Published in Volume
124, Issue 6
(June 2, 2014)J Clin Invest.
Copyright © 2014, American Society for Clinical
Peripheral nervous system plasmalogens regulate Schwann cell differentiation
1Nerve Regeneration Group, Instituto de Biologia Molecular e
Celular (IBMC), Porto, Portugal. 2Instituto de Ciências Biomédicas
Abel Salazar (ICBAS), University of Porto, Porto, Portugal. 3Biology
Department, Boston College, Chestnut Hill Massachusetts, USA. 4Academic
Medical Center, University of Amsterdam, Amsterdam, The Netherlands.
5Biochemistry Center Heidelberg, University of Heidelberg, Heidelberg,
Address correspondence to: Pedro Brites, Instituto de Biologia Molecular e Celular,
Nerve Regeneration, Rua do Campo Alegre 823, 4150-180, Porto, Portugal. Phone:
351226074900; Fax: 351226099157; E-mail: firstname.lastname@example.org.
Authorship note: Tiago Ferreira da Silva, Jessica Eira, and André T.
Lopes contributed equally to this work.
First published April 24, 2014
Rhizomelic chondrodysplasia punctata (RCDP) is a developmental disorder characterized by
hypotonia, cataracts, abnormal ossification, impaired motor development, and intellectual
disability. The underlying etiology of RCDP is a deficiency in the biosynthesis of ether
phospholipids, of which plasmalogens are the most abundant form in nervous tissue and
myelin; however, the role of plasmalogens in the peripheral nervous system is poorly
defined. Here, we used mouse models of RCDP and analyzed the consequence of plasmalogen
deficiency in peripheral nerves. We determined that plasmalogens are crucial for Schwann
cell development and differentiation and that plasmalogen defects impaired radial sorting,
myelination, and myelin structure. Plasmalogen insufficiency resulted in defective protein
kinase B (AKT) phosphorylation and subsequent signaling, causing overt activation of
glycogen synthase kinase 3β (GSK3β) in nerves of mutant mice. Treatment with
GSK3β inhibitors, lithium, or 4-benzyl-2-methyl-1,2,4-thiadiazolidine-3,5-dione
(TDZD-8) restored Schwann cell defects, effectively bypassing plasmalogen deficiency. Our
results demonstrate the requirement of plasmalogens for the correct and timely
differentiation of Schwann cells and for the process of myelination. In addition, these
studies identify a mechanism by which the lack of a membrane phospholipid causes
neuropathology, implicating plasmalogens as regulators of membrane and cell signaling.
Plasmalogens, glycerophospholipids with a 1-O-alkenyl ether bond at the
sn-1 position of the glycerol backbone, represent the most abundant form of
ether phospholipids. In vitro studies have highlighted the role of plasmalogens as mediators
of the physical properties of membranes, as antioxidants, and as pools of lipid mediators
(1). The biosynthesis of ether phospholipids is
initiated in peroxisomes by the activity of glyceronephosphate O-acyltransferase (GNPAT) and
alkylglyceronephosphate synthase (AGPS) (2). Mutations
in GNPAT, AGPS, or in PEX7, which targets
AGPS in peroxisomes, lead to extremely reduced levels of plasmalogens and to rhizomelic
chondrodysplasia punctata (RCDP) (3–5). RCDP is a peroxisomal disorder characterized by
multiple tissue defects, ranging from bone, eye, and heart, to nervous tissue involvement
Nevertheless, the pathophysiology behind defective plasmalogens is poorly understood. The
phenotype of mouse models for RCDP has supported the etiology of the disease (8). Pex7- and Gnpat-KO
mice, lacking plasmalogens in all tissues, are characterized by a severe phenotype, with
early postnatal death and hypotonia in a subset of mice, whereas KO mice surviving past the
first 2 weeks display impaired growth, cataracts, and infertility (9, 10). Decreased transcript levels
of Pex7 and Agps impair ether phospholipid synthesis in
Pex7 and Agps hypomorphic mice, respectively, causing
partial decreases in plasmalogen levels. In these mutants, the residual levels of
plasmalogens are thought to prevent the hypotonia and early lethality observed in KO mice
Nevertheless, bone, lens, and testicular defects in the hypomorphic mice mirror those of KO
mice. Pex7- and Gnpat-KO mice, models of RCDP type 1 and
type 2, respectively, share a defect in ether phospholipid biosynthesis, although
Pex7-KO mice also have defects in fatty acid β- and
α-oxidation (9, 13). As in the human RCDP variants, the phenotype and tissue pathology of
Pex7- and Gnpat-KO mice are extremely similar,
highlighting that the defect in plasmalogens is the major player in disease presentation and
Plasmalogens are enriched in nervous tissue, especially in white matter, where they
localize to the cytoplasmic apposition of myelin (14,
15). Brain imaging of classical RCDP patients
indicates impaired myelination, with hypointensity on T1-weighted images and reduced levels
of choline, with increased mobile lipids and myo-inositol on MR spectroscopy (6, 16). A
plasmalogen deficiency in mice contributes to astrocytosis, microgliosis, and myelin loss in
the CNS (13). Impaired peripheral nerve conduction in
Pex7-KO mice (13) and the
description of peripheral neuropathy in a subset of RCDP patients lacking the rhizomelia and
ossification defects (17) prompted us to determine
the role of plasmalogens and the consequences of their deficiency in the peripheral nervous
system (PNS). In the PNS, Schwann cells are the glia responsible for maintaining axonal
health and myelination (18). Prior to myelination and
concomitant with Schwann cell differentiation, large-caliber axons destined for myelination
are segregated from axon bundles by Schwann cells in a process called radial sorting (19). Differentiated, promyelinating Schwann cells then
undergo transcriptional, biochemical, and morphologic changes as they initiate axonal
wrapping with a myelin sheath (20, 21). In the PNS, the extent of myelin deposition around
individual axons is driven by the axonal expression of neuregulin 1 (NRG1) and by the
signaling cascade initiated in Schwann cells expressing the ErbB receptors (22). In the adult PNS, Remak bundles are composed of
immature, nonmyelinating Schwann cells that continue to associate with bundles of
small-caliber axons that did not undergo radial sorting (23).
Here, we investigated the role of plasmalogens in myelination of the PNS. Our results
demonstrate that plasmalogens are crucial for Schwann cell differentiation, as their absence
impairs axonal sorting and myelination. Moreover, we found that plasmalogens and myelin
basic protein (MBP) are two crucial players that coordinate myelination in the PNS.
Mechanistically, we demonstrate that plasmalogens are essential for the correct
phosphorylation and activation of protein kinase B (AKT). In mutant sciatic nerves, reduced
AKT activity resulted in overt activation of glycogen synthase kinase 3β
(GSK3β). Finally, we found that the exogenous inhibition of GSK3β with lithium
or 4-benzyl-2-methyl-1,2,4-thiadiazolidine-3,5-dione (TDZD-8) rescued the impairment in
Schwann cell differentiation and axonal sorting of plasmalogen-deficient mice. Together, our
novel findings reveal the neuropathology behind a deficiency in plasmalogens and elucidate
the mechanism by which a membrane phospholipid is essential for Schwann cell differentiation
and function, implicating plasmalogens as regulators of membrane and cell signaling.
Plasmalogen deficiency causes defects in radial sorting and myelination. To assess whether Pex7- and Gnpat-KO mice had defects
in PNS development, sciatic nerves from 5- and 15-day-old (P5 and P15, respectively) mice
were analyzed by electron microscopy. In nerves from P5 WT mice, we observed axon bundles
composed of small-caliber axons (Figure 1A, arrow)
and sorted axons (Figure 1A, arrowhead) that
displayed a 1:1 relationship with Schwann cells. However, in sciatic nerves from P5
Gnpat-KO mice, the bundles contained small- and large-caliber axons
(Figure 1A, asterisks). Consistent with a failure in
the process of radial sorting, we observed an increased number of axons per bundle (Figure
1B) and a decreased number of sorted,
promyelinating fibers (Figure 1C) in nerves from P5
Pex7- and Gnpat-KO mice. Two main consequences of
impaired radial sorting were observed in nerves from Gnpat-KO mice. In
what seemed to be an attempt by Schwann cells to rescue the impaired radial sorting, we
observed an increase in Schwann cells engulfing single, small-caliber axons in nerves from
P15 Gnpat-KO mice (Figure 1A, white
arrowheads). Ultrastructural analysis of sciatic nerves from adult
Pex7-KO mice revealed a reorganization of Remak bundles, since we
observed a reduction in the number of axons in these bundles (Figure 1D), without changes in the density of these unmyelinated fibers (Figure
1E). Additionally, the presence of large-caliber
axons in Remak bundles caused Schwann cells to myelinate these structures (Supplemental
Figure 1A; supplemental material available online with this article; doi:10.1172/JCI72063DS1). Surprisingly, this abnormal myelination of Remak
bundles was not temporary. In sciatic nerves from 1.5-year-old Gnpat-KO
mice, we continued to observe myelination of Remak bundles that contained a few
large-caliber axons (Supplemental Figure 1B).
Plasmalogen deficiency causes defects in axonal sorting and myelination. (A) Electron microscopic analysis of sciatic nerves from P5 and P15 WT and
Gnpat-KO mice. Bundles in P5 Gnpat-KO nerves
contained large axons (asterisks), whereas in WT nerves these axons had been sorted
(arrowhead), and the bundles contained very small-caliber axons (arrow). At P15, sciatic
nerves from Gnpat-KO mice had Remak bundles with only 1 axon
(arrowheads). Scale bars: 2 μm. (B) Composition of axon bundles in
sciatic nerves from P5 WT, Pex7-KO, and Gnpat-KO mice.
*P = 0.031; **P = 0.011. (C) Density of
sorted axons in sciatic nerves from P5 WT, Pex7-KO, and
Gnpat-KO mice. *P = 0.003. (D)
Composition of Remak bundles in nerves from adult WT and Pex7-KO mice.
*P = 0.013. (E) Density of unmyelinated fibers (UMF) in
Remak bundles of nerves from adult WT and Pex7-KO mice.
(F) Quantification of myelin thickness by g ratio in sciatic nerves at P15.
Results are graphed as boxes with a line at the mean and whiskers from the minimal to
maximal values. *P = 0.005. (G) DRG cocultures of neurons
and Schwann cells from WT and Gnpat-KO mice stained for neuronal
βII-tubulin (green) and for the myelin protein MBP (red). Scale bars: 200
μm. (H) Density of myelin segments in DRG cocultures from WT and
Gnpat-KO mice. *P = 0.001. (I) Length
of individual myelin segments in myelinating cocultures. *P =
During postnatal development, from P5 to P20, nerves from Pex7 and
Gnpat-KO mice displayed a generalized state of hypomyelination
(Supplemental Figure 2). Determination of the g ratio revealed a defect in myelination
with decreased myelin thickness in Pex7- and
Gnpat-KO–derived nerves (Figure 1F). In vitro analysis of myelination using cocultures of dorsal root ganglia
(DRG) neurons and Schwann cells (Figure 1G) revealed
that Schwann cells from Gnpat-KO mice produced fewer and smaller myelin
segments when compared with those in WT Schwann cells (Figure 1, H and I). These findings demonstrate that deficiency in plasmalogens
impairs Schwann cell function, affecting radial sorting and myelination and leading to PNS
Plasmalogens and MBP coordinate myelination in the PNS. To further address the role of plasmalogens in myelination, we analyzed myelin content in
sciatic nerves from adult Pex7-KO mice. Surprisingly, we observed normal
myelin thickness in mutant nerves (Figure 2A),
suggesting a rescue of the defective myelination during postnatal development. However,
x-ray diffraction of fixed sciatic nerves showed an increase in myelin period of
Pex7-KO nerves (Figure 2B),
suggesting less compaction or reduced stability of myelin in the absence of plasmalogens.
The overall organization of myelinating Schwann cells in teased fiber preparations from
Gnpat-KO nerves was abnormal, with an increase in noncompact myelin due
to the increased number of Schmidt-Lanterman incisures (WT 1.7 ± 0.4 incisures/100
μm, Gnpat KO 4.7 ± 1.4 incisures/100 μm;
P = 0.0028; Figure 2C) and with
fragmented and dispersed DRP2-labeled appositions (Figure 2D).
Plasmalogens and MBP coordinate myelination. (A) Quantification of myelin thickness by g ratio in sciatic nerves from
3-month-old WT and Pex7-KO mice. Results are the mean ± SEM.
(B) X-ray diffraction analysis of sciatic nerves from WT and
Pex7-KO mice. Results are graphed as the mean ± SEM.
P = 0.01. (C) Immunofluorescence analysis of teased
fibers from adult WT and Gnpat-KO nerves stained with F-actin (red).
Arrows point to Schmidt-Lanterman incisures. Scale bars: 10 μm. (D)
Immunofluorescence analysis of sciatic nerve teased fibers stained with an antibody
against DRP2 (green) showing abnormal apposition in mutant nerves. Scale bars: 10
μm. (E) PPD-stained cross sections of sciatic nerves from P30 mice.
Scale bars: 25 μm. (F) Quantification of the degree of myelination by
g ratio in sciatic nerves from P30 mice. Results are graphed as boxes with a line at the
mean and whiskers from the minimal to maximal values. *P < 0.001.
(G) Calculated motor nerve conduction velocities (MNCV) in 3-month-old
WT, shi, Pex7-KO, Pex7:shi DM,
Gnpat-KO, and Gnpat:shi DM mice. *P
< 0.0001; #P = 0.001.
We hypothesized that the achievement of myelination in the absence of plasmalogens could
be mediated by the action of other myelin components. Studies of PNS myelin of shiverer
(shi) mice, which lack MBP, a major constituent of PNS myelin, showed
that other myelin components, namely, P0 and P2, are compensatory, allowing
shi mice to attain normal myelination and compaction (24). To further investigate whether plasmalogens were
crucial for myelination, we generated Pex7:shi and
Gnpat:shi double-mutant (DM) mice. Phenotypically, the DMs shared the
features of Pex7- and Gnpat-KO mice and the
characteristic shivering caused by MBP deficiency (data not shown). When compared with
single mutants and WT mice, nerves from Pex7:shi DM mice were
characterized by a severe hypomyelination (Figure 2E)
without axonal loss (WT 248,704 ± 15,639 axons/mm2; DM 243,884 ±
15,851 axons/mm2; P = 0.434). Myelin thickness was reduced in
Pex7-KO nerves, but the double deficiency of plasmalogens and MBP in
Pex7:shi DM mice caused a pronounced defect in myelination as judged by
the high g ratio values (Figure 2F). At the
functional level, the single mutants had defects in nerve conduction, but in DM mice, the
combined deficiency of MBP and plasmalogens affected nerve conduction by less than half
the normal values (Figure 2G). These findings
indicate that in the absence of plasmalogens, the presence of normal amounts of MBP
(Supplemental Figure 2B) is sufficient to achieve normal amounts of myelin. Our results
highlight the possible coordination between membrane phospholipids and myelin components
to attain normal myelination and show that plasmalogen deficiency impairs the organization
of myelin and myelinating Schwann cells.
Defects in plasmalogens impair regeneration and preservation of axons and
myelin. To further investigate the role of plasmalogens in Schwann cells and myelin, we performed
sciatic nerve crush in adult mice. Histological and morphometric analyses performed 15
days after crush in the distal segment of crushed nerves from WT and
Pex7-KO mice revealed reduced density of remyelinated axons in mutant
nerves (Figure 3, A and B). In addition, the extent
of remyelination was reduced, showing a higher g ratio in mutant nerves (Figure 3C). Our ultrastructural analysis highlighted that the
defect in remyelination was not due to impaired axonal regeneration, as crushed nerves
from Pex7-KO mice had a 3-fold increase in the density of demyelinated
fibers (Figures 3, D and E), i.e., large-caliber
axons surrounded by a Schwann cell but lacking myelin sheaths (Figure 3D, asterisks). These results indicate that plasmalogen deficiency
primarily affects the ability of Schwann cells to remyelinate regenerating axons.
Plasmalogens are important players during remyelination of the PNS. (A) PPD-stained cross sections of the distal segment of sciatic nerves 15
days after nerve crush. Scale bars: 10 μm. (B) Degree of regeneration
as measured by the density of myelinated axons in the distal segment 15 days after
sciatic nerve crush. *P = 0.014. (C) Extent of impaired
regeneration as measured by g ratio determination. Results are graphed as boxes with a
line at the mean and whiskers from the minimal to maximal values. *P =
0.029. (D) Electron microscopic analysis of the distal segment of crushed
sciatic nerves from WT and Pex7-KO mice. In WT nerves, remyelination of
regenerating axons was evident (arrowheads), whereas Pex7-KO axons of a
similar caliber were devoid of myelin (asterisks). Scale bars: 5 μm.
(E) Density of axons lacking myelin (demyelinated axons) in the distal
segment following sciatic nerve crush. Error bars represent SEM. *P =
Analysis of sciatic nerves from aged Pex7- and Gnpat-KO
mice (mean age 17 ± 3.3 months) revealed axonal loss and demyelination (Figure 4A), with decreased numbers of myelinated axons (Figure
4B) and myelin loss in medium- and large-caliber
axons (Figure 4C). Ultrastructural analysis showed
axons enwrapped by Schwann cells but lacking myelin (Figure 4D, asterisk) and production of redundant basal lamina by mutant Schwann cells
(Figure 4D, arrowheads). The endoneurium of
plasmalogen-deficient mice was filled with Schwann cell processes, and normal-appearing
myelinating Schwann cells also extended processes in an apparent attempt to engulf and
enwrap demyelinated and normally myelinated axons (Supplemental Figure 3). In aged
Pex7- and Gnpat-KO mice, attempts at remyelination by
mutant Schwann cells were evident by the identification of myelinating images (Figure
4E) that are characteristic of the initial stages
of myelination in newborn mice. However, the process of remyelination seemed to be
ineffective and abortive, with the formation of onion bulb–like structures (Figure
4F) and axonal loss. Together, our results reveal
that during normal development and following nerve regeneration, plasmalogens are crucial
for Schwann cells to sort axons, myelinate, and to sustain axons and myelin levels in the
Deficiency in plasmalogens impairs the ability of Schwann cells to sustain
myelinated axons. (A) PPD-stained cross sections of sciatic nerves from 1.5-year-old WT and
Pex7-KO mice revealing demyelination and loss of axons in mutant
nerves. Scale bars: 10 μm. (B) Density of myelinated axons in sciatic
nerves from WT and Pex7-KO mice. Error bars represent SEM.
*P = 0.012. (C) Quantification of the degree of
myelination by g ratio in sciatic nerves from 1.5-year-old WT and
Pex7-KO mice. *P = 0.026. Error bars represent SEM.
(D) Electron microscopic analysis of sciatic nerves from representative
1.5-year-old WT and Pex7-KO mice showing an axon devoid of myelin
(asterisk), severely demyelinated axons (arrows), and the presence of extended Schwann
cell processes (arrowheads) throughout the perineurium. Scale bars: 2 μm.
(E) Schwann cell from an aged Gnpat-KO mouse undergoing
remyelination of a demyelinated axon. Scale bar: 2 μm. (F) Engulfment
of a demyelinated axon by 2 Schwann cells, generating concentric deposition of membrane
processes, similar to the formation of onion bulbs. Scale bar: 2 μm.
Plasmalogens regulate AKT activation at the plasma membrane. To elucidate the mechanisms underlying the effects of plasmalogen deficiency in Schwann
cells, we analyzed key kinases in sciatic nerves from P15 WT and Gnpat-KO
mice. Levels of phosphorylated c-Jun N-terminal kinase (JNK), signal transducer and
activator of transcription 3 (STAT3), extracellular signal–regulated kinase 1/2
(ERK1/2), and downstream targets of the GTP-binding proteins of the Rho family (namely,
N-WASP, myosin light chain, and cofilin) were normal in Gnpat-KO nerves
(data not shown). However, we observed a 3-fold decrease in AKT phosphorylation (Figure
5A) and in the phosphorylation of two of its
targets, GSK3β (Figure 5B) and c-RAF (Figure
5C). The defective phosphorylation of AKT was
unrelated to altered levels of phosphorylated PDK1 (Figure 5D) or PTEN (Figure 5E), two upstream
regulators of AKT activation. The levels of phosphorylated AKT and GSK3β at Ser9
were reduced in nerve lysates from Gnpat-KO mice at P4 (Supplemental
Figure 4, A and B), but were normal in lysates from Gnpat-KO mice at 3
months of age (Supplemental Figure 4C). During the active period of axonal sorting, we
also observed a decrease in the proliferation rate of Schwann cells. Pulse-chase in vivo
experiments with BrdU showed reduced BrdU incorporation in sciatic nerves from
Gnpat-KO mice at P4 (Figure 5F).
Defects in plasmalogens result in impaired AKT activation and signaling. (A) Western blot analysis and quantification of AKT phosphorylation
(p-AKT) in sciatic nerve lysates of P15 WT and Gnpat-KO mice.
*P = 0.018; **P = 0.006.
(B–E) Quantification of phosphorylated forms of
GSK3β at Ser9 (B), c-RAF at Ser259 (C), PDK1 at Ser241
(D), and PTEN at Ser380 (E) in sciatic nerves from WT and
Gnpat-KO mice. *P < 0.02. (F)
Density of BrdU-positive cells in nerves from P4 WT and Gnpat-KO mice.
*P = 0.020. (G) Western blot analyses of p-AKT and
p-ERK1/2 in serum-starved MEFs from WT and Gnpat-KO mice stimulated
with 10% FBS. (H and I) Quantification of p-AKT at Ser473
(H) and Thr308 (I) in primary WT and
Gnpat-KO Schwann cells after stimulation with NRG1. (J)
Western blot analysis of total and p-AKT in cytosolic and membrane fractions of
serum-starved MEFs from WT and Gnpat-KO mice stimulated with 10% FBS.
Western blot analysis of caveolin 1 (CAV1), GAPDH, and peroxisomal thiolase (ACAA1) used
to control membrane fractions and cytosolic fractions and to monitor lack of solubilized
organelles in cytosolic fractions, respectively. (K) DRG cocultures from WT
and Gnpat-KO mice treated with DMSO (control) or with SC79, stained for
neuronal βIII-tubulin (green) and MBP (red). Scale bars: 100 μm.
(L) Density of myelin segments in DRG cocultures from WT and
Gnpat-KO mice after DMSO and SC79 treatment. *P <
0.002. (M) Length of individual myelin segments in myelinating cocultures.
*P < 0.01. Error bars represent SEM in all graphs.
Phosphorylation of AKT was also impaired in serum-starved mouse embryonic fibroblasts
(MEFs) from Gnpat-KO mice after stimulation with 10% FBS (Figure 5G and Supplemental Figure 4D). However, FBS stimulation
was able to normally activate ERK1/2 in serum-starved MEFs from KO mice (Figure 5G and Supplemental Figure 4E). We next investigated
whether the NRG1/ErbB signaling pathway would also be affected by the plasmalogen
deficiency. We measured the levels of AKT phosphorylation in primary Schwann cells from WT
and Gnpat-KO mice after stimulation with NRG1, the endogenous axonal
signal that regulates Schwann cell differentiation and myelination (18). We detected decreased levels of phosphorylation, indicative of
impaired AKT activation in Schwann cells from Gnpat-KO mice (Figure 5, H and I). The recruitment of AKT to the plasma
membrane plays a critical role in its activation by PDK1 and by the mammalian TOR complex
2 (25, 26).
Compared with WT MEFs, the plasmalogen deficiency in Gnpat-KO MEFs
impaired the FBS-induced phosphorylation of AKT at the membrane (Figure 5J and Supplemental Figure 4F), despite normal levels and
localization of PDK1 (Supplemental Figure 4G). Failure to activate AKT at the membrane was
also evident by the decreased levels of phosphorylated AKT in the cytosolic fraction
(Figure 5J). To determine whether the defect in AKT
activation plays a critical role in the impaired myelination observed in DRG cocultures,
we performed in vitro myelination assays in the presence of SC79. This small compound was
shown to allow AKT activation in the cytosol, bypassing the need for AKT to be targeted to
the membrane (27). Treatment of DRG cocultures from
Gnpat-KO mice with SC79 (Figure 5K) increased the number and length of myelin segments (Figure 5, L and M), indicating the rescue in myelination. These
results highlight that plasmalogens are important for the correct recruitment and
activation of AKT at the membrane and that their absence causes a signaling defect
independently of ligand-receptor activation.
Defective differentiation of Schwann cells in plasmalogen-deficient mice is rescued
by treatment with GSK3β inhibitors. AKT promotes Schwann cell differentiation and myelination in part through the inhibitory
phosphorylation of GSK3β at Ser9 (28). We
therefore tested whether treatment with LiCl, a GSK3β inhibitor (29), would rescue the Schwann cell defects observed in
Gnpat-KO mice. The addition of LiCl to the culture medium rescued the
myelination defect of DRG mixed cultures from Gnpat-KO mice (Figure 6A). Therefore, we treated WT and
Gnpat-KO mice with LiCl or NaCl in two treatment schemes (Figure 6B). Following LiCl treatment for 1 week from P7,
analysis of nerves from Gnpat-KO mice revealed decreased phosphorylation
of GSK3β at Tyr216 (Figure 6C) and increased
phosphorylation of GSK3β at Ser9 (Figure 6D).
We found that higher levels of GSK3β phosphorylation at Ser9 were due to increased
phosphorylation of AKT (Figure 6, E and F), since
lithium dissociates the protein phosphatase 2A–containing (PP2A-containing)
complex, decreasing the dephosphorylation of AKT (30).
Treatment with GSK3β inhibitors restores Schwann cell differentiation and
radial sorting defects. (A) In vitro myelination after treatment with NaCl (control) or LiCl.
*P = 0.001. (B) Strategy for in vivo treatments. For the
assessment of LiCl on the phosphorylation of AKT and GSK3β, mice were injected
with LiCl on alternating days from P7 to P15 (upper diagram). For the assessment of LiCl
on nerve pathology, mice were injected daily with LiCl from P1 to P6 (lower diagram).
(C and D) Quantification of p-GSK3β at Tyr216
(*P = 0.04) (C) and Ser9 (*P = 0.017)
(D) in nerves from P15 mice. (E and F)
Quantification of p-AKT at Ser473 (E) and Thr308 (F) in nerves
from P15 mice. *P = 0.01. (G and H)
Quantification of SOX2 (G) and OCT6 (H) levels in nerves from
P6 WT and Gnpat-KO mice. *P = 0.03 (G);
P = 0.038 (H). (I) Ultrastructural analysis
of sciatic nerves from P6 Gnpat-KO mice treated with NaCl and LiCl.
Lines and asterisks indicate axon bundles on control and lithium-treated nerves,
respectively. Scale bars: 5 μm. (J) Number of axons in bundles of
sciatic nerves from P6 mice treated with NaCl and LiCl. *P = 0.0013.
(K) Density of sorted axons (promyelinating stage) in sciatic nerves from
P6 mice treated with NaCl and LiCl. *P = 0.0022. (L)
Number of axons in bundles of sciatic nerves from P4 mice treated with DMSO and TDZD-8.
*P < 0.007. (M) Density of sorted axons in sciatic nerves from P4
WT and Gnpat-KO mice treated with DMSO and TDZD-8. *P
Lithium treatment during the first 6 days of life induced Schwann cell differentiation in
Gnpat-KO nerves, as levels of SOX2, a marker of immature Schwann cells,
were decreased (Figure 6G), and levels of OCT6, a
marker of differentiated Schwann cells, were increased (Figure 6H). Ultrastructural analysis of sciatic nerves from LiCl-treated
Gnpat-KO nerves showed improved myelination (Figure 6I) and rescue in axonal sorting, with normal numbers of
axons in bundles (Figure 6J) and normal density of
sorted promyelinating Schwann cells containing a single axon (Figure 6K). Similar results were obtained when newborn WT and
Gnpat-KO mice were treated for 4 days with TDZD-8, a highly specific
inhibitor of GSK3β (31). In sciatic nerves
from P4 Gnpat-KO mice treated with TDZD-8, we observed a decrease in the
number of axons per bundle (Figure 6L) and an
increase in the number of sorted axons (Figure 6M).
Taken together, our results indicate that plasmalogen deficiency affects Schwann cell
differentiation and function, causing defects in axonal sorting, myelination, assembly of
myelin, and nerve conduction through the impairment of AKT activation at the plasma
membrane and via GSK3β.
In this study, we report that a plasmalogen defect in the PNS primarily affects Schwann
cells and that plasmalogens are crucial for Schwann cells at two developmental time points.
During the early postnatal period, plasmalogens are important for axon-glia interaction and
the myelination process. The recognition of axons by Schwann cells or the ability to
actively segregate axons destined for myelination is impaired by the deficiency in
plasmalogens. The presence of unsorted axons caused abnormal myelination of the Remak
bundles and reorganization of these structures. Additionally, impaired myelination,
increased regions of noncompact myelin, and defective compartmentalization of Schwann cells
in mutant nerves highlight the role of plasmalogens in the correct assembly of myelin and
the organization of Schwann cells. Surprisingly, the extent of myelin thickness was normal
in adult mutant mice, which led us to hypothesize, that like a deficiency in MBP, which has
minimal effects in the PNS despite the complete lack of myelin in the CNS, other myelin
components may contribute to achieving normal myelination (24, 32). MBP, known to modulate the
apposition of the cytoplasmic leaflets of myelin membranes through electrostatic
interactions, is also involved in the partition of myelin into compact and noncompact myelin
(33). To further investigate the role of
plasmalogens during myelination, we asked whether depleting myelin of plasmalogens and MBP,
two major constituents of myelin at the cytoplasmic apposition, would impair myelination. In
DM mice, we found a major impairment in myelination and nerve conduction, indicating that
plasmalogens are important regulators of myelination. In addition, we found that upon
Wallerian degeneration, injury-induced remyelination was also affected by the deficiency in
plasmalogens, although axonal regeneration seemed to be unaffected. Failure to maintain
myelin levels was also evident in sciatic nerves from aged mutant mice, in which we observed
extensive demyelination, axonal loss, and failed attempts to remyelinate the PNS.
Our results highlight that in mice, a deficiency in plasmalogens is sufficient to impair
PNS myelination and to cause age-dependent peripheral neuropathy. As such, the evaluation of
the PNS in RCDP type 2 and type 3 patients should be informative. In Refsum’s
disease, the single defect in α–oxidation of phytanic acid is known to cause
peripheral neuropathy (34). Previously, we identified
a group of atypical RCDP type 1 patients with mild mutations in PEX7 and a Refsum-like
presentation, which included retinitis pigmentosa and nerve hypertrophy (17). In addition to the accumulation of phytanic acid,
these patients also had a defect in plasmalogen synthesis. Based on our current findings, we
propose that RCDP type 1 patients should also be investigated for defects in nerve
conduction and pathology. In these patients, the peripheral neuropathy can be aggravated by
the combined defect in plasmalogen synthesis and phytanic acid accumulation.
In the present study, we analyzed the phosphorylation levels of several kinases involved in
PNS development to understand how a defect in plasmalogens impairs Schwann cell
differentiation and Schwann cell–mediated myelination. The results obtained in nerves
of Gnpat-KO mice during the active period of myelination in the PNS
revealed an impairment of AKT activation. Our observations of impaired activation of the AKT
pathway following induction with FBS or NRG1, which was not mediated by defects in upstream
regulators, suggested a more direct role of plasmalogens in AKT activation rather than of
multiple defects in several ligand-receptor interactions and activations. AKT activation by
phosphorylation at the plasma membrane was impaired in plasmalogen-deficient cells. In
mutant cells, AKT was found in membrane fractions, but upon FBS stimulation, AKT was not
phosphorylated despite the correct localization and phosphorylation of PDK1. The requirement
of AKT to associate with raft microdomains for efficient activation and signal transduction,
combined with the enrichment of plasmalogens in lipid rafts, suggests that the defect in
plasmalogens affects the correct compartmentalization of AKT, and thus its activation by
phosphorylation (25, 34). The AKT signaling pathway is important for lens fiber differentiation,
chondrocyte differentiation and maturation, adipogenesis, and spermatogenesis (35–38).
Based on our results showing that a plasmalogen defect impairs AKT activation in Schwann
cells, we hypothesize that in tissues affected by a plasmalogen defect (e.g., lens,
cartilage, adipocytes, and testis), impaired AKT signaling modulates the observed pathology,
namely, cataracts, impaired ossification, lipodystrophy, and loss of spermatocytes (39).
In sciatic nerves from Gnpat-KO mice, we found that impaired AKT
activation resulted in dysregulated phosphorylation of GSK3β. GSK3β activity and
its role in the regulation of transcription are known to modulate Schwann cell
differentiation and myelination (28, 40). Active GSK3β, phosphorylated at Tyr216, is
inhibited by AKT phosphorylation at Ser9 (41). In
nerves from Gnpat-KO mice, we observed increased levels of Tyr216
phosphorylation and decreased levels of Ser9 phosphorylation, which, combined, indicate
overtly active GSK3β (41). Lithium, a known
inhibitor of GSK3β, can directly inhibit the kinase through Mg2+
competition and can indirectly inhibit GSK3β by promoting further inhibition through
Ser9 phosphorylation (42, 43). Lithium administration to Gnpat-KO mice was able to
normalize GSK3β phosphorylation status, induced the stimulation of Schwann cell
differentiation, rescued the radial sorting defect, and improved myelination independently
of the plasmalogen defect. Similar findings were obtained when Gnpat-KO
mice were treated with TDZD-8, a highly specific inhibitor of GSK3β. Treatment of
plasmalogen-deficient mice with alkyl-glycerol rescues the biochemical defect and pathology
in several tissues, but has minimal effects on restoring plasmalogen levels in nervous
tissue (39). As such, improving the defective
AKT/GSK3β signaling pathway may have clinical potential in combined therapeutic
interventions with alkyl-glycerol during the active period of PNS myelination and in mildly
affected patients displaying signs of peripheral neuropathy.
In summary, our data reveal the role of plasmalogens in the neuropathic course of two
models of RCDP (see proposed model in Figure 7),
showing that plasmalogen deficiency severely impairs the ability of Schwann cells to
differentiate and mature, thus causing defects in myelination and nerve conduction. Through
the identification of the mechanism by which a defect in plasmalogens mediates the
pathology, we unraveled that ether phospholipids are crucial for the activation of the AKT
pathways and identified a candidate therapeutic intervention that overcomes the plasmalogen
deficiency and rescues the signaling pathway and Schwann cell differentiation.
Schematic representation of a proposed model depicting the consequences of
plasmalogen deficiency in the PNS. The deficiency in plasmalogens at the plasma membrane of Schwann cells causes reduced
phosphorylation of AKT, leading to an overt activation of GSK3β by reducing the
inhibitory phosphorylation at Ser9. Active GSK3β inhibits Schwann cell
differentiation, impairing axonal sorting and myelination. Lithium administration to
inhibit GSK3β is able to rescue Schwann cell differentiation and maturation in the
absence of plasmalogens. During aging, lack of plasmalogens causes demyelination and
axonal loss, and Schwann cells extend processes in failed attempts to remyelinate
existing axons. Plasmalogens are depicted by gray-colored phospholipids; ҂,
impaired signaling; →, induction; ┬, inhibition.
Mouse strains, procedures, and treatments. WT, Pex7, and Gnpat mice have been previously described
(9, 10). KO
and WT littermates of both sexes were obtained from mating of heterozygous mice. Shiverer
mice (shi; Mbpshi/J) were obtained from The Jackson
Laboratory. Pex7:shi and Gnpat:shi DM mice were obtained
from F2 mice after crossing heterozygous Pex7 or Gnpat
mice with homozygous shi mice.
For nerve crush, 2-month-old WT (n = 4) and Pex7-KO
(n = 6) mice were anesthetized (i.p. 100 mg/kg ketamine and 1 mg/kg
medetomidine), and the exposed right sciatic nerve was injured (15-second crush 2 times)
at the level of the notch with a fine hemostat. The contralateral nerve was used as a
control. Postsurgical analgesia (1 mg/kg butorphanol s.c.) was performed twice a day for 3
days. Nerve conduction (WT, n = 6; shi,
n = 4; Pex7-KO, n = 3;
Pex7:shi DM, n = 4; Gnpat-KO,
n = 5; Gnpat:shi DM, n = 3) was
determined as previously described (44).
For the BrdU incorporation assay, BrdU in 0.9% NaCl and 7 mM NaOH were injected i.p. at a
dosage of 50 mg/kg in WT and Gnpat-KO mice (n = 4 for
each genotype) 4 and 20 hours before the collection of sciatic nerves.
LiCl or NaCl at a dosage of 50 mg/kg was injected s.c. daily from P0.5 to P6 (control WT,
n = 6; Gnpat-KO, n = 6; lithium WT,
n = 7; Gnpat-KO, n = 6) or on
alternating days from P7 to P15 (control WT, n = 7;
Gnpat-KO, n = 7; lithium WT, n = 7;
Gnpat-KO, n = 7). TDZD-8 (Sigma-Aldrich) at 5 mg/kg or
DMSO at 20% (v/v) was injected s.c. daily from P0.5 to P4 (control WT, n=
4; Gnpat-KO, n = 4; TDZD-8 WT, n = 5;
Gnpat-KO, n = 5). Mice were used under standard
conditions and had free access to food and water. Experiments and mouse manipulations were
performed in compliance with the institutional guidelines and recommendations of the
Federation for Laboratory Animal Science Association (FELASA) and were approved by the
National Authority for Animal Health (DGAV; Lisbon, Portugal).
Histological and morphological analyses. Sciatic nerves were dissected and fixed by immersion on 4% glutaraldehyde in 0.1 M
cacodylate buffer (pH 7.4) for 1 week. After postfixation with 1% OsO4 in 0.1 M
cacodylate buffer (pH 7.4) for 2 hours, nerves were dehydrated and embedded in Epon
(Electron Microscopy Sciences). Sections (1-μm-thick) were stained for 10 minutes
with 1% p-phenylenediamine (PPD) in absolute methanol, dried, and mounted on a drop of DPX
(Merk). The entire area of the nerve was photographed on an Olympus optical microscope
equipped with an Olympus DP 25 camera and Cell B software, and images were imported into
Photoshop (Adobe). For g ratio analyses, the Photoshop recording measurement tool was used
to determine the area of the axon and the entire myelinated fiber (axon plus myelin).
Derived diameters were used to calculate the g ratio in 200 to 300 individual fibers per
genotype and age (P5 mice: WT, n = 7; Pex7-KO,
n = 4; Gnpat-KO, n = 4; P15 mice: WT,
n = 5; Pex7-KO, n = 4;
Gnpat-KO, n = 4; P30 and P90, n = 4
for all genotypes; 1.5-year-old: WT, n = 6; Pex7-KO,
n = 6; Gnpat-KO, n = 5). For the
determination of the density of myelinated fibers (MFs), the total number of MFs was
divided by the area of the entire cross section of the sciatic nerve.
Ultrathin sections (60-nm) prepared on a Leica ultramicrotome were placed on 200-mesh
copper grids (Electron Microscopy Sciences) and were counterstained, first with alcoholic
uranyl acetate solution (2% w/v; 10 minutes), then by aqueous uranyl acetate solution (2%
w/v; 10 minutes) and lead citrate (4% w/v; 10 minutes). Grids were observed on a JEOL
JEM-1400 transmission electron microscope equipped with an Orious Sc1000 digital camera.
Ten to 15 nonoverlapping images were obtained and used for all determinations.
Cell culture and in vitro myelination. MEFs were prepared from E12 embryos and cultured in high-glucose DMEM with 10% FBS and
antibiotics. Primary Schwann cells were isolated from sciatic nerves of 4-day-old WT and
Gnpat-KO mice and cultured in high-glucose DMEM with 10% normal horse
serum (NHS) and antibiotics. Schwann cells were serum starved for 12 hours before
stimulation with 30 ng/ml recombinant human neuregulin 1 β EGF-like domain (R&D
Systems). The lysates were prepared after 0, 7, and 15 minutes at 37°C. Serum-starved
MEFs were stimulated with 10% FBS for 0, 1, 3, and 5 minutes. The lysates were prepared in
PBS containing 0.3% Triton X-100, protease inhibitors (cOmplete, Mini; Roche), and 2 mM
In vitro myelination was performed as described (45). Mixed cultures of DRG neurons and Schwann cells from WT (n
= 7) and Gnpat-KO (n = 4) embryos were maintained in
high-glucose MEM with 2 mM L-glutamine, 10% FBS, 50 ng/ml nerve growth factor (NGF) (2.5S;
Millipore), and 1% penicillin-streptomycin, and were then plated onto 13-mm-diameter
Matrigel-coated coverslides (1:10 in DMEM). After 24 hours at 37°C, cells were
cultured for 10 days in neurobasal medium supplemented with 4 g/l glucose, 2 mM
L-glutamine, 50 ng/ml NGF, and 1× B27 (Gibco). Myelination was induced at 10 days in
vitro (div) by the daily addition of 50 μg/ml ascorbic acid to high-glucose MEM
medium supplemented with 2 mM L-glutamine, 10% FBS, and 50 ng/ml NGF. For lithium
treatment in myelination assays, medium was supplemented with 3 μM forskolin and 16
mM NaCl (control) or 16 mM LiCl starting at div 7. For treatment with SC79, medium was
supplemented with 0.0008% v/v DMSO or with 0.4 μg/ml SC79 (Millipore) in DMSO
starting at div 7. In both treatment schemes, cells were fixed and processed for
immunofluorescence at 20 div.
Antibodies. The antibodies used were: anti–βIII-tubulin (1:600; 1967-1; Epitomics);
anti-MBP (1:250; MAB386; Millipore); anti-DRP2 (1:1,000; ref. 46); anti–Ser473-AKT; anti–Thr308-AKT;
anti–pan-AKT; anti–Ser259-c-RAF; anti–Ser9-GSK3β (all at
1:1,000; catalog 9916; Cell Signaling Technology); anti–β-tubulin (1:1,000;
catalog 5142; Cell Signaling Technology); anti–caveolin 1 (1:5,000; C13630; BD
Transduction Laboratories); anti-ACAA1 (1:1,000; HPA007244; Sigma-Aldrich); anti-GAPDH
(1:1,000; catalog 5174; Cell Signaling Technology); and anti-BrdU (1:2,000; Ab6326;
Abcam). For immunofluorescence, Alexa Fluor 488– and Alexa Fluor
568–conjugated secondary antibodies were used (1:500; Invitrogen). For detection of
BrdU, nerves from P4 WT and Gnpat-KO mice were fixed in carnoys and
processed for immunofluorescence in paraffin sections pretreated with 2N HCl. For Western
blotting, horseradish peroxidase–conjugated secondary antibodies were used
(1:5,000; Jackson ImmunoResearch).
Immunofluorescence. In the in vitro myelination assays, fixed and permeabilized cells were incubated for 1
hour at room temperature (RT) with blocking buffer (5% normal donkey serum [NDS] in PBS).
Primary antibodies (rabbit anti–βIII-tubulin and rat anti-MBP) were diluted
in blocking buffer and incubated for 16 hours at 4°C. After washing with PBS,
secondary antibodies were diluted in blocking buffer and incubated for 1 hour at RT. After
washing, coverslides were mounted on microscope slides with Vectashield containing DAPI
(Vector Labs). Slides were analyzed by epifluorescence on a Zeiss Axio Imager Z1
microscope equipped with an Axiocam MR3.0 camera and Axiovision 4.7 software.
For teased fiber preparations, sciatic nerves from 9-month-old WT and
Gnpat-KO mice (n = 3 for each genotype) were dissected
and fixed by immersion with 4% PFA in PBS for 30 minutes. Under a dissection microscope,
the perineurium was removed, and small nerve bundles were separated using 30-gauge
needles. Nerve bundles were blocked and permeabilized by incubation for 1 hour at RT in
blocking buffer containing 0.1% Triton X-100. Bundles were incubated with
rhodamine-conjugated phalloidin (1:400; Invitrogen) or with rabbit anti-DRP2 (46) for 16 hours at 4°C. After washing with PBS,
bundles were incubated for 2 hours at RT with secondary antibodies. Nerve bundles were
then washed in PBS and placed in a drop of Vectashield containing DAPI. Single teased
fibers were prepared using 30-gauge needles, sealed, and viewed by confocal microscopy on
a Leica TCS SP5 II.
Biochemical analyses. For Western blot analysis, lysates of sciatic nerves were prepared by sonication in lysis
buffer, and 25 μg protein was run on 12% SDS-PAGE gels. Blots were developed with
ECL (Pierce) and exposed to Hyperfilm (Amersham). Films were scanned on a Molecular Imager
GS800, and Quantity One (Bio-Rad) was used for quantifications. The ProteoExtract
subcellular proteome extraction kit (Calbiochem) was used to obtain membrane and cytosolic
fractions of cultured MEFs from WT and Gnpat-KO mice. Three independent
experiments were performed.
X-ray diffraction. Sciatic nerves from WT (n = 6) and Pex7-KO
(n = 7) mice were fixed by immersion on 2% glutaraldehyde in 0.12 M
phosphate buffer (pH 7.4). Diffraction experiments and data analysis were carried out as
detailed previously (47). For the determination of
the relative amount of myelin by x-ray diffraction, the quotient M/M+B was used, in which
the denominator includes the total x-ray scatter coming from the volume of nerve subtended
by the x-ray beam (consisting of the multilamellar myelin, M, and background, B), and the
numerator is the total intensity coming from the multilamellar myelin (the peak
intensities above background), as previously described (47).
Statistics. Results were expressed as the mean ± SEM, and data were analyzed with Prism software
(GraphPad Software). To compare 2 groups, nonparametric Mann-Whitney U
tests were used, and P < 0.05 was considered a significant difference.
For comparison of more than 2 groups, 1-way ANOVA tests were used, followed by
Tukey’s multiple comparison tests.
View Supplemental data
We thank Paula Sampaio for microscopy support, Paula Magalhães for genotyping, and
Isabel Carvalho, Sofia Lamas, and Fátima Martins for excellent animal care. We are
grateful to P. Brophy (University of Edinburgh) for the DRP2 antibody and to M. Baes (K.U.
Leuven) for providing the Gnpat mouse strain. This work was funded by the
Research Foundation of the European Leukodystrophy Association (ELA 2008-009C4, ELA
2010-042C5), by FEDER Funds through the Operational Competitiveness Program –
COMPETE, and by national funds through the FCT – Fundação para a
Ciência e a Tecnologia under the project FCOMP-01-0124-FEDER-015970
(PTDS/SAU-ORG/112406/2009). P. Brites is an FCT Investigator, and T. Ferreira da Silva was
supported by the FCT (SFRH/BD/88160/2012).
Conflict of interest: The authors have declared that no conflict of interest
Citation for this article:J Clin Invest.
Adrienne Luoma’s present address is: Committee on Immunology, University of
Chicago, Chicago, Illinois, USA.
Robin L. Avila’s present address is: Department of Neurology, The University of
Chicago, Chicago, Illinois, USA.
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