Peroxisomal dysfunctions cause lysosomal storage and axonal Kv1 channel redistribution in peripheral neuropathy.

Impairment of peripheral nerve function is frequent in neurometabolic diseases, but mechanistically not well understood. Here, we report a novel disease mechanism and the finding that glial lipid metabolism is critical for axon function, independent of myelin itself. Surprisingly, nerves of Schwann cell-specific Pex5 mutant mice were unaltered regarding axon numbers, axonal calibers, and myelin sheath thickness by electron microscopy. In search for a molecular mechanism, we revealed enhanced abundance and internodal expression of axonal membrane proteins normally restricted to juxtaparanodal lipid-rafts. Gangliosides were altered and enriched within an expanded lysosomal compartment of paranodal loops. We revealed the same pathological features in a mouse model of human Adrenomyeloneuropathy, preceding disease-onset by one year. Thus, peroxisomal dysfunction causes secondary failure of local lysosomes, thereby impairing the turnover of gangliosides in myelin. This reveals a new aspect of axon-glia interactions, with Schwann cell lipid metabolism regulating the anchorage of juxtaparanodal Kv1-channels.

Introduction

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, Kv1.1 (KCNA1) and Kv1.2 (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). Kv1 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).

Results

Cnp-Cre::Pex5 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).

Figure 1.

Schwann cell-specific PEX5-deficiency causes peroxisome dysfunction and peripheral neuropathy in the absence of axonal loss or dysmyelination.

(A) Scheme of normal (left) and impaired (right) PEX5-dependent peroxisomal protein import. PTS1, peroxisomal targeting signal type 1; PEX5, peroxisomal biogenesis factor peroxin 5. (B) Catalase (red) is present in peroxisomes (PMP70; green) of controls, but is localized in the cytoplasm of mutant fibers. PMP70, peroxisomal membrane protein 70; DAPI-stained SC nuclei are depicted in blue; Scale bar, 10 µm. (C, D) Lipid mass spectrometry of nerve lysates from controls and mutants aged 9 months indicates peroxisomal dysfunction. Peroxisomal products (PEO-) and its corresponding plasmalogens (PEP-) are reduced. Specific substrates of peroxisomal β-oxidation, VLCFA, are accumulated in mutant nerves. Statistics: means ± s.d.; n = 6; Student’s T-test; ***p<0.001. (E–G) Functional impairment of mutant compared to control sciatic nerves is assessed by electrophysiology at the age of 2 months (n = 4). To evoke significant responses of all measured nerves, larger stimulus intensity is required for mutant (0.155mA) as compared to control (0.135mA) nerves. Peak amplitudes plotted against increasing stimulus intensity indicates earlier response saturation of mutant nerves. CAP, compound action potentials; au, arbitrary units; NCV, nerve conduction velocities (each curve representing single-nerve mean responses across intensities). (H) g-ratio analysis by electron microscopy of sciatic nerves as measure of myelin thickness (n = 3; ≥100 randomly chosen axons per nerve at age 2 months). (I) Axonal analysis by methylene blue-stained semithin cross-sections of sciatic nerves of control and mutant mice at the age of 2 months (n = 3). Scale bar, 10 µm. (J) Immunostained teased sciatic nerve fibers of mutants and controls at 2 and 9 months show normal paranodal localization of CASPR (red) and anchoring protein neurofascin (NF155; green). (K, L) Intact transverse bands (arrows) at paranodal loops (PNLs) between axon (Ax) and myelin (M) by electron microscopy in a 9-month mutant nerve, even when PNLs harbor enlarged vesicles (top inset, arrowheads). Mutant nerves display normal adherens junctions (AJ) between loops (bottom inset, asterisk). Scale bars, 500 nm; scale bars in insets, 250 nm.

DOI: http://dx.doi.org/10.7554/eLife.23332.003

(A) PCR on genomic tail DNA using primers specifically generating a 300 bp amplicon after Cre-mediated excision of Pex5-floxed exons 11–14. (B) qRT-PCR reveals a 50% (corresponding to the fraction of SC nuclei) reduction of Pex5 mRNA level in mutant nerves (n = 6). (C) Quantitative mass spectrometry of sciatic nerve lysates. Plasmalogens were abundant in a control nerve (top, green arrows at peaks). The same plasmalogens were hardly detectable in a mutant nerve lysate (bottom, red arrows at peaks). (D) Analysis of VLCFA by gas chromatography–mass spectrometry in control and mutant mice. Statistics: means ± s.e.m.; n = 6; *p<0.05; **p<0.01; ***p<0.001, Student’s T-test (B, D).

DOI: http://dx.doi.org/10.7554/eLife.23332.004

(A) Photographs of electrophysiology setup with a pair of suction electrodes. (B) Pex5 mutant nerves display smaller and delayed responses when elicited at stimulus intensities of 0.17mA (left) or 0.22mA (right). (C) In vivo electrophysiology of controls and PEX5 cKO shows a greater decline of the compound muscle action potential (CMAP) amplitudes measured at more proximal versus distal stimulation indicating conduction blocks. The ratios of the amplitudes elicited by proximal vs. distal stimulation are depicted. Statistics: n = 5, means ± s.d. **p<0.01, Student’s T-test.

DOI: http://dx.doi.org/10.7554/eLife.23332.005

(A) Chromogenic immune-staining for myelin protein zero (MPZ/P0) on paraffin-embedded sciatic nerve sections at 2 months. Scale bar, 50 µm. (B) Abundance of major PNS myelin proteins P0 and PMP22, as well as fatty-acid-binding protein P2, and DM20 were not significantly altered by quantification of Western blot analysis of sciatic nerve lysates obtained at the age of 2 months (n = 4; each normalized to control levels; 1.0). Only the covalently lipid-binding protein PLP showed significant reduction in mutant as compared to control nerves. Alpha-Tubulin served as loading control. Statistics: mean ± s.e.m. *p<0.05; **p<0.01; ***p<0.001; n.s., not significant, Student’s T-test. (C) Electron micrographs of sciatic nerve cross-section showing myelin sheaths of animals aged 2 months. Scale bars, 10 nm. (D) Internodal lengths measured using teased fibers of 2-month-old PEX5 mutant and control animals. Statistics: n = 3, mean ± s.d. *p<0.05, Student’s T-test. (E) Chromogenic immune-staining for APP on longitudinal paraffin-embedded sciatic nerve sections. Scale bar, 50 µm.

DOI: http://dx.doi.org/10.7554/eLife.23332.006

(A, D) Chromogenic immunostaining for MAC-3 (A) and CD3 (D) on longitudinal paraffin-embedded sciatic nerve sections of mice aged 2 and 9 months. (B, C) Mast cells (arrow) visualized and quantified using entire semithin (500 nm) methylene-blue-stained cross-sections of mouse sciatic nerves at 2 months of age (n = 4). Statistics: means ± s.e.m. *p<0.05; **p<0.01; ***p<0.001; Student’s T-test.

DOI: http://dx.doi.org/10.7554/eLife.23332.007

Figure 1—figure supplement 1.

Successful PEX5 ablation causes peroxisomal dysfunction.

(A) PCR on genomic tail DNA using primers specifically generating a 300 bp amplicon after Cre-mediated excision of Pex5-floxed exons 11–14. (B) qRT-PCR reveals a 50% (corresponding to the fraction of SC nuclei) reduction of Pex5 mRNA level in mutant nerves (n = 6). (C) Quantitative mass spectrometry of sciatic nerve lysates. Plasmalogens were abundant in a control nerve (top, green arrows at peaks). The same plasmalogens were hardly detectable in a mutant nerve lysate (bottom, red arrows at peaks). (D) Analysis of VLCFA by gas chromatography–mass spectrometry in control and mutant mice. Statistics: means ± s.e.m.; n = 6; *p<0.05; **p<0.01; ***p<0.001, Student’s T-test (B, D).

DOI: http://dx.doi.org/10.7554/eLife.23332.004

Peroxisomal dysfunction in myelinating SC was confirmed by lipid mass spectrometry (Figure 1c, Figure 1—figure supplement 1c), showing reduced plasmalogens (PEP-) and its precursor alkylated phosphatidyl-ethanolamines (PEO-; 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).

Figure 1—figure supplement 2.

Electrophysiology of mouse sciatic nerves.

(A) Photographs of electrophysiology setup with a pair of suction electrodes. (B) Pex5 mutant nerves display smaller and delayed responses when elicited at stimulus intensities of 0.17mA (left) or 0.22mA (right). (C) In vivo electrophysiology of controls and PEX5 cKO shows a greater decline of the compound muscle action potential (CMAP) amplitudes measured at more proximal versus distal stimulation indicating conduction blocks. The ratios of the amplitudes elicited by proximal vs. distal stimulation are depicted. Statistics: n = 5, means ± s.d. **p<0.01, Student’s T-test.

DOI: http://dx.doi.org/10.7554/eLife.23332.005

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).

Figure 1—figure supplement 3.

Myelin analysis of mouse sciatic nerves.

(A) Chromogenic immune-staining for myelin protein zero (MPZ/P0) on paraffin-embedded sciatic nerve sections at 2 months. Scale bar, 50 µm. (B) Abundance of major PNS myelin proteins P0 and PMP22, as well as fatty-acid-binding protein P2, and DM20 were not significantly altered by quantification of Western blot analysis of sciatic nerve lysates obtained at the age of 2 months (n = 4; each normalized to control levels; 1.0). Only the covalently lipid-binding protein PLP showed significant reduction in mutant as compared to control nerves. Alpha-Tubulin served as loading control. Statistics: mean ± s.e.m. *p<0.05; **p<0.01; ***p<0.001; n.s., not significant, Student’s T-test. (C) Electron micrographs of sciatic nerve cross-section showing myelin sheaths of animals aged 2 months. Scale bars, 10 nm. (D) Internodal lengths measured using teased fibers of 2-month-old PEX5 mutant and control animals. Statistics: n = 3, mean ± s.d. *p<0.05, Student’s T-test. (E) Chromogenic immune-staining for APP on longitudinal paraffin-embedded sciatic nerve sections. Scale bar, 50 µm.

DOI: http://dx.doi.org/10.7554/eLife.23332.006

Figure 1—figure supplement 4.

Young Pex5 mutant nerves lack signs of inflammation.

(A, D) Chromogenic immunostaining for MAC-3 (A) and CD3 (D) on longitudinal paraffin-embedded sciatic nerve sections of mice aged 2 and 9 months. (B, C) Mast cells (arrow) visualized and quantified using entire semithin (500 nm) methylene-blue-stained cross-sections of mouse sciatic nerves at 2 months of age (n = 4). Statistics: means ± s.e.m. *p<0.05; **p<0.01; ***p<0.001; Student’s T-test.

DOI: http://dx.doi.org/10.7554/eLife.23332.007

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 Nav1.6 and Kv7.2, 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 Kv1.1 displayed an abnormal localization, frequently displaced into the adjacent internodal region, and to a minor extent into paranodes. Kv1.1 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 Kv1 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 Kv1 channels using Kv1.2 antibody (data not shown). At 9 months, we determined a twofold overall increase of Kv1.1 in nerve lysates (Figure 2c,d). However, as most Kv1.1 is expressed by unmyelinated fibers, the increase of internodal Kv1.1 in myelinated axons may be significantly higher. Neuronal CASPR2, the cis-anchor for Kv1 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 Kv1.1 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 Kv1.1/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.

Figure 2—figure supplement 1.

Normal distribution of nodal proteins and associated anchors in Pex5 mutant nerves.

Immune-stained sciatic teased fibers of 2- and 9-month-old mice. (A) Nodal sodium (Nav1.6; left) and potassium (Kv7.2; right) channels are normally localized and flanked by paranodal CASPR (red, scale bars, 5 µm). (B, C) Ion channel anchoring proteins βIV-spectrin (green) (B) and ankyrinG (green) (C) are normally localized to nodes of Ranvier and flanked by paranodal CASPR (red). Scale bars, 5 µm. (D) Staining for 4.1G (green), which unlike 4.1B is abundantly present at paranodes, shows normal localization (arrowheads). Potassium channel Kv1.1 clusters (red) are located at juxtaparanodes but in mutants additionally localize along internodes. (E) Triple-staining of Nav1.6 (blue), paranodal CASPR (red), and juxtaparanodal potassium channels Kv1.1 (green) shows preservation of Kv1.1 at juxtaparanodes and at mesaxonal lines (arrows) of mutant fibers. In addition, ectopic internodal Kv1.1 patches are depicted. Scale bar, 10 µm.

DOI: http://dx.doi.org/10.7554/eLife.23332.009

Figure 2.

Abnormal distribution of axonal ion channels Kv1.1, its anchoring proteins, and associated lipids in conditional Pex5 mutant nerves.

(A, B) Immune-stained sciatic teased fibers of 2- and 9-month-old mice reveal normally localized nodal sodium channels (Nav1.6; blue) flanked by paranodal CASPR (red; scale bar, 10 µm). Only mutant nerves show extra Kv1.1+ patches within internodes. This is progressive with age shown by the corresponding quantification at different ages, starting at P19, revealing a shift of Kv1.1 primarily to internodes (IN), and (not significantly) to paranodes (PN). n = 3 for P19 and 1 month; n = 5 for 2 months; n = 7 for>9 months. Statistics: means ± s.e.m, two-way ANOVA followed by Student’s T-test. *p<0.05; **p<0.01; ***p<0.001; n.s., not significant. (C, D) Western blot analysis of sciatic nerve lysates of 9-month-old mice shows increased Kv1.1 protein abundance in mutant nerve lysates. β-III Tubulin served as loading control for normalization (n = 4). (E–G) Kv1-anchoring proteins (green) CASPR2 and TAG-1 showed in addition to normal juxtaparanodal localization an ectopic internodal (arrow heads) and occasionally a partial overlap (asterisk) with paranodal CASPR (red) in mutant sciatic teased fibers. Both proteins colocalized at original (white arrows) and ectopic (arrow heads) positions (bottom panels). Quantification of TAG-1/Kv1.1-positive ectopic clusters is depicted, (G). Blue arrows indicate nodes of Ranvier. Scale bars, 10 µm. (H) qRT-PCR analysis (n = 5) for TAG-1 showing a tendency of increased mRNA expression in mutant compared to control nerves at 2 months. (I) Mass spectrometry of sciatic nerve lysates (n = 3) obtained at the age of 9 months shows percentage of GD1 species per genotype containing 41, 43, or 44 C-atoms (‘:’ separates number of double-bonds and ‘;’ separates number of hydroxyl-groups). Only 0.74% of GD1 in control lysates contain 41 carbons, species with more carbons are absent, while highly enriched in mutant nerve lysates. (J) Antibody-staining for gangliosides of nerves from mice aged 9 months shows GD1a+ vesicles abnormally distended into mutant internodes (green, top). GM1 is localized at paranodes of control fibers, but extends into mu internodes. Dotted lines indicate borders of myelinated fibers. Blue arrows indicate nodes of Ranvier. Scale bars, 10 µm. Statistics: means ± s.d.; *p<0.05; **p<0.01; ***p<0.001; n.s., not significant, Student’s T-test.

DOI: http://dx.doi.org/10.7554/eLife.23332.008

Immune-stained sciatic teased fibers of 2- and 9-month-old mice. (A) Nodal sodium (Nav1.6; left) and potassium (Kv7.2; right) channels are normally localized and flanked by paranodal CASPR (red, scale bars, 5 µm). (B, C) Ion channel anchoring proteins βIV-spectrin (green) (B) and ankyrinG (green) (C) are normally localized to nodes of Ranvier and flanked by paranodal CASPR (red). Scale bars, 5 µm. (D) Staining for 4.1G (green), which unlike 4.1B is abundantly present at paranodes, shows normal localization (arrowheads). Potassium channel Kv1.1 clusters (red) are located at juxtaparanodes but in mutants additionally localize along internodes. (E) Triple-staining of Nav1.6 (blue), paranodal CASPR (red), and juxtaparanodal potassium channels Kv1.1 (green) shows preservation of Kv1.1 at juxtaparanodes and at mesaxonal lines (arrows) of mutant fibers. In addition, ectopic internodal Kv1.1 patches are depicted. Scale bar, 10 µm.

DOI: http://dx.doi.org/10.7554/eLife.23332.009

GM1-labeling by CTB (red) of sciatic teased fibers from mice aged 9 months shows a defined paranodal staining in control (left) but is more distended in mutant fibers (right). Area in rectangles (top) is magnified in bottom panels. Axons are stained for TUJ1 (green). CTB, fluorescently tagged subunit B of cholera toxin.

DOI: http://dx.doi.org/10.7554/eLife.23332.010

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).

Figure 2—figure supplement 2.

Abnormal distribution of GM1 ganglioside in the perinodal region of mutant nerves by CTB-staining.

GM1-labeling by CTB (red) of sciatic teased fibers from mice aged 9 months shows a defined paranodal staining in control (left) but is more distended in mutant fibers (right). Area in rectangles (top) is magnified in bottom panels. Axons are stained for TUJ1 (green). CTB, fluorescently tagged subunit B of cholera toxin.

DOI: http://dx.doi.org/10.7554/eLife.23332.010

Figure 3.

Hallmarks of lysosomal storage disorders in Pex5 mutant nerves.

(A) Double-staining of sciatic teased fibers displayed giant GD1a+ (green) vesicles that colocalized with LAMP1 (red) in internodes only of mutant animals (rectangular inset magnified in right panels). Dotted lines indicate borders of myelinated fibers; scale bar in inset, 5 µm. (B) Electron microscopy of a teased and sectioned mutant sciatic nerve depicting paranodal vesicle accumulations. Scale bar, 500 nm. (C) Immuno-electron microscopy of sciatic nerves from mice aged 9 months identifies enlarged LAMP1+ vesicles (arrows) within paranodal loops (PNLs). M, myelin; Ax, axon. Scale bar, 500 nm. (D, E) Immunolabeling of teased sciatic nerves at the age of 2 months shows enlarged LAMP1+ (left, green) or LIMP-2+ (right, green) compartments close to CASPR+ paranodes (red) of mutant nerves. Scale bars, 10 µm. (F–H) Analysis of sciatic nerve lysates by Western blotting (n = 4; β-III Tubulin, served as loading control for normalization) and qRT-PCR (n = 6) for LAMP1 and LIMP-2 showing increased protein, but decreased mRNA levels in mutants. (I) Immunostaining of control sciatic fibers shows close association of peroxisomal PMP70 (green) and LAMP1 on lysosomal membranes (red) in the paranodal region, suggesting physical interactions. A wide-field flourescent image after deconvolution (250 nm z-stack width) is shown. Scale bar, 10 µm. Dotted lines indicate borders of the myelinated fiber. (J) Lysosomal enzyme activities of nerve lysates was enhanced in mutants compared to controls as assessed by assays using substrates of α-Mannosidase (left) and β-Hexosaminidase (right; n = 4). Statistics: means ± s.e.m. *p<0.05; **p<0.01; ***p<0.001; n.s., not significant, Student’s T-test (G, H, J). Blue arrows indicate nodes of Ranvier (D, E, I).

DOI: http://dx.doi.org/10.7554/eLife.23332.011

DOI: http://dx.doi.org/10.7554/eLife.23332.012

(A–C) Immunofluorescence showing normal abundance of early/late endosomal and autophagosomal vesicles (green) in the vicinity of nodes of Ranvier (indicated by paranodal CASPR or blue arrow) of teased fibers obtained from 9-month-old animals. RAB7, Ras-related protein 7 (late endosomes; top); EEA1, early endosome-associated protein 1 (middle); ATG5, autophagy-related protein 5 (autophagosomes; bottom). Scale bars, 10 µm.

DOI: http://dx.doi.org/10.7554/eLife.23332.013

Lysosomal accumulation within paranodal loops.

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.

Figure 3—figure supplement 1.

Characterization of paranodal vesicle inclusions in Pex5 mutant nerves.

(A–C) Immunofluorescence showing normal abundance of early/late endosomal and autophagosomal vesicles (green) in the vicinity of nodes of Ranvier (indicated by paranodal CASPR or blue arrow) of teased fibers obtained from 9-month-old animals. RAB7, Ras-related protein 7 (late endosomes; top); EEA1, early endosome-associated protein 1 (middle); ATG5, autophagy-related protein 5 (autophagosomes; bottom). Scale bars, 10 µm.

DOI: http://dx.doi.org/10.7554/eLife.23332.013

Theoretically, complete peroxisomal failure in Cnp-Cre::Pex5 mice could perturb many aspects of SC metabolism in the observed neuropathy. However, we found that Cnp-Cre::Hsd17b4 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 Kv1.1 clusters (Figure 4—figure supplement 1a–e), confirming the important role for glial lipid metabolism in maintaining axonal membrane composition, and function.

Figure 4—figure supplement 1.

Ectopic, internodal Kv1-channel clusters in nerves of conditional MFP2 knockout mice.

(A) Fluorescent image of aged (19 months) Cnp-Cre::Mfp2 (MFP2 cKO) sciatic teased nerve fibers show normal distribution of nodal Nav1.6 (blue), paranodal CASPR (red), and juxtaparanodal Kv1.1 (green), as in controls. Mutant fibers exhibit in addition ectopic internodal Kv1.1 clusters (arrow head). Scale bar, 10 µm. (B) Quantification of ectopic Kv1.1 channel distribution at paranodes (PN) or internodes (IN). Statistics: means ± s.e.m. n = 3; *p<0.05; **p<0.01; ***p<0.001, Student’s T-test. (C) Enzyme assay measuring lysosomal β-Hexosaminidase in nerve lysates from animals aged 16 months showing increased activity in mutant nerve lysates. (D, E) Functional impairment of MFP2 mutant compared to control sciatic nerves assessed by ex vivo electrophysiology at age 16 months (n = 6). Mfp2 mutant nerves display smaller responses (red curves) as compared to controls (black curves) when elicited at stimulus intensities of 0.15mA (left) or 0.155mA (right). Peak amplitudes plotted against increasing stimulus intensity indicate reduced compound action potentials of MFP2 mutant nerves (red dots) compared to controls (black dots) at all intensities measured. Statistics: Two-way ANOVA, effect of genotype, F(1200)=27.37; p=0.

DOI: http://dx.doi.org/10.7554/eLife.23332.015

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 Kv1.1 positive channels as observed in both conditional (Pex5 and Mfp2) mutants. The quantification showed that mislocalization of internodal Kv1.1 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).

Figure 4.

Normal nerve morphology, but ectopic ion channels and lysosomal accumulates in 22 months aged Abcd1 null mutants.

(A–C) Analysis of nerve morphology using entire methylene blue-stained semithin cross-sections (A, C) and g-ratios as measure of myelin thickness by electron microscopy of sciatic nerves (≥200 randomly chosen axons per nerve) obtained from control and Abcd1 knockout mice (n = 4). Scale bar, 10 µm. (D, E) Immune-stained teased sciatic nerve fibers obtained from mutants and controls with corresponding quantifications show progressively abnormal Kv1.1 localization (green), while Nav1.6 (blue) and CASPR (red) are normally localized. Scale bars, 10 µm. (n = 4, 2 mo; n = 6, 22 mo). Statistics: means ± s.e.m, Student’s T-test *p<0.05; **p<0.01; ***p<0.001; n.s., not significant. (F–H) Analysis of lysosomes reveals LAMP1+ and LIMP-2+ accumulations (green) by immune-stained sciatic teased fibers (n = 3) and a mild increase of lysosomal β-Hexosaminidase activity measured using sciatic nerve lysates (n = 4). Blue arrows indicate nodes of Ranvier. Scale bars, 10 µm. Statistics: means ± s.e.m, two-way ANOVA followed by Bonferroni (B) or Student’s T-test (H). *p<0.05; **p<0.01; ***p<0.001; n.s., not significant.

DOI: http://dx.doi.org/10.7554/eLife.23332.014

(A) Fluorescent image of aged (19 months) Cnp-Cre::Mfp2 (MFP2 cKO) sciatic teased nerve fibers show normal distribution of nodal Nav1.6 (blue), paranodal CASPR (red), and juxtaparanodal Kv1.1 (green), as in controls. Mutant fibers exhibit in addition ectopic internodal Kv1.1 clusters (arrow head). Scale bar, 10 µm. (B) Quantification of ectopic Kv1.1 channel distribution at paranodes (PN) or internodes (IN). Statistics: means ± s.e.m. n = 3; *p<0.05; **p<0.01; ***p<0.001, Student’s T-test. (C) Enzyme assay measuring lysosomal β-Hexosaminidase in nerve lysates from animals aged 16 months showing increased activity in mutant nerve lysates. (D, E) Functional impairment of MFP2 mutant compared to control sciatic nerves assessed by ex vivo electrophysiology at age 16 months (n = 6). Mfp2 mutant nerves display smaller responses (red curves) as compared to controls (black curves) when elicited at stimulus intensities of 0.15mA (left) or 0.155mA (right). Peak amplitudes plotted against increasing stimulus intensity indicate reduced compound action potentials of MFP2 mutant nerves (red dots) compared to controls (black dots) at all intensities measured. Statistics: Two-way ANOVA, effect of genotype, F(1200)=27.37; p=0.

DOI: http://dx.doi.org/10.7554/eLife.23332.015

Myelin TAG-1 is stabilized by gangliosides (GS, light green) within juxtaparanodes (JXP). For normal turnover (left) of JXP domains, SC require lysosomes (purple) and peroxisomes (dark green) that degrade GS (indicated by curved pink arrow). When GS degradation is perturbed (right), indicated by impaired peroxisomes (red), GS accumulate within enlarged lysosomes (purple), storage vesicles (gray), and likely within JXP. Excess GS within JXP stabilize the protein complex containing TAG-1 (yellow), CASPR2 (red) and Kv1.1 (blue) proteins, which thereby escape normal turnover, leading to domains breaking-off and drifting into internodes. Extra clusters possibly allow more K+ (blue dots) efflux (curved gray arrow).

DOI: http://dx.doi.org/10.7554/eLife.23332.016

Discussion

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 Kv1-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 Kv1 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 Kv1 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 Kv1-/CASPR2-/TAG-1 clusters, but this cannot be distinguished by TAG-1 immunofluorescence.

Figure 4—figure supplement 2.

Hypothetical model for lipid turnover of the juxtaparanodal anchoring complex of Kv1-channels.

Myelin TAG-1 is stabilized by gangliosides (GS, light green) within juxtaparanodes (JXP). For normal turnover (left) of JXP domains, SC require lysosomes (purple) and peroxisomes (dark green) that degrade GS (indicated by curved pink arrow). When GS degradation is perturbed (right), indicated by impaired peroxisomes (red), GS accumulate within enlarged lysosomes (purple), storage vesicles (gray), and likely within JXP. Excess GS within JXP stabilize the protein complex containing TAG-1 (yellow), CASPR2 (red) and Kv1.1 (blue) proteins, which thereby escape normal turnover, leading to domains breaking-off and drifting into internodes. Extra clusters possibly allow more K+ (blue dots) efflux (curved gray arrow).

DOI: http://dx.doi.org/10.7554/eLife.23332.016

Kv1-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 Kv1 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 Kv1.1 channel protein. Also, the ectopic Kv1.1+ 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 Kv1 into paranodes, is the underlying cause of conduction slowing. In contrast, the progressive shift of Kv1 toward internodes correlates well with disease progression. Theoretically, Kv1-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 Kv1.1 function (Zhou et al., 1998). On the other hand, a ‘gain-of-function’ effect by opening additional ectopic internodal Kv1 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 Kv1 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.

Materials and methods

Animals

Cnp1 and Pex5 mice with C57/Bl6 genetic background were genotyped as described (Kassmann et al., 2007). To check for recombination, genomic DNA was isolated from tail biopsies using the nexttec kit according to the manufacturer’s instructions. PCR was performed with sense (5’- CCAACGTCTGCCCATTCCTCCACCTG-3’) and antisense primers (5’- TTTGAGGATGGGAAGCAGTGCT −3’) generating a 330 bp amplicon after Cre-mediated excision of the floxed Pex5 gene in conditional knockout mice. Hsd17b4 (Mfp2 mice and Abcd1 knockout mice with C57/Bl6 genetic background were genotyped by PCR as described (Verheijden et al., 2013; Forss-Petter et al., 1997). Animals were maintained in individually ventilated cages under SPF conditions.

Immunofluorescence

For visualization of gangliosides sciatic nerves of Cnp-Cre::Pex5 mice and controls were dissected and pre-teased in artificial cerebrospinal fluid (ACSF; 126 mM NaCl, 3 mM KCl, 1,25 mM NaH2PO4, 26 mM NaHCO3, 2 mM MgSO4, 10 mM Glucose, 2 mM CaCla2). Fiber bundles were incubated for 1.5 hr at room temperature with antibodies specific for GM1 or GD1a (kindly provided by Hugh Willison, Glasgow). Samples were washed in ACSF, incubated with isotype-specific fluorescent secondary antibodies (Alexa-488 IGg3 and IG2b goat anti-mouse; Thermo Fisher Scientific, Massachusetts, USA), again washed in ACSF, fixed in 4% paraformaldehyde (PFA) for 5 min, then washed in PBS containing 0.1 M glycine, teased on glass slides, air-dried, and stored at −20°C. For other antibody-labeling, sciatic nerves were dissected, teased and stored at −20°C as described previously (Kassmann et al., 2011). For staining protein antigens, frozen teased fibers were fixed in 4% paraformaldehyde (PFA) for 5 min, permeabilized in methanol at −20°C for 3–5 min, PBS-washed, and blocked for 1 hr in PBS containing 10% horse serum and 0.05% Triton X-100. Primary antibodies diluted in blocking solution were applied overnight at 4°C, specimens were washed in PBS, and incubated for 1 hr with fluorescent secondary antibodies Alexa-488, Alexa-555 (Thermo Fisher Scientific, 1:2000), and DyLight 633 (1:1000, YO Proteins, Sweden). Cell nuclei were visualized with 4′, 6′- diamidino-2-phenylindole (DAPI; 1:10.000, Thermo Fisher Scientific). Fibers were mounted in AquaPolymount (Polysciences, Pennsylvania, USA). Fluorescent images were acquired with an inverted Zeiss Axio Observer.Z1 equipped with an Axiocam MRm (Zeiss, Germany).

Immunohistology

Mouse sciatic nerves were fixed overnight in 4% PFA, embedded in paraffin, and longitudinally sectioned (5 µm). For labeling, Dako LSAB2 system (Dako, Denmark) was used according to the manufacturer’s directions. All samples were analyzed by light microscopy (Zeiss Axiophot).

Semithin sections and electron microscopy

Mice were anesthetized with avertin (100 μl/10 g bodyweight) and perfused intracardially with 15 ml HBSS (Lonza, Switzerland), followed by 30 ml of fixative (0.2% glutaraldehyde, 4% paraformaldehyde in PBS). Sciatic nerves were dissected and post-fixed for 2–4 hr in 4% followed by overnight incubation in 1% PFA. Epon embedding of sciatic nerves was performed as described previously (Kassmann et al., 2007).

Semithin cross-sections (500 nm) of sciatic nerves were incubated with methylene blue/Azur II and axons of entire cross-sections of sciatic nerves were analyzed by light microscopy (Zeiss Axiophot). Quantification of axon number was performed using Fiji software.

Preparation of ultrathin cryosections (50 nm) was performed according to the Tokuyasu technique, and immunolabeling was carried out as previously described (Werner et al., 2007). Antibody binding was visualized with protein A-gold (1:60, 10 nm).

Ultrathin sections (50 nm) were contrasted with 1% uranyl acetate and lead citrate. Sections were examined with a LEO EM 912AB electron microscope (Zeiss), and pictures were taken with an on-axis 2048 X 2048 CCD camera (Proscan). Myelin thickness was evaluated using Fiji software by g-ratio analysis (dividing the axonal by the fiber diameter) derived from at least 100 randomly chosen fibers per nerve.

Protein biochemistry

Sciatic nerves were transferred to 200 µl (per nerve) ice-cold lysis buffer containing 1% Triton X-100, 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% NP-40, and protease inhibitors (Complete, Roche, Switzerland). Samples were homogenized using Precellys 24 (VWR International, Dealware, USA), centrifuged for 10 min at 1000 g to remove cellular nuclei, and protein concentration of the supernatant was determined using Lowry assay (Bio-rad Laboratories, California, USA). Blotting membranes were incubated overnight at 4°C with primary antibodies. Western Lightning + ECL Kit (Perkin Elmer, Massachusetts, USA) was used to label protein bands, which were detected using Intas ChemoCam Image system. At least four biological replicates were analyzed. Quantification was performed in Fiji.

Antibodies

4.1G (1:100; kindly provided by E. Peles) AnkG (1:100; kindly provided by M. Rasband), APP (1:1000; MAB348, Merck Millipore, Massachusetts, USA), ATG5 (1:100; pab50264 Covalab, France), CASPR (mk, m; 1:1000; clone K65/35 NeuroMab, California, USA), CASPR2 (1:500; kindly provided by E. Peles), Catalase (1:200; C0979 Sigma-Aldrich, Missouri, USA), CD3 (1:150; MCA1477 Serotec, MorphoSys, Germany), EEA1 (1:300; ab2900 Abcam, Cambridge, United Kingdom), Kv1.1 (1:50; sc11184 Santa Cruz Biotechnology, Texas, USA), Kv1.1 (1:50; clone 20/78 NeuroMab), Kv7.2 (1:2000; PA1-929 ThermoSicentific), LAMP1 (WB, 1:400; IF, 1:200; IEM, 1:200; 553792 BD Bioscience, New Jersey, USA), LIMP-2 (WB, 1:250; IF, 1:2000; kindly provided by J. Blanz), MAC-3 (1:400; 553322 BD Bioscience), Nav1.6 (1:500; ASC-009 Alomone labs, Israel), NF155 (1:1000; kindly provided by P. Brophy), P0 (1:1000 [Archelos et al., 1993]); P2 (1:500; sc-49304 Santa Cruz), PLP (1:5000 [Jung et al., 1996]), PMP22 (1:1000; SAB4502217 Sigma), PMP70 (1:600; ab3421 Abcam), RAB7 (1:300; R4779 Sigma), TAG-1 (1:500; kindly provided by E. Peles), ßiv spectrin (1:400; kindly provided by M. Rasband), α-Tubulin (1:1000; T 5168 Sigma), III β-Tubulin (1:1000; Covance, New Jersey, USA).

Lysosomal enzyme activity assay

Sciatic nerves were homogenized using Precellys 24 (twice for 20 s at 5000 rpm) in 150 µl ice-cold lysis buffer containing 10 mM Tris-HCl pH 7.4, 150 mM NaCl, 5 mM EDTA, 0.5% Triton X-100, 1 x PEFA, and Protease inhibitors (Complete, Roche). Nerve lysates were centrifuged for 10 min at 15,000 g to remove cellular debris and nuclei. The supernatant was incubated with 10 mM of the substrate (i.e. p-nitrophenyl-a-D-mannopyranosid or p-Nitrophenyl-N-acetyl-ß-D-glucosaminide; Sigma) dissolved in 0.2 M citrate buffer, and the reaction was stopped by addition of 500 µl 0.4 M glycine NaOH (pH 10.4; adjusted with 1 M NaOH) to stop enzyme activity. Samples were centrifuged for 10 min at 15,000 g, and the supernatants of two technical replicates were measured at 405 nm with an Eon microplate spectrophotometer (BioTek, Vermont, USA).

Quantitative real-time PCR

Total RNA was extracted from sciatic nerve lysates as described previously (Fünfschilling et al., 2012). Reactions were performed with four technical replicates of nerves obtained from six different animals per group.

PCR primers were specific for β-actin (forward 5′-CTTCCTCCCTGGAGAAGAGC and reverse 5′-ATGCCACAGGATTCCATACC), Lamp1 (forward 5’-CCTACGAGACTGCGAATGGT and reverse 5’-CCACAAGAACTGCCATTTTTC), Limp-2 (forward 5’-TGGAGATCCTAACGTTGACTTG and reverse 5’-GGCCAGATCCACGACAGT), and Pex5 (forward 5’-CACATCCGCTTCCTATGACA and reverse 5’-AAAAGGCTGAGGGTGGTCA).

Mass spectrometry

To quantify PE (diacyl), PE O- (alkanyl-acyl), and PE P- (alkenyl-acyl) species, acidic (PE and PE O-) or neutral (PE P-) lipid extractions were performed as described (Bligh and Dyer, 1959). Typically, 1–5 µl of a 1:10 dilution of total lysates were measured. As lipid standards, 50 pmol of PE standards and 40–60 pmol of a PE P- standards were used. Standard syntheses were performed as described (Özbalci et al., 2013; Paltauf and Hermetter, 1994). After solvent evaporation under a gentle nitrogen flow at 37°C, lipids were re-suspended in 50 µl 10 mM ammonium acetate in methanol. PE/PE O- and PE P- species were analyzed in neutral loss or precursor ion mode selecting for class-specific fragments on a QTRAP5500 (Sciex, Massachusetts, USA) equipped with a NanoMate devise (Advion, New York, USA), employing MS settings as described (Özbalci et al., 2013). Data evaluation was done using LipidView (AB Sciex) and an in-house-developed software (ShinyLipids). Species annotated as PE O- mainly contain plasmenylethanoamines but also minor amounts of PE P- species and PE species with odd-numbered fatty acids. Extraction of gangliosides was performed as described (Sampaio et al., 2011), with the exception that the first neutral extraction was performed using chloroform:methanol as a 17:1 (v/v) solution, followed by a chloroform:methanol 2:1 (v/v) extraction. For quantification of gangliosides, GD1a and GD1b 50 pmol N-CD3-Stearoyl GM3 and N-CD3-Stearoyl-GM1 (Matreya, Pennsylvania, USA) were used as internal ganglioside standards. Following evaporation, samples were re-suspended in 100 µl methanol. Gangliosides were subjected to UHPLC-MS analysis, using a CSH C18 column (1 × 150 mm, 1.7 µm particles, Waters) coupled to a QExactive high-resolution Orbitrap mass spectrometer (Thermo) equipped with an ESI source.

For GD1a/b quantification, 30 µl aliquots of the aqueous and the chloroform:methanol (2:1) phase were transferred to Eppendorf cups, evaporated and resuspended in 50 µl buffer containing 60% of mobile phase A (acetonitrile:water; 60:40 (v/v) with 10 mM ammonium formate and 0.1% formic acid) and 40% mobile phase B (isopropanol:acetonitrile, 90:10 (v/v) with 10 mM ammonium formate and 0.1% formic acid). Of each sample, 10 µl was subjected to UPLC separation (Dionex, California, USA), using a multistep gradient. Full MS scans were acquired for 30 min in negative ion mode (600–1800 m/z) with automatic gain control target set to 1 × 106 ions. The maximum injection time was set to 200 ms and a FHWM resolution of 140,000 (at m/z 200). In addition to Full MS scans, all ion fragmentation scans in negative ion mode were performed at a resolution of 70,000, scanning a mass range of 120–600 m/z with a normalized collision energy set to 30 eV.

Data evaluation of Full-MS scans (profile spectra) was performed using MassMap (MassMap, Germany). Therefore, data files obtained were converted to mzXML-files using the software Proteowizard. mzXML-Files were then converted to mmp-Files using the LC-MS data evaluation software MassMap.

VLCFA determination was performed using 10 µl of a 1:10 dilution of total membrane fractions. 500 pmol of C25 fatty acid (Dr. Ehrenstorfer GmbH, Germany) and C23 fatty acid (Matreya) was used as internal standards. Prior to extraction, glass tubes where washed with chloroform containing 1% acetic acid. The samples were resuspended in 80 µl of toluene. 1 ml acetonitrile-hydrochloric acid (conc.) (4:1) was added and incubated for 2 hr at 90°C. After cooling to room temperature, 2 ml hexane were added, the samples were vortexed, incubated for 5 min and centrifuged for 2 min at 2000 rpm. The upper phase was transferred to a new glass tube. After evaporation of the organic phase under a gentle stream of nitrogen, lipid extracts were resuspended in 100 µl chloroform:methanol:water (50:45:5, v/v/v) with 0.1% ammonium hydroxide solution. Of each sample, 5 µl was transferred to a 96-well plate containing 10 µl 0.01% piperidine in methanol and 5 µl MeOH. Samples were subjected to mass spectrometric analysis on a QSTAR Elite (Sciex) instrument equipped with a NanoMate (Advion). TOF scans were performed over a mass range of 100–500 m/z in negative ion mode. Data sets were processed and evaluated using LipidView (Sciex).

Electrophysiology, ex vivo

Following cervical dislocation mouse sciatic nerves were rapidly transferred into a perfusion chamber filled with gassed ACSF at 37°C containing in mM: 126 NaCl, 3 KCl, 26 NaHCO3, 1.25 NaH2PO4, 25 glucose, 2.5 MgSO4 and 2 CaCl2; a pH of 7.4 was ensured by continuous gassing with carbogen (95% O2/5% CO2). Nerves’ lengths were measured and were then allowed to adapt for approximately 30 min before electrophysiological recordings began. Suction electrodes backfilled with ACSF were used for stimulation and recording. Electrical stimulation was performed at different intensities ranging between 0.13 mA and 0.3 mA, and increased manually in step-width of 0.05 mA. Inter-stimulus intervals were 3 s and each of the stimulus intensities was used 10 times. CAPs were continuously measured. Recorded signals were acquired at 100 kHz, amplified x100 by Ext-2F (NPI Electronic, Germany), and a further 50-fold by SR560 (Stanford Research Systems, California, USA), and filtered. Recordings were controlled by patchmaster software (HEKA, Germany) with an EPC9 amplifier interface (HEKA). Stimulus intensities were altered using Stimulus Isolator A385 (World Precision Instruments).

Analysis of electrophysiology was performed using Matlab. Each nerve was analyzed separately. Responses triggered but the same stimulus repetitions were averaged together. Nerve conduction velocity was calculated for each of the stimulus intensities as the mean across trials of the ratio between the length of the nerve and the time between peaks of stimulus-artifact (first small positive peak) and response (third positive peak).

Electrophysiology, in vivo

Mice aged 9 months were anesthetized with ketaminhydrochloride (100 mg kg−1)/xylazin hydrochloride (8 mg kg−1). Steel needle electrodes were placed subcutaneously, one pair at sciatic notch (proximal stimulation), a second pair at the tibial nerve above the ankle (distal stimulation). Supramaximal square wave pulses lasting 100 ms were delivered using a Toennies Neuroscreen (Jaeger, Germany). Compound muscle action potential (CMAP) was recorded from the intrinsic foot muscles.

Image processing

Digital images were processed and quantified with ZEN 2012 software (Zeiss) and/or Fiji.

Statistical analysis

All numerical values are shown as the mean ± s.e.m.; n = 3–6, unless stated otherwise. Statistical significance was determined using GraphPad Prism5 by two-way ANOVA or the two-tailed Student's t test for unpaired samples assuming unequal variance and p values below 0.05 were marked as significant: *p<0.05; **p<0.01; ***p<0.001). Normal distribution was assumed, but not formally tested.

Study approval

Mice were kept in the mouse keeping facility of the Max-Planck-Institute of Experimental Medicine at 12 hr light/dark cycle. All experiments were executed according to the German Animal Protection Law and approved by the government agency of the State of Lower Saxony, Germany.

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Peroxisomal dysfunctions cause storage of lysosomes and axonal Kv1 channel redistribution in peripheral neuropathy" for consideration by eLife. Your article has been favorably evaluated by Gary Westbrook (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors. The reviewers have opted to remain anonymous. The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

In this short report, Kleinecke et al. describe the consequences of peroxisome dysfunction upon Pex5 deletion in Schwann cells. The phenotype is strikingly less severe than that in that observed when Pex5 is deleted in oligodendrocytes as described in a previous paper from this group, and interestingly suggests peroxisome dysfunction has severe consequences on the function of neighboring lysosomes. Indeed, the authors demonstrate that there is mislocalization of lysosomal proteins, gangliosides and other proteins including Kv1.1 channels. The authors suggest that these changes have important effects on nerve physiology. Lastly, some of the observed phenotypes were also observed in mice deficient in Abcd1. The authors propose an interesting model that suggests lipids on Schwann cell membranes stabilize certain adhesion proteins and their associated ion channels. Disruption of these lipids leads to Kv1.1 channel mislocalization which in turn is proposed to lead to nerve conduction changes.

Essential revisions:

Although the presented data suggest very interesting models, the correlative nature of this data could be strengthened by additional experiments.

1) The examination of conditional knockouts for the peroxisomal protein Mfp2 for similar phenotypes that were observed in Pex5 cKOs would strengthen the conclusions and provide some mechanistic insight. For example, if many of the findings observed in the Pex5 cKOs, particularly the abnormal CAP recordings, could also be demonstrated in the Mfp2 cKOs, it could be said with greater confidence that manipulation of lipid metabolism is specifically producing these changes. Similarly, the connection to human disease could be strengthened if the CAP recording abnormalities are also found in Abcd1 KOs.

2) Additional studies on nerve physiology are needed. The studies on nerve conduction show clear changes in conduction velocity and CAP amplitude. The authors conclude this is a function of ectopic internodal K+ channels leading to conduction block. Some additional experiments are needed to support these conclusions. Decreased internodal distance could also decrease conduction velocity, whether this could be a progressive change is unclear, but it should be ruled out. Conduction block is typically defined clinically by stimulating at a distal point and a proximal point, and comparing amplitudes. If the distal amplitude is greater than the proximal, this is evidence of conduction block (increased failure to propagate and action potential with increased distance). This should be tested. It is unclear that the decrease in CAP could not be simply the result of lower amplitude action potentials, or a complete failure in excitability of a subset of axons (larger, faster conducting axons, for example, which could also explain the velocity changes).

3) Furthermore, examination of the localization of Kv1.1 channels should be done at an earlier time point. It's not clear from the data whether membrane complexes break off and diffuse into the adjacent internodal and paranodal regions, or if localization is simply misspecified during the initial formation of myelination. Although the progressive mislocalization of Kv1.1 channels to the internode with age supports the authors' model, it would be helpful to examine an earlier time point when sciatic nerve myelination has just completed.

4) Lastly, to strengthen the proposal that ganglioside accumulation underlies mislocalization of Kv1.1, quantitation of the GPI-anchored protein TAG-1 and CASPR2 would be useful to rule out for possible alterations in expression that may be altered by lipid metabolism.

Essential revisions:

Although the presented data suggest very interesting models, the correlative nature of this data could be strengthened by additional experiments.

1) The examination of conditional knockouts for the peroxisomal protein Mfp2 for similar phenotypes that were observed in Pex5 cKOs would strengthen the conclusions and provide some mechanistic insight. For example, if many of the findings observed in the Pex5 cKOs, particularly the abnormal CAP recordings, could also be demonstrated in the Mfp2 cKOs, it could be said with greater confidence that manipulation of lipid metabolism is specifically producing these changes. Similarly, the connection to human disease could be strengthened if the CAP recording abnormalities are also found in Abcd1 KOs.

We agree that a similar pathology of PEX5 and MFP2 cKO nerves would strengthen our conclusions. We have therefore analyzed sciatic nerves from aged (16 months old) MFP2 mutants, which demonstrated nearly 3-fold lysosomal enzyme activity (new Figure 4—figure supplement 1C). Moreover, electrophysiological recordings revealed reduced CAPs of mutants compared to age-matched controls, as predicted. (new Figure 4—figure supplement 1D).

We also performed the requested electrophysiological recordings of ABCD1 animals. At age 2 months, there were no differences yet compared to controls. This was not unexpected, because the mislocalization of Kv1 potassium channels at that age affected only a minor portion of internodes (original Figure 4E). We note, that in human AMN patients the corresponding PNS symptoms will not manifest before the 2nd to 3rd decade of life, and it is well known that mouse and man differ in disease expression and disease kinetics (specifically for mutations of the X-linked ABCD1 gene; Ferrer et al., Brain Pathol., 2010). Indeed, ABCD1 knockout mice showed a significant mislocalization of potassium channels only at 22 months of age (original Figure 4E), which corresponds to the human 7th – 8th decade of life. At that age we measured reduced compound muscle action potential (CMAP) compared to younger mutants, however there was already a major overlap with other aging-dependent changes of conductivity that also affect wild-type mice. Thus, it was not possible to simply separate nerve aging effects from ABCD1 genotype-dependent changes of conductivity. Although, we noted a clear trend (P=0.12) between the groups, we prefer to not add these data into the current manuscript. Waiting for another cohort of ABCD1 mutant with the age of 22 months is unfortunately not possible within the time frame of the revision.

2) Additional studies on nerve physiology are needed. The studies on nerve conduction show clear changes in conduction velocity and CAP amplitude. The authors conclude this is a function of ectopic internodal K

As requested, we have addressed the possibility that conduction blocks are responsible for the reduced CAPs in PEX5 mutant nerves. Here, we have turned to in vivo analysis and have assessed the ratio of compound muscle action potentials (CMAP) after proximal and distal nerve stimulation. As shown in new Figure 1—figure supplement 2C, the distal amplitude was significantly (p<0.01) greater than the proximal amplitude, as predicted for axonal conduction blocks. A sentence referring to these new data has been added in the third paragraph of the Results.

To investigate a possible impact of altered internodal length on conduction velocity (NCV) we analyzed PEX5 mutant nerves at age 2 months. Here, the average internodal length of mutant fibers (623 µm) was approximately 10% shorter compared to controls (691 µm), as summarized in the fourth paragraph of the Results and depicted in new Figure 1—figure supplement 3D. Although significant, this length difference should by itself not result in slowed conduction, because shortening the internodal length accelerates nerve conduction to a flat maximum, as shown by the Brophy lab. That paper demonstrated unaltered NCV despite a 30% reduced internodal length in the ΔPDZ-Prx mouse mutant (Wu et al., Curr. Biol., 2012).

3) Furthermore, examination of the localization of K

For the original submission we had analyzed Kv1.1 in PEX5 mutant and control nerves at P19 (original Figure 2), i.e. shortly after juxtaparanodes have formed in development. As requested, we have extended the observational time points to P17 and then to P7.

At P7 and also at P17 Kv1.1 appeared not yet completely clustered, and in the majority of myelinated fibers (mutants and controls) the staining included a diffuse signal in nodal/paranodal and paranodal/ juxtaparanodal regions. In these immature fibers that lacked regular Kv1.1-positive juxtaparanodes, we never observed abnormal internodal clusters.

Likewise, in ABCD1-mutant nerves a comparison of young (2mo) and old (22 mo) sciatic nerves revealed significant internodal clusters only at age 22 months (as shown in Figure 4E). This strengthens our model of an initially normal formation of juxtaparanodes and a secondary breaking off. We have added a clarifying sentence in the last paragraph of the Results.

4) Lastly, to strengthen the proposal that ganglioside accumulation underlies mislocalization of K

We agree that alterations of a cell's lipid metabolism could affect (e.g. SREBP-dependent) gene expression and result in overexpression as a hypothetical cause of ectopic protein localization. We have therefore analyzed the mRNA expression levels of TAG-1, which is the Kv1.1 anchor expressed by Schwann cells, but could not detect significant differences (p=0.42) as shown in NEW Figure 2H. Note that CASPR2 is expressed in the neuronal/axonal compartment.