerves. The dura matter enclosing the spinal intrathecal space merges with the epineurial connective tissue layers at the central ends of peripheral nerves (Haller and Low, 1971) as they transit to the spinal roots, so that the subarachnoid and endoneurial space are in continuity (Himango and Low, 1971; McCabe and Low, 1969) allowing CSF to enter the endoneurial fluid (Pettersson, 1993). Pressure gradients are likely to promote the diffusion of particles from the subarachnoid space into the peripheral nerves given that the intrathecal CSF pressure at about 10 mm Hg is higher compared to 3-5 mm Hg in DRGs and 1-2 mm Hg in peripheral nerves (Mizisin and Weerasuriya, 2011). Different studies have demonstrated that AAV and LV vectors can be safely delivered by lumbar intrathecal injection and can access both the CNS and PNS resulting in efficient gene expression in DRGs, spinal roots and peripheral nerves (Beutler et al., 2005; Glatzel et al., 2000; Kagiava et al., 2016; Vulchanova et al., 2010) in addition to the CNS (Hordeaux et al., 2015). In particular, intrathecal AAV9 vector injections showed promising results for CNS expression (Bevan et al., 2011; Duque et al., 2015). Furthermore, intrathecal administration offers the possibility to reach the nervous system while evading the anti-AAV-neutralizing antibodies (Gray et al., 2013). Ongoing clinical trials based on intrathecal delivery aim to treat GAN (NCT02362438) and Batten disease (NCT02725580). Furthermore, preclinical studies for SMA have been done with intrathecal injections in rodents and larger animal models (Passini et al., 2014) and intrathecal injections are used for ASO therapy. Recent findings in large animal models of lysosomal storage diseases provide encouraging evidence 12

that intrathecally delivered AAV vectors could provide expression in peripheral nerves (Gurda and Vite, 2019). Even though intravenous delivery would be the most convenient route of administering gene therapy to neuropathy patients, there are many issues associated with it. A very high viral load is required in order to achieve relatively low transduction efficiency. Different AAV serotypes are likely to have variable tissue tropism after intravenous injection (Duque et al., 2009; Schuster et al., 2014b). They also differ in their rate of viral clearance from the blood as well as in their expression kinetics, with AAV7 and 9 having the fastest onset of expression (Zincarelli et al., 2008). In recent years, many AAV serotypes have been developed and tested for gene delivery to different tissues (Challis et al., 2019; Chan et al., 2017; Rahim et al., 2011; Schuster et al., 2014a). Both the AAV9 and AAVrh10 serotypes have been shown to provide a high expression throughout the CNS after intravenous administration, especially in neonatal mice (Foust et al., 2009; Tanguy et al., 2015) and rats (Jackson et al., 2015), indicating efficient transport through the BBB, especially during development. Intravenous delivery of these vectors is currently used in clinical trials for spinal muscular atrophy (SMA) treatment (ClinicalTrials.gov Identifier: NCT02122952). However, it remains to be shown whether efficient transduction of Schwann cells could be achieved by intravenous administration of AAV vectors including in adult animals and patients. 5. Proof of concept studies in demyelinating CMT 5.1. Gene replacement therapy in models of CMT1X Based on the pathogenesis of CMT1X and cellular mechanisms of Cx32 mutations with cellautonomous loss of function, we developed a gene addition approach for Schwann cell targeted delivery of WT Cx32 using a lentiviral vector. Intraneural (Sargiannidou et al., 2015a) or intrathecal (Kagiava et al., 2016; Kagiava et al., 2018) GJB1 gene delivery in Cx32 KO mice resulted in improvement by about 50% in the ratios of abnormally myelinated fibers and reduction of inflammatory cells, both main features of the neuropathy in this model. The use of the myelin protein zero (Mpz) promoter restricted the gene expression in myelinating Schwann cells as indicated by double staining with different cell markers. The intrathecal approach (Kagiava and Kleopa, 2018) allowed more widespread expression in spinal roots and distal nerves, leading also to functional benefit. Motor performance in rotarod testing was improved by 5-fold in the lower speed and 7-fold in the higher speed. Cell targeted gene delivery improved also the electrophysiological properties of Cx32 KO 13

mice as indicated by the quadriceps muscle contractility and in vitro motor nerve conduction velocity studies. Muscle contractility was increased by 13 % while conduction velocities increased by 28 % although both parameters failed to reach the values of wild type mice of the same age, indicating only a partial phenotype rescue (Kagiava et al., 2016). Quantification of the abnormally myelinated fibers and foamy macrophages confirmed the beneficiary effects of the gene addition with improvement in both pathological features in lumbar anterior roots, in femoral motor nerves and in mid-sciatic nerves (Figure 1A-B). Subsequently we tested the gene addition therapy in models expressing representative CMT1X mutants on a Cx32 KO mice background (Sargiannidou et al., 2009) including the ER-retained T55I and the Golgi-retained R75W and N175D mutants (Kagiava et al., 2018). The ER-retained T55I KO mice showed a similar therapeutic response as the Cx32 KO, but in the transgenic lines expressing the Golgi-retained R75W or N175D mutants the therapeutic response was greatly abolished. The presence of these mutants appeared to reduce the trafficking of virally delivered WT Cx32 to paranodal areas resulting in loss of phenotype rescue (Kagiava et al., 2018). Thus, gene addition did not improve functional and morphological properties of the Golgi-retained mutants indicated by the behavioral analysis showing no improvement in the rotarod and foot grip tests while morphological analysis showed either no (R75W) or partial (N175D) improvement of pathological changes. In contrast, the ER-mutant T55I improved in all aspects. Reflecting the results of in vivo testing, in vitro analysis co-expressing various CMT1X mutants with the WT Cx32 confirmed that certain Golgi-retained mutants may interfere with the WT Cx32. In particular, immunostaining showed co-localization of the WT Cx32 with certain mutants in the Golgi and reduced or completely lost formation of GJ plaques on the cell membrane. Moreover, coimmunoprecipitation experiments confirmed that some (but not all) Golgi-retained CMT1X mutants (R75W, M93V, N175D) interact directly with co-expressed WT Cx32, in contrast to ER-retained mutants (Kyriakoudi et al., 2017). Thus, although most CMT1X mutants cause loss of Cx32 function (Sargiannidou et al., 2009), and no WT-mutant Cx32 interaction can take place under normal conditions in this Xlinked disease, it appears that CMT1X mutants that exit the ER and reach the Golgi may potentially interfere with virally delivered WT Cx32 during hexamer formation. This challenge needs to be overcome in future efforts to develop a gene therapy approach for CMT1X, either by increasing the level of virally delivered WT Cx32, as can be achieved by AAV vectors, or by silencing the mutant allele in combination with replacing the WT Cx32. 5.2. Gene replacement therapy for CMT4C 14

Using a similar Schwann cell targeted approach as for CMT1X, we generated a lentiviral vector to deliver the human SH3TC2 cDNA along with myc tag driven by the 1.1Kb Mpz promoter. Since Sh3tc2-/- mice develop a very early phenotype, we optimized the intrathecal vector delivery at 3 weeks of age, to maximize the therapeutic benefit. After lumbar intrathecal injection in Sh3tc2-/- mice, the SH3TC2 protein was expressed and localized correctly in the perinuclear Schwann cell cytoplasm in lumbar roots and sciatic nerves, and co-localized with interacting protein Rab11. A treatment trial in groups of Sh3tc2-/- mice resulted in significant phenotype improvement by 3 months of age in treated compared to mock-treated mice. Morphological analysis of lumbar roots and sciatic nerves in fully treated mice revealed improvement of g-ratios and myelin thickness as well as in the number of demyelinated fibers compared to mock-treated littermates (Schiza et al., 2019) (Figure 1CD). Furthermore, plasma sample analysis confirmed the beneficiary results of gene delivery ameliorating the neurofilament light levels, a marker of axonal damage with clinical relevance (Sandelius et al., 2018). Behavioral and electrophysiological analysis showed that Sh3tc2-/- mice significantly improved in motor performance after treatment. In addition, electrophysiological analysis showed improvement of the motor nerve conduction velocities in the fully treated mice compared to mock, but as with the Cx32 KO studies, without reaching the WT values. Finally, the nodal elongation that has been highlighted as a feature of CMT4C pathology both in biopsied nerves from patient and in the mouse model (Arnaud et al., 2009) was also partly rescued in the treatment group. This finding indicates that SH3TC2 function that plays a role on the development and maintenance of myelinating Schwann cells was restored (Schiza et al., 2019). These results provide a proof of principle that gene replacement can rescue the CMT4C model and thus should be further pursuit to provide a therapeutic approach to treat CMT4C patients. Work that remains to be done towards this direction is to demonstrate also post-onset therapeutic response in older Sh3tc2-/- mice, as well as to increase the rates and levels of SH3TC2 expression for example by testing AAV-mediated delivery, to maximize the therapeutic rescue. 6. Challenges and future perspectives Although these initial studies in CMT1X and CMT4C models of demyelinating neuropathy provide a proof of principle that Schwann cell targeted gene replacement therapy can rescue the respective phenotypes, overall expression rates in Schwann cells following LV vector injection were relatively low and therapeutic effects were partial. Thus, better vector biodistribution and higher gene expression levels are likely to provide a more robust therapeutic response. Alternative vectors such as the AAV could offer significant advantages 15

if their efficacy to target Schwann cells is further demonstrated. Another issue to be clarified will be the extend of viral vector biodistribution to the PNS following intrathecal delivery, not only in rodents, but also in larger animals which are closer to human size, such as nonhuman primates. In conclusion, recent progress in developing gene therapy targeted to myelinating Schwann cells in the PNS has generated optimism for the possibility to replace or silence genes associated with inherited demyelinating neuropathies. Further optimization of viral vectors, delivery methods, biodistribution and promoters is needed so that this effort can lead to clinical translation of treatments for CMT patients.

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Figure legend

Figure 1: Improved pathology of peripheral nerves following gene replacement therapy in models of demyelinating neuropathies. A-B: Images of semithin sections of toluidine-stained femoral motor nerves from 8-month old Gjb1-/-/Cx32 KO mice that were intrathecally injected either with the mock-vector (Mpz-Egfp) in A, or therapeutic vector (Mpz-GJB1) in B. There is improvement in pathology with fewer abnormally myelinated and remyelinated (r) or demyelinated (*) fibers in the treated mouse model of CMT1X. C-D: Semithin sections of toluidine-stained mid-sciatic nerves from 3-month old Sh3tc2-/- mice that were intrathecally injected either with the mock-vector (Mpz-Egfp) in C, or therapeutic vector (Mpz-SH3TC2) in D, show improvement of dys-and demyelinating pathology with fewer demyelinated (*) fibers and thicker myelin sheaths in the treated mouse model of CMT4C.Scale bar in A: 10 μm.

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Table 1: Classification and cellular mechanisms of demyelinating CMT Neuropathies Type

Gene

(OMIM)

(OMIM)

Gene Function

Disease mechanism

Autosomal dominant demyelinating neuropathies (CMT1) HNPP (162500)

PMP22 (601097)

Compact myelin protein, regulation of myelin thickness and maintenance

Gene dosage effects (duplication in CMT1A, deletion in HNPP), impaired regulation of myelin biosynthesis, alteration of mRNA processing, destabilization of myelin, demyelination and/or excessive myelin formation

CMT1B (118200)

MPZ (159440)

Compact myelin protein, myelin compaction

Impaired myelin compaction, retention in endoplasmic reticulum (ER) and unfolded protein response

CMT1C (601098)

LITAF (603795)

Endosomal sorting and cell signaling

Impaired protein degradation in early endosomes

CMT1D (607678)

EGR2 (129010)

Transcription and mRNA processing

Impaired expression of myelin-related genes

CMT1E (118300)

PMP22 (601097)

Compact myelin protein, regulation of myelin thickness and maintenance

Retention and accumulation of mutant PMP22 in ER, unfolded protein response

CMT1F (607734)

NEFL (162280)

Component of axonal cytoskeleton

Loss of myelinated axons

CMT1G (618279)

PMP2 (170715)

Myelin protein serving lipid dynamics, membrane stability

Aberrant transport of fatty acids, impaired organization of compact myelin, short internodes

CMT1A (118220)

X-linked demyelinating / intermediate neuropathy CMT1X (302800)

GJB1 (304040)

Formation of gap junctions through non-compact myelin layers

Disturbed axonal and myelin homeostasis, impaired Schwann cell-axon signaling, axonal degeneration and demyelination

Autosomal recessive demyelinating neuropathies (CMT4) CMT4A (214400)

GDAP1 (606598)

Outer mitochondrial membrane protein, regulates mitochondrial dynamics

Abnormal mitochondrial dynamics

CMT4B1 (601382)

MTMR2 (603557)

Dual specific phosphatase, dephosphorylates PI(3,5)P2 endosomal sorting

Increased PI(3,5)P2, altered ERK1/2 and AKT signaling, abnormal vesicular trafficking, focally folded myelin sheaths

CMT4B2 (604563)

SBF2 / MTMR13 (607697)

Phosphatase that functions as a scaffold for MTM1, interacts with MTMR2

CMT4B3

SBF1

Endosomal sorting and cell

29

Dysregulated membrane homeostasis

(615284)

(603560)

signaling

CMT4C (601596)

SH3TC2 (608206)

Endosomal sorting and cell signaling, interacts with Rab11

Impaired perinuclear endocytic recycling compartment, dysregulation of myelination, nodal widening

CMT4D (601455)

NDRG1 (605262)

Phosphoprotein that is downstream of integrin signaling

Impaired Schwann cell signaling and demyelination

CMT4E (605253)

EGR2 (129010)

Transcription factor

Impaired expression of myelin-related genes

CMT4F (614895)

PRX (605725)

PDZ domain protein that links dystroglycan to the actin cytoskeleton

Disrupted Schwann cell cytoskeleton and basal membrane adhesion

CMT4G (605285)

HK1 (142600)

Glucose metabolism at the outer mitochondrial membrane

Disturbed mitochondrial function

CMT4H (609311)

FGD4 (11104)

Schwann cell cytoskeleton and basal lamina adhesion

Disrupted actin binding, disturbed cytoskeleton

CMT4J (611228)

FIG4 (609390)

Phosphatase, complexed with Vac14 and Fab1 kinase, activates PI(3,5)P2

Disrupted PI(3,5)P2 signaling, impaired autophagy

CMT4K (616684)

SURF1 (185620)

Assembly factor of mitochondrial complex IV (COX)

COX deficiency, abnormal mitochondrial respiratory chain function

30

Highlights: 

Demyelinating CMT results from mutations in Schwann cell genes



Intrathecal vector delivery could provide a translatable access to PNS



Schwann cell-targeted gene expression using myelin-specific promoters



Encouraging gene replacement studies in CMT1X and CMT4C models

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