Nerve cells transmit messages along their length in the form of electrical signals. Much like an electrical wire, the nerve fiber or axon is coated by a multiple-layered insulation, called the myelin sheath. However, unlike electrical insulation, the myelin sheath is regularly interrupted to expose short regions of the underlying nerve. These exposed regions and the adjacent regions underneath the myelin contain ion channels that help to propagate electrical signals along the axon.

Peroxisomes are compartments in animal cells that process fats. Genetic mutations that prevent peroxisomes from working properly can lead to diseases where the nerves cannot transmit signals correctly. This is thought to be because the nerves lose their myelin sheath, which largely consists of fatty molecules.

The nerves outside of the brain and spinal cord are known as peripheral nerves. Kleinecke et al. have now analyzed peripheral nerves from mice that had one of three different genetic mutations, preventing their peroxisomes from working correctly. Even in cases where the mutation severely impaired nerve signaling, the peripheral nerves retained their myelin sheath.

The peroxisome mutations did affect a particular type of potassium ion channel and the anchor proteins that hold these channels in place. The role of these potassium ion channels is not fully known, but normally they are only found close to regions of the axon that are not coated by myelin. However, the peroxisome mutations meant that the channels and their protein anchors were now also located along the myelinated segments of the nerve’s axons. This redistribution of the potassium ion channels likely contributes to the peripheral nerves being unable to signal properly.

In addition, Kleinecke et al. found that disrupting the peroxisomes also affected another cell compartment, called the lysosome, in the nerve cells that insulate axons with myelin sheaths. Lysosomes help to break down unwanted fat molecules. Mutant mice had more lysosomes than normal, but these lysosomes did not work efficiently. This caused the nerve cells to store more of certain types of molecules, including molecules called glycolipids that stabilize protein anchors, which hold the potassium channels in place. A likely result is that protein anchors that would normally be degraded are not, leading to the potassium channels appearing inappropriately throughout the nerve.

Future work is now needed to investigate whether peroxisomal diseases cause similar changes in the brain. The results presented by Kleinecke et al. also suggest that targeting the lysosomes or the potassium channels could present new ways to treat disorders of the peroxisomes.

Schwann cells, the myelinating cells of the peripheral nervous system (PNS), contain peroxisomes in distant cytoplasmic compartments, including myelin channels and cytosolic loop regions close to nodes of Ranvier (Kassmann et al., 2011). Peroxisomes degrade fatty acids derived from myelin lipids and generate precursors of myelin plasmalogens (Waterham et al., 2016). Lysosomal compartments are also present in these nodal regions (Gatzinsky et al., 1997), and growing evidence from in vitro studies suggests interactions between both types of organelles (Baarine et al., 2012; Chu et al., 2015; Thai et al., 2001).

Peroxisomal dysfunction is caused by mutations of genes encoding peroxisomal proteins or peroxisomal biogenesis factors (Waterham et al., 2016). In humans, loss-of-function mutations of ABCD1 are responsible for the disease X-linked Adrenoleukodystrophy (X-ALD). This peroxisomal ATP-binding cassette (ABC-) transporter mediates the import of very long-chain fatty acids (VLCFA) into the organelle. In consequence, ABCD1-deficient peroxisomes are not capable of importing and degrading VLCFA that are specific substrates of peroxisomal β-oxidation (Kemp et al., 2012). A more severe impairment of peroxisomes is caused by lack of the Hsd17b4 gene (also called multifunctional protein 2; Mfp2 gene) that encodes a central enzyme of peroxisomal β-oxidation. In MFP2-deficient cells, the β-oxidation of virtually all peroxisome-specific substrates, including VLCFA, is inhibited (Verheijden et al., 2013). A complete disruption of the organelle is observed in the absence of peroxisome biogenesis factor peroxin 5 (PEX5). This cycling receptor recognizes proteins with a peroxisomal targeting sequence type 1 (PTS1) and is involved in their transfer into peroxisomes. PEX5-dependent protein import applies to the majority of peroxisomal enzymes. Thus, PEX5-deletion disrupts peroxisomal function substantially (Waterham et al., 2016).

Schwann cell lipid metabolism is rate-limiting for myelination and is important for maintenance of axonal integrity (Saher et al., 2011; Viader et al., 2013), which requires in addition to membrane wrapping the assembly of nodal, paranodal, and juxtaparanodal membrane proteins (Rasband and Peles, 2015). The juxtaparanodal domain of myelinated axons harbors voltage-gated shaker-type potassium channels, K_v1.1 (KCNA1) and K_v1.2 (KCNA2; Chiu and Ritchie, 1980; Robbins and Tempel, 2012), which also align the inner mesaxon as a thin band (Arroyo et al., 1999). Associated with connexin-29 hemichannels (Rash et al., 2016), their clustering and anchoring at juxtaparanodes requires the neuronal membrane proteins CASPR2 and TAG-1, the latter expressed by glia and neurons (Poliak et al., 1999b; Traka et al., 2003). K_v1 channels have been proposed to play a role in regulating fiber excitability (Baker et al., 2011; Glasscock et al., 2012), but the exact in vivo function of these fast-opening/slowly inactivating channels remains unknown (Arancibia-Carcamo and Attwell, 2014).

Cnp-Cre::Pex5^flox/flox mice, termed cKO or 'mutants' in the following, lack peroxisomal protein import in Schwann cells (Figure 1a; Figure 1—figure supplement 1a). The PNS of these mice is well myelinated and unlike the CNS (Kassmann et al., 2007) without immune-mediated injury, in agreement with pilot observations (Kassmann et al., 2011). Upon closer inspection, we determined about 50% genomic recombination, corresponding to the fraction of Schwann cell (SC) nuclei in sciatic nerves (Figure 1—figure supplement 1b). Teased fiber preparations, stained for PMP70, revealed peroxisomes as puncta. In mutant nerves, these were import-deficient 'ghosts', as evidenced by cytoplasmic catalase, normally a luminal peroxisomal marker (Figure 1b).

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Peroxisomal dysfunction in myelinating SC was confirmed by lipid mass spectrometry (Figure 1c, Figure 1—figure supplement 1c), showing reduced plasmalogens (PE_P-) and its precursor alkylated phosphatidyl-ethanolamines (PE_O-; Wanders, 2014). Also VLCFA were increased, indicating the accumulation of peroxisomal β-oxidation substrates (Figure 1d; Figure 1—figure supplement 1d).

We determined nerve conduction velocity (NCV) by electrophysiology of isolated sciatic nerves (to avoid possible contributions of altered muscle endplates) at the age of 2 months (Figure 1e–g; Figure 1—figure supplement 2a). For all stimulus intensities tested, responses of mutant nerves were different from controls (Figure 1—figure supplement 2b). Compound action potentials (CAPs) and NCV were diminished in mutants (mean: 28 ± 4.7 m/s) compared to controls (41.5 ± 3.6 m/s; Figure 1e). Thresholds to evoke a signal were only slightly elevated (155µA versus 135µA), but the maximal response was 50% of control (Figure 1f,g). Also, in vivo recordings revealed significantly reduced compound muscle action potentials (CMAPs) in mutant mice (Kassmann et al., 2011). This was more enhanced when stimulating proximally than distally, which clinically defines conduction blocks (Figure 1—figure supplement 2c).

We suspected that reduced nerve conduction would be explained by demyelination. Surprisingly, by immunohistochemistry and Western blot analysis structural myelin proteins, including PMP22, MPZ/P0, and P2, were not significantly altered (Figure 1—figure supplement 3a,b). Only PLP, a minor PNS myelin protein, showed significant reduction. Also by electron microscopy (EM), myelin thickness, periodicity, and compaction were indistinguishable (Figure 1h, Figure 1—figure supplement 3c). Next, we determined internodal length in teased fiber preparations, which was shorter in mutant (623 nm) than in control fibers (691 nm; Figure 1—figure supplement 3d), but unlikely sufficiently reduced to cause a slower conduction by itself (Wu et al., 2012). Importantly, while the reduced CAP suggested significant axon loss at 2 months, the morphometric analysis of entire nerve cross-sections revealed normal axonal numbers and calibers (Figure 1i). Thus, reduced CAPs were most likely caused by functional conduction blocks rather than physical axon loss. Signs of axonal transport defects (APP accumulations) were not observed (Figure 1—figure supplement 3e), and also, indirect signs of neurodegeneration (neuroinflammation) were not significantly enhanced in nerves at 2 months (Figure 1—figure supplement 4a–d).

Disruptions of axo-glial junctions might cause leak currents and thus conduction failure (Arancibia-Carcamo and Attwell, 2014). Yet, both neurofascin-155 (NF155) and axonal CASPR, that is adhesion proteins that mediate axo-glial contacts (Desmazieres et al., 2014; Normand and Rasband, 2015), were normally localized (Figure 1j). By EM, we never found the detachment of paranodal loops (Figure 1k,l), even where abnormally enlarged by accumulated vesicles (Kassmann et al., 2011). Next, we examined the distribution of ion channels (Devaux et al., 2004; Rasband and Peles, 2015) in sciatic nerve teased fibers from 2- and 9-month-old mutants and controls. Nodal Na_v1.6 and K_v7.2, as well as their anchoring proteins, were normally flanked by CASPR-positive paranodes (Figure 2—figure supplement 1a–c). Surprisingly, the juxtaparanodal potassium channel protein K_v1.1 displayed an abnormal localization, frequently displaced into the adjacent internodal region, and to a minor extent into paranodes. K_v1.1 clusters were in the majority of cases still maintained at juxtaparanodes, suggesting that internodal mislocalization is secondary (Figure 2—figure supplement 1d,e; Figure 2a). Indeed, the pathology of internodal K_v1 channels progressively increased with age frequently presenting more than one extra cluster along one internode in aged mutants, while shifts into the nodal direction were not progressive (Figure 2a,b; Figure 2—figure supplement 1e). We confirmed the observation of displaced juxtaparanodal K_v1 channels using K_v1.2 antibody (data not shown). At 9 months, we determined a twofold overall increase of K_v1.1 in nerve lysates (Figure 2c,d). However, as most K_v1.1 is expressed by unmyelinated fibers, the increase of internodal K_v1.1 in myelinated axons may be significantly higher. Neuronal CASPR2, the cis-anchor for K_v1 channels in the axonal membrane, and TAG-1 on SC are essential for channel clustering at juxtaparanodes (Hivert et al., 2016; Poliak et al., 1999a; Savvaki et al., 2010). Both proteins colocalized with K_v1.1 at regular and ectopic internodal positions (Figure 2e–g). TAG-1 expression was slightly but not significantly elevated (Figure 2h). In contrast, staining for myelin-associated glycoprotein (MAG) did not reveal a connection between Schmidt-Lanterman incisures and the ectopic K_v1.1/TAG-1/CASPR2-positive clusters (data not shown). Although, we cannot exclude that enhanced expression of TAG-1 contributes to the formation of ectopic protein clusters, the observations suggest that in mutant nerves entire membrane patches of the juxtaparanodal compartment accumulate, break-off, and drift into the internodal region.

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Gangliosides are glycosphingolipids important for stabilizing membrane-spanning proteins and axon-glia interactions (Susuki et al., 2007). We analyzed sciatic nerve lysates from 9-month-old mutants by lipid mass spectrometry and noted that many more gangliosides contained VLCFA (a 14-fold enrichment; Figure 2h). To determine their subcellular distribution, we stained teased fibers with a fluorescently tagged cholera-toxin subunit B (CTB), which binds GM1 gangliosides. Also by immunostaining, GM1 was restricted to paranodes in wild-type nerves but widely dispersed into the internodal myelin of mutant nerves (Figure 2—figure supplement 2; Figure 2i). Likewise, ganglioside GD1a could be immunostained as enlarged puncta (>5 µm) in internodal myelin of mutant nerves (Figure 3a). These GD1a-containing vesicles that colocalized with LAMP1 were a frequent finding in mutant internodes, but rarely observed in controls (Figure 3a, left).

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Lysosomal accumulation within paranodal loops.

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Paranodal loops in conditional mutants showed vesicular inclusions of variable size and electron-density (Figure 3b). By immune-labeling of ultra-thin cryosections and teased fibers, the majority of these were LAMP1-positive (Figure 3c,d; Figure 3—source data 1). Even more striking, lysosomal LIMP-2 marked small puncta in controls (<500 nm), but giant compartments within mutant paranodes (Figure 3e, right). Strongly increased abundance of LAMP1 and LIMP-2 was confirmed by Western blotting (Figure 3f,g). Decreased mRNA levels (Figure 3h) suggest that the enrichment of LAMP1 and LIMP-2 marks organelle accumulation rather than enhanced lysosomal biogenesis.

Lysosomal markers are found in autophagosomes and early/late endosomes (Saftig and Klumperman, 2009). We thus phenotyped accumulated vesicles with antibodies specific for EEA1, RAB7, and ATG5. Neither marker was increased at mutant paranodes (Figure 3—figure supplement 1a–c). Therefore, the majority of accumulating vesicles within mutant paranodes are likely bona fide lysosomes, serving the degradation of myelin-associated proteins/lipids, including gangliosides (Luzio et al., 2007; Saftig and Klumperman, 2009). To determine the relationship between paranodal lysosomes (Gatzinsky et al., 1997) and peroxisomes, we immunostained for PMP70 and LAMP1, revealing a close spatial association (Figure 3i) as reported for cultured cells (Chu et al., 2015). We determined higher activities of lysosomal α-mannosidase and β-hexosaminidase in mutant nerve lysates (Figure 3j), and note that increased compartment size and enzymatic activities are known phenomena of lysosomal disorders (Sardiello and Ballabio, 2009), yet associated with organelle dysfunction.

Theoretically, complete peroxisomal failure in Cnp-Cre::Pex5^flox/flox mice could perturb many aspects of SC metabolism in the observed neuropathy. However, we found that Cnp-Cre::Hsd17b4^flox/flox mice (referred to as MFP2 conditional knockout mice in the following), which specifically lack peroxisomal β-oxidation (Verheijden et al., 2013), share the key features of this novel phenotype, including ectopic internodal K_v1.1 clusters (Figure 4—figure supplement 1a–e), confirming the important role for glial lipid metabolism in maintaining axonal membrane composition, and function.

To investigate whether this new pathomechanism, identified in two mouse models with a glia-specific mutation, is also relevant for a human genetic disease, we analyzed ABCD1-deficient mice, a model for X-ALD/AMN (Forss-Petter et al., 1997). Sciatic nerve axons and myelin were morphologically intact, even at 22 months of age (Figure 4a–c). However, ABCD1-mutants exhibited a similar mislocalization of K_v1.1 positive channels as observed in both conditional (Pex5 and Mfp2) mutants. The quantification showed that mislocalization of internodal K_v1.1 was temporally progressive. In accordance with our model of membrane patches drifting into internodes, significant alterations were observed only at 22 months of age, but not yet at 2 months (Figure 4d–e). This pathology was accompanied by lysosomal accumulates in myelinated fibers (Figure 4f–h).

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Our data suggest an important pathomechanism of peroxisomal dysfunction in peripheral nerves is the secondary impairment of lysosomes. For example, gangliosides, which are frequently esterified with VLCFA and 2-hydroxy fatty acids (Chrast et al., 2011; Sandhir et al., 2000), must be degraded in lysosomes. However, subsequent β-oxidation of VLCFA and 2-hydroxy fatty acids requires peroxisomes (Foulon et al., 2005; Wanders, 2014). Thus, our mouse model provides first in vivo evidence for functional interactions between lysosomes and peroxisomes, so far only discussed for cultured cells (Chu et al., 2015).

One consequence of lysosomal impairment is the perturbation of normal ganglioside turnover. Gangliosides are in close association with K_v1-anchoring proteins (Loberto et al., 2003) and provide stability to the juxtaparanodal clusters (Susuki et al., 2007), which resemble giant membrane 'lipid rafts'. As mechanism for ectopic localization of axonal K_v1 channels, we suggest the following working model (Figure 4—figure supplement 2): Lack of ganglioside breakdown causes their accumulation in the glial juxtaparanodal membrane. Thereby, clusters of K_v1 and associated anchoring proteins (TAG-1 and CASPR2) on the axonal side gain abnormal stability, break-off, diffuse into the internodal region, and escape from regular turnover. It is likely that also glial TAG-1 remains anchored to the axonal K_v1-/CASPR2-/TAG-1 clusters, but this cannot be distinguished by TAG-1 immunofluorescence.

K_v1-channels are fast-opening, voltage-gated channels that are activated by mild voltage-shifts and thought to regulate fiber excitability (Glasscock et al., 2012). The integrity of K_v1 channels is also impaired in CASPR2-/- mice, but perturbation of nerve function is not a feature (Poliak et al., 2003). Unlike PEX5 mutant nerves, CASPR2-/- nerves do not exhibit elevated levels of K_v1.1 channel protein. Also, the ectopic K_v1.1+ clusters are situated directly adjacent to juxtaparanodes in CASPR2-deficient fibers, not along the internodes as observed in PEX5 mutants. Considering the severe progressive decline of nerve function in PEX5 mutant mice, which becomes clearly more affected with age (data not shown), it is unlikely that a rather non-progressive pathology, that is the shift of K_v1 into paranodes, is the underlying cause of conduction slowing. In contrast, the progressive shift of K_v1 toward internodes correlates well with disease progression. Theoretically, K_v1-mediated currents in the internode could hyperpolarize the axonal membrane and dampen the conductivity of spiking axons. Such ‘feedback regulation’ might prevent hyperexcitability, which is a feature of mice lacking K_v1.1 function (Zhou et al., 1998). On the other hand, a ‘gain-of-function’ effect by opening additional ectopic internodal K_v1 channels, may contribute to a disturbed equilibrium of axonal ions causing slowed NCV and conduction blocks. Direct proof of this model would require patch clamping single axons underneath myelin, which is impossible at present.

Peripheral neuropathy has been associated with numerous metabolic diseases, including AMN. In this clinically milder variant of ABCD1 loss-of-function mutations, the reduced NCV was assumed to reflect demyelination, which is a hallmark in the brain of severely affected patients. Similarly, conditional PEX5 knockout mice display a cerebral inflammatory demyelination, while such pathology is absent in sciatic nerves. This remarkable discrepancy between the peripheral and central nervous system pathology is currently unexplained and presumably related to differently responding cell populations (Kassmann, 2014). Also in ABCD1-deficient nerves neither demyelination nor axon degeneration, but ectopic axonal K_v1 channels was a progressive pathological feature as early as at 2 months of age, preceding the disease onset by more than 1 year (Pujol et al., 2002). Our present findings therefore expand the emerging role of myelinating glial cells in maintaining axonal membrane composition and thereby influencing axonal function, independent of myelin itself.

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Author details

1. Sandra Kleinecke

   Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

   Contribution

   SK, Mouse breeding, histology, light microscopy, data analysis, incl. statistics, and interpretation of data

   Competing interests

   No competing interests declared.

2. Sarah Richert

   Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

   Contribution

   SR, Conceptualization, Investigation, Mouse breeding, histology, light microscopy, data analysis, incl. statistics, and interpretation of data

   Competing interests

   No competing interests declared.

3. Livia de Hoz

   Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

   Contribution

   LdH, Formal analysis, Methodology, Data analysis, incl. statistics, and interpretation of electrophysiology data

   Competing interests

   No competing interests declared.

4. Britta Brügger

   University of Heidelberg, Biochemistry Center (BZH), Heidelberg, Germany

   Contribution

   BB, Software, Formal analysis, Funding acquisition, Investigation, Methodology, Lipid mass spectrometry: data analysis, incl. statistics, and interpretationof data

   Competing interests

   No competing interests declared.

5. Theresa Kungl

   Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

   Contribution

   TK, Conceptualization, Investigation, Histology, light microscopy, data analysis, incl. statistics, and interpretation of data

   Competing interests

   No competing interests declared.

6. Ebrahim Asadollahi

   Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

   Contribution

   EA, Software, Investigation, Methodology

   Competing interests

   No competing interests declared.

7. Susanne Quintes

   Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

   Contribution

   SQ, Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared.

8. Judith Blanz

   Unit of Molecular Cell Biology and Transgenic, Institute of Biochemistry, University of Kiel, Kiel, Germany

   Contribution

   JB, Formal analysis, Methodology

   Competing interests

   No competing interests declared.

9. Rhona McGonigal

   Institute of Infection, Immunity, and Inflammation, University of Glasgow, Glasgow, United Kingdom

   Contribution

   RM, Formal analysis, Validation, Investigation, Methodology

   Competing interests

   No competing interests declared.

10. Kobra Naseri

    Birjand University of Medical Sciences, Birjand, Iran

    Contribution

    KN, Investigation, Visualization, Methodology

    Competing interests

    No competing interests declared.

11. Michael W Sereda

    Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

    Contribution

    MWS, Formal analysis, Methodology, Data analysis, incl. statistics, and interpretation ofelectrophysiology data

    Competing interests

    No competing interests declared.

12. Timo Sachsenheimer

    University of Heidelberg, Biochemistry Center (BZH), Heidelberg, Germany

    Contribution

    TS, Investigation, Methodology

    Competing interests

    No competing interests declared.

13. Christian Lüchtenborg

    University of Heidelberg, Biochemistry Center (BZH), Heidelberg, Germany

    Contribution

    CL, Investigation, Methodology

    Competing interests

    No competing interests declared.

14. Wiebke Möbius

    Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

    Contribution

    WM, Investigation, Methodology

    Competing interests

    No competing interests declared.

    "This ORCID iD identifies the author of this article:" 0000-0002-2902-7165

15. Hugh Willison

    Institute of Infection, Immunity, and Inflammation, University of Glasgow, Glasgow, United Kingdom

    Contribution

    HW, Formal analysis, Methodology

    Competing interests

    No competing interests declared.

16. Myriam Baes

    Department of Pharmaceutical and Pharmacological Sciences, Cell Metabolism, KU Leuven- University of Leuven, Leuven, Belgium

    Contribution

    MB, Resources, Provided floxed mice andrevised the article critically for important intellectual content

    Competing interests

    No competing interests declared.

17. Klaus-Armin Nave

    Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

    Contribution

    K-AN, Funding acquisition, Writing—review and editing, Revising thearticle critically for important intellectual content, Final approval of the version to be published

    Competing interests

    K-AN: Reviewing editor, eLife

    "This ORCID iD identifies the author of this article:" 0000-0001-8724-9666

18. Celia Michèle Kassmann

    Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Göttingen, Germany

    Contribution

    CMK, C.M.K. supervised theproject, designed experiments, analyzed data, and wrote the manuscript

    For correspondence

    kassmann@em.mpg.de

    Competing interests

    No competing interests declared.

    "This ORCID iD identifies the author of this article:" 0000-0003-0993-9455

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