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The membrane-bound receptor for platelet-derived growth factor A (PDGFRα) is crucial for controlling the production of oligodendrocytes (OLs) for myelination, but regulation of its activity during OL differentiation is largely unknown. We have examined the effect of increased sulfated content of galactosylceramides (sulfatides) on the regulation of PDGFRα in multipotential neural precursors (NPs) that are deficient in arylsulfatase A (ASA) activity. This enzyme is responsible for the lysosomal hydrolysis of sulfatides. We show that sulfatide accumulation significantly impacts the formation of OLs via deregulation of PDGFRα function. PDGFRα is less associated with detergent-resistant membranes in ASA-deficient cells and showed a significant decrease in AKT phosphorylation. Rescue experiments with ASA showed a normalization of the ratio of long versus short sulfatides, restored PDGFRα levels, corrected its localization to detergent-resistant membranes, increased AKT phosphorylation, and normalized the production of OLs in ASA-deficient NPs. Moreover, our studies identified a novel mechanism that regulates the secretion of PDGFRα in NPs, in glial cells, and in the brain cortex via exosomal shedding. Our study provides a first step in understanding the role of sulfatides in regulating PDGFRα levels in OLs and its impact in myelination.
Oligodendrocyte progenitor cells (OPCs)2 are primarily generated from undifferentiated multipotential neural precursors (NPs) during early embryogenesis under the influence of morphogenic molecules, such as Shh (sonic hedgehog) (1,–4). Multiple mechanisms are developmentally coordinated to regulate the differentiation of OPCs into functional oligodendrocytes (OLs) (5,–7). Of these, signaling through the platelet-derived growth factor receptor α (PDGFRα) is essential in controlling the proliferation and survival of OPCs (8,–10). PDGFRα is expressed in NPs (11) and marks the appearance of the first PDGF-responsive OPCs during early neurogliogenesis (12). Numerous studies have highlighted the relevance of PDGFRα signaling in oligodendrogenesis. For example, PDGFRα+ cells isolated from the embryonic spinal cord exclusively generate myelinating oligodendrocytes (12, 13). Furthermore, the formation of mature OLs is significantly reduced upon ablation of PDGFRα+ cells (12) and in mice with targeted disruption of its ligand, PDGF-AA (14). Functionally, the activity of PDGFRα depends in part on its localization to detergent-resistant membrane (DRM) domains in the plasma membrane and interaction with integrin complexes (15,–17). The association of PDGFRα with DRMs underlines the possibility that conditions affecting DRMs may impact PDGFRα function and, therefore, the myelination program.
Previously, we found that exogenous sulfated galactosylceramides (sulfatides) significantly reduced the formation of OLs from NPs in vitro (18). One interpretation from these results is that sulfatides might repress the sensitivity of OPCs to proliferating signals such as PDGF-AA, thereby reducing the pool of progenitors available for differentiation into mature OLs. Sulfatides are crucial sphingolipids in myelin architecture (19) and have been found to negatively regulate the maturation of OPCs into differentiated OLs (20, 21), but the mechanism is still unknown. Sphingolipids, including sulfatides, have structural and functional roles in DRMs (22, 23) and participate in caveolar and exosomal biogenesis (22, 24,–26). Therefore, sulfatides have an intrinsic potential to modulate the activity of membrane-bound receptors such as PDGFRα by altering membrane domains such as DRMs.
In this study, we tested the hypothesis that sulfatides contribute to the regulation of oligodendrogenesis by modulating PDGFRα function. We found that increased sulfatide levels in NPs lead to a reduced production of OPCs and OLs. Furthermore, we observed a diminished association of PDGFRα with DRMs, repressed AKT phosphorylation, and exacerbated secretion of PDGFRα via exosomes. We present evidence that exosomal secretion of PDGFRα is a natural process in glial cells in vitro and during myelination of the murine cortex, when sulfatides are highly produced.
Heterozygous ASA+/− breeders (obtained from Dr. Gieselmann and back-crossed in the C56BL/6 background) were maintained in standard housing conditions, under the approval of the Animal Care and Use Committee. ASA+/+ and ASA−/− embryos at 16.5 days of gestation and 3-day-old newborns were used in our experiments. ASA+/+ and ASA−/− mice at 7, 14, and 21 days were killed without sex distinctions for immunocytochemical and in vivo exosome isolation studies.
NPs were isolated from ASA+/+ and ASA−/− embryonic day 16.5 telencephalon by mechanical dissociation and maintained as proliferating spheres in the presence of 10 ng/ml FGF-2 and 20 ng/ml EGF (27). Cultures of NPs obtained from different litters were used between passages 3 and 10 with identical results (n = 5–6). For differentiation assays, NPs were mechanically dissociated and seeded at a density of 7.5 × 104 cells/cm2 onto coverslips precoated with Matrigel (BD Biosciences) for 1 h at room temperature. Cultures were maintained for 3 or 7 days (3 or 7 DIV) in the absence of growth factors and in the presence of 2% fetal bovine serum (differentiated medium). Differentiated medium containing 2% FBS showed traces of Alix and Rab5B (data not shown). In some experiments, differentiated cells were exposed to PDGF-AA (Peprotech) at a concentration of 20 ng/ml for 1 day after plating. For Western blot analyses, NP spheres were collected 5 days after proliferation or 7 days after plating for differentiation.
Analysis of proteolytic degradation of the PDGFRα were performed utilizing 2 × 106 dissociated ASA+/+ and ASA−/− NPs. NPs were exposed to 10 μm MG132 or 10 mm NH4CL in basal proliferating medium conditions. Cells incubated with MG132 or NH4Cl for 6 h were collected for protein expression analysis of the PDGFRα as described below (see “Immunoblotting”). Because MG132 was dissolved in DMSO, DMSO-treated ASA+/+ and ASA−/− NPs were included as controls. All experiments were repeated three times. Additionally, three independent experiments were performed exposing 4 × 106 dissociated ASA+/+ and ASA−/− NPs to PDGF-AA ligand at a concentration of 20 ng/ml for 30 min before cell collection. For these experiments, NPs were starved for 3 h of growth factors present in the proliferating medium (EGF and basic FGF) and exposed to MG132 or NH4Cl as described above. After starvation, NP metabolism was slowed down by an ice bath for 15 min, cells were pelleted, and medium was preserved on ice. Pelleted NPs were exposed to fresh medium containing PDGF-AA for 30 min on ice. Cells were pelleted and washed to remove unbound ligand and resuspended in their original medium with MG132 or NH4Cl for an additional 30 min at 37 °C. NPs were pelleted, washed with PBS, and collected for Western blot analysis. NP pellets were analyzed for PDGFRα and downstream signaling for AKT, phospho-AKT (Ser308 and Thr478), MAPK p42/44, and phospho-MAPK p42/44 (Thr202/Tyr204).
For analysis of the effects of exogenous sulfatides (Avanti), ASA+/+ NP spheres were dissociated and incubated with two pulses of 10 μm sulfatides at days 1 and 3 and collected after 5 days.
Primary glial cultures were prepared from ASA+/+ and ASA−/− as described previously (28). Cell suspensions were seeded in 75-cm2 tissue culture flasks coated with 10 μg/ml poly-l-lysine at equal cell densities. Medium was changed 4 days after plating. Glial conditioned medium was collected at 7, 14, and 21 DIV and stored at −20 °C.
CG4 cells were cultured as described previously (29), except that B104 conditioned medium was replaced with 10 ng/ml PDGFA (BD Biosciences) and 5 ng/ml basic FGF (Peprotech). After 5 days in culture, supernatants were processed for exosomal isolation and cell pellets for Western blot analysis as described below.
NP lysates (equivalent to 2 mg of protein) were equilibrated with a solution of 90% sucrose to give a final solution of 45% sucrose and subjected to ultracentrifugation using a sucrose gradient (24). Twelve 0.25-ml fractions were collected and analyzed by immunoblotting.
Cholesterol levels of ASA+/+ and ASA−/− NPs from each sucrose fraction (50 μl) were measured using the fluorometric Amplex Red cholesterol assay kit (Invitrogen). Fluorometric analysis was performed using the Beckman Coulter DTX 880 multimode detector.
Sulfatides were extracted from total lysates of NP spheres at 5 days (ASA+/+, ASA−/−, and ASA−/− NPs treated with two pulses of 5, 15, or 25% ASA conditioned medium (ASACM) at day 1 and 3 of dissociated spheres) and DRM fractions (ASA+/+ and ASA−/− NPs) as described previously with some modifications (30,–32). C12:0 sulfatide was used as an internal standard (0.5 μm). Fifty-microliter aliquots from each sample were used for protein quantification. Ten volumes of chloroform/methanol (CHCl3/CH3OH (2:1, v/v)) were added to each sample and incubated with agitation at room temperature for 2 h. Each sample was filtered, and 0.2 volumes of 0.9% NaCl was added. Mixtures were vortexed and centrifuged at 2000 rpm for 5 min. The lower phase was dried under N2 at 45 °C. Lipid extracts were dissolved in CHCl3/0.6 n NaOH, CH3OH (1:1, v/v) and incubated for 2 h at room temperature with agitation. Extracts were washed with water and centrifuged at 2000 rpm. The lower phase was retained, washed with CHCl3/CH3OH/H2O (3:48:47, v/v/v), and centrifuged at 2000 rpm. The lower phase was evaporated, and the dry product was dissolved in CHCl3 (500 ml/10 mg of product). Lipid extracts were fractionated on an SPE-NH2 column as described (33). Briefly, upon activation with hexane, fractional elution was carried out, and sulfatides were eluted with CHCl3/CH3OH/3.6 m aqueous NH4C2H3O2 (30:60:8, v/v/v) (as a last fraction), dried with N2 at 45 °C, and stored at −20 °C until use. Samples were dissolved in 60 ml of 5 mm NH4HCO2 in CH3OH, and 2 ml were injected onto the UPLC column. The four major sulfatide isoforms enriched in white matter (C16:0, C18:0, C24:0, and C24:1) were analyzed using UHPLC-MS/MS on a Shimadzu Nexera/LCMS-8040 triple quadrupole mass spectrometer (Kyoto, Japan). A Waters Acquity UPLC C18 column (2.1 × 50 mm, 1.7 mm) was used to separate sulfatides. The following gradient was used: 0–3 min, from 65 to 83% A; 3–3.5 min, from 83 to 95% A, held at 95% A for 1.5 min (solvent A, CH3CN; solvent B, 5 mm NH4HCO2 in water, pH 4.5). The flow rate was 0.6 ml/min, the autosampler temperature was 20 °C, and the column oven temperature was 45 °C. All analyte measurements were done using negative ion electrospray mass spectrometry with collision-induced dissociation and selected reaction monitoring. Selected reaction monitoring transitions monitored were as follows: C12:0, m/z 722 to 97; C16:0, m/z 778 to 97; C18:0, m/z 806 to 97; C24:1, m/z 888 to 97; C24:0, m/z 890 to 97.
Conditioned media from NPs (3 × 107 cells) and primary glial cultures (3 × 107) (n = 3 for each group) were filtered (0.22 μm) and subjected to ultracentrifugation at 100,000 × g for 90 min. Pellets were resuspended in cold PBS and ultracentrifuged at 100,000 × g for 90 min. Exosome pellets were dissolved in lysis buffer as described below, and equal amounts of proteins (4 μg) were separated by SDS-PAGE and transferred to nitrocellulose, followed by immunodetection of PDGFRα, EGFR, and the exosomal marker Rab-5b. Exosomes were isolated from ASA+/+ and ASA−/− brain cortices at 7, 14, and 21 days postnatal as described (34). The final exosomal pellet was subjected to a sucrose gradient centrifugation at 200,000 × g for 16 h at 4 °C. A total of seven fractions were collected (fraction a, top; fraction g, bottom). Fractions b, c, and d, enriched in the exosomal marker Rab-5b, were each resuspended in 40 μl of lysis buffer. Western blot analysis was performed on a half of the volume of each fraction. For transmission electron microscopy, exosomes were fixed in 2% paraformaldehyde and stained as described (35). Samples were imaged using a 120-kV transmission electron microscope, JEOL JEM-1220, and equipped with a Gatan Es1000W 11-megapixel CCD camera.
The deficiency in ASA activity in ASA−/− NPs was rescued by cross-correction using ASA-supplemented cell medium (36). Briefly, ASA-secreting HeLa cells were grown in 10% FCS DMEM/F-12 before conditioning. Cells were incubated with serum-free cell medium for 4 days. Conditioned media were filtered through a 0.45-μm filter and kept frozen until use. For correction experiments, 5, 15, or 25% ASACM or 25% control medium (DMEM/F-12 medium) was added to ASA−/− NP culture medium. ASA activity was measured utilizing 4-methylumberyl sulfate derivative substrate as described by Porter et al. (37).
RNA was purified using TRIzol (Invitrogen) and reverse-transcribed with Superscript III (Invitrogen). SYBR Green-based real-time PCR analysis was performed using primers specific for the PDGFRα sequence. The 60 S acidic ribosomal protein P0 (RPLP0) was used as an internal standard. Primers were optimized on a standard curve with an efficiency between 90 and 110% and a correlation coefficient higher than and 0.990. PCR analysis was calculated using the ΔΔCt method. The following primers were used: PDGFRα forward, 5′-ATATGATCTTTCTGTGGTTTAA-3′; reverse, 5′-CACTGCTTGGCAGAGCTACCT-3′; RPLP0 forward, 5′-CACGAAGCTAACGACTATCGC-3′; reverse, 5′-CTCTAGGGACTCGTTCGTGC-3′.
Samples were homogenized in lysis buffer (25 mm Tris-HCl, pH 7.4, 150 mm NaCl, 5 mm EDTA, 1% Triton, mammalian protease inhibitor mixture, 1 mm PMSF, 1 mm okadaic acid, and 2 mm sodium orthovanadate) on ice for 30 min with vortexing at 10-min intervals. After centrifugation at 10,000 × g, 10 μg of protein/sample was loaded onto 4–12% 1.5-mm precast NuPage gels (Invitrogen) and electrophoresed using an XCell Sure-lock vertical electrophoresis system (Invitrogen). After transferring to nitrocellulose membrane, blots were blocked with milk/BSA solution and incubated with the relevant antibodies. Immunoreactive products were detected using peroxidase-labeled secondary antibodies and ECL chemiluminescent substrate (Pierce). Immunoblots were semiquantified using ImageJ analysis software (National Institutes of Health).
For differentiation studies, cultures were maintained for 3 or 7 DIV in the absence of growth factors and in the presence of 2% fetal bovine serum. Cells were fixed with 4% paraformaldehyde for 20 min and washed twice with PBS. Cells were incubated with primary antibodies at room temperature for 1 h or overnight at 4 °C, washed extensively with PBS, and then incubated with Alexa-conjugated secondary antibodies (Molecular Probes). Coverslips or chamber slides were mounted with Prolong Gold antifade reagent with DAPI (Invitrogen) and imaged using either DM5500 Q Microscope with a Leica DFC500 camera or Zeiss Meta 510 confocal microscope. Cell counting was performed in the ImageJ software by counting cell-specific markers (NG2 for oligodendrocyte precursor cells, O4 for intermediate oligodendrocytes, glial fibrillary acidic protein (GFAP) for astrocytes, and DAPI for the nuclei). Three images of each coverslip or chamber slide were counted, and all experiments were repeated at least three times with different primary NPs cultures. The cell percentage was calculated for each specific marker with respect to total DAPI+ cells. Brains were collected after perfusion with PBS and 4% paraformaldehyde. Sections were cryoprotected in 20% sucrose, embedded in OCT, and sectioned at 20 μm on a cryostat. Tissue sections were immunostained for myelin basic protein (MBP) and PDGFRα and developed using the diaminobenzidine-ABC kit (Vector Laboratories).
The following antibodies were used in this study: actin (1:3000; Sigma), AKT (1:1000; Cell Signaling), Alix (1:500; Millipore), ASA (1:750; a gift from Dr. V. Gieselmann, Rheinische Friedrich-Wilhelms University, Bonn, Germany), caveolin 1 (1:1000; Cell signaling), EGFR (1:500; Upstate), p44/42 MAPK (1:2000; Cell Signaling), flotillin 2 (1:2000; BD Biosciences), GFAP (1:1000; Millipore), MBP (1:500; a gift from Dr. E. R. Bongarzone (University of Illinois, Chicago, IL)), HSP40 (1:1000; Cell Signaling), HSP60 (1:1000; Cell Signaling), phospho-AKT-Thr308 (1:1000; Cell Signaling), phospho-AKT-Ser473 (1:1000; Cell Signaling), phospho-p44/42 MAPK (1:1000; Cell Signaling), chondroitin sulfate proteoglycan, NG2 (1:300; Millipore), PDGFRα (1:200; Santa Cruz Biotechnology, Inc., and Cell Signaling), O4 (1:30; a gift from Dr. Bongarzone), proteolipid protein (PLP) (1:50; a gift from Dr. Bongarzone), and Rab-5b (1:200; Santa Cruz Biotechnology).
Data were analyzed by Student's t test or one-way analysis of variance, considering a p value of <0.05 as significant. Results are given as the means ± S.E. All analyses were performed using GraphPad Prism version 5.0d (GraphPad Software, Inc., La Jolla, CA).
Previous studies performed in ASA−/− animals during the peak of myelination showed a delayed expression of mRNA for myelin proteins: MBP, myelin and lymphocyte protein, and PLP, which suggests a defect in the differentiation program of OLs in the ASA−/− context (38). We have examined the expression of PDGFRα and MBP by immunohistochemistry in the cortex of ASA+/+ and ASA−/− mice during the period of active myelination at postnatal days 7, 14, and 21 (Fig. 1, A and B). Our analysis showed a reduced content of the PDGFRα (Fig. 1A) and MBP (Fig. 1B) in the ASA−/− cortex. Immunoblotting analysis confirmed the immunohistochemical findings. PDGFRα protein levels were significantly reduced in the mutant cortex at all ages examined (Fig. 1, C and F). MBP (Fig. 1, D and F) and PLP (Fig. 1E) were also significantly reduced in the ASA−/− brain.
To better understand the possible effect of ASA deficiency on the production of OLs, multipotential NPs were isolated from embryonic day 16.5 of ASA−/− and ASA+/+ mice, and NPs were differentiated for 7 days. ASA−/− NPs produced significantly fewer O4+ intermediate OLs (Fig. 2, A and B) and more GFAP+ astrocytes (Fig. 2, A and B) than cells with normal levels of ASA activity. A marked decrease of immature NG2+ OPCs was also determined in ASA−/− cells differentiated for 3 DIV (3.4 ± 0.6% in ASA−/− cells versus 8.3 ± 0.5% in ASA+/+ cells), indicating that the decrease in the production of oligodendroglial cells starts at early stages of in vitro differentiation. Sulfatide-treated ASA+/+ NPs showed a significant reduction in the numbers of O4+ cells as was observed in differentiated ASA−/− NPs (Fig. 2B). Immunoblotting analysis of protein extracts from differentiated ASA−/− NPs confirmed a failure of these cultures to produce OLs that express MBP (Fig. 2C). Measurement of four major sulfatide isoforms by UHPLC-MS/MS showed a 2-fold significant increase in sulfatides in ASA−/− NP spheres with respect to ASA+/+ NP spheres (Fig. 2D).
Because the PDGFRα is a critical signal for the generation of OPCs and their timely maturation into OLs, we studied its expression in ASA−/− NPs. PDGFRα is crucial for regulating the number and survival of OPCs/OL during myelination (9,–11, 39,–41), mainly by regulating the activity of the AKT or ERK1/2 pathway (42,–44). To determine whether this receptor was involved in the phenotype of ASA−/− cells, various experiments were performed. Immunoblotting analyses showed a significant reduction of PDGFRα protein levels in proliferating and differentiating ASA−/− cells (Fig. 3A). This decrease appeared specific to PDGFRα because control experiments did not show significant changes in levels of EGFR (Fig. 3A). Next, we measured the response of ASA−/− cells to the mitogenic stimulus of exogenous PDGF-AA. Differentiating ASA−/− and ASA+/+ NP cultures were incubated in the presence of PDGF-AA and pulsed with BrdU on day 2. Immunocytochemistry of NG2 showed significantly fewer BrdU+/NG2+ OPCs in ASA−/− cultures (Fig. 3B). Immunoblotting analysis of PDGFRα levels showed reduced levels of the receptor in PDGF-AA-stimulated NPs (Fig. 3C), confirming a failure of ASA−/− cells to respond to the growth factor. Functional studies of PDGFRα looking at AKT and MAPK ERK1/2 phosphorylation (Fig. 3, D–F) confirmed a severe failure of ASA−/− cells to phosphorylate serine 473 and threonine 308 residues of AKT upon stimulation with PDGF-AA (Fig. 3E).
Our previous experiments indicated a reduction of PDGFRα in ASA−/−-deficient NPs. This reduction could be caused by different defects affecting protein production (e.g. gene transcription), degradation (e.g. proteosome), and secretion (45,–47). Real-time PCR analyses of PDGFRα mRNA levels (Fig. 4A) and immunoblotting analyses for stress response markers (heat shock proteins HSP40 and HSP60; not shown) did not find significant differences in ASA−/− and ASA+/+ NPs. Furthermore, blockage of proteasome activity with MG132 and of lysosomal function with NH4Cl showed no significant differences in PDGFRα protein levels in ASA−/− cells grown in basal culture conditions or stimulated with PDGF-AA (Fig. 4, B and C). Together, these results indicate that defects in gene transcription, stress-related gene translation deficiencies, and altered protein degradation are unlikely to be causes for the reduction of PDGFRα protein levels in ASA−/− cells.
Receptor shedding via exosomal secretion has been shown to be an important mechanism by which cells regulate signaling with their environment (48). Exosomes are vesicles of ~20–100 nm in diameter generated through the formation of multivesicular bodies (49). Of relevance, OPCs and OLs are known to shed various proteins and lipids through exosomal secretion in vitro (50, 51). To determine whether PDGFRα is associated with this process, exosomes were prepared from conditioned cell culture media of ASA+/+ and ASA−/− NPs. Electron microscopy examination of exosomal preparations showed that ASA−/− and ASA+/+ have similar stereotypical profiles of circular vesicles with diameters ranging from ~25 to 100 nm (Fig. 4, D and E). Immunoblotting analysis of exosomal protein extracts showed significant increase in PDGFRα levels in ASA−/− exosomes (Fig. 4F), which represents about 3–4% of the reduction in PDGFRα observed in ASA−/− NPs. In contrast, receptor levels in ASA+/+ exosomes were significantly lower in ASA+/+ and absent in mock medium (Fig. 4, F–H). Treatment of ASA+/+ NPs with 10 μm exogenous sulfatides also led to an exosomal release of PDGFRα, although less intense than in ASA−/− cells, possibly due to ASA activity in ASA+/+ cells (Fig. 4F). Levels of EGFR in exosomes were significantly reduced in ASA−/− when normalized against the exosomal housekeeping protein Alix but not against Rab-5b (Fig. 4, G and H). This may reflect compositional changes in exosomes from mutant cells.
Association of PDGFRα with DRMs is needed for its function in OPCs (33, 34). To examine whether this association was affected in ASA−/− cells, a Lubrol-sucrose gradient method that fractionates plasmalemma DRMs from other non-DRM membranes (lysosomes, Golgi apparatus, etc.) was used (52). Immunoblotting analyses showed a sharp reduction (~5% of the total level of receptor) of PDGFRα in DRMs (fractions 4–6) from ASA−/− cells (Fig. 5A). DRM markers flotillin 2 and caveolin 1 were also significantly reduced in mutant DRMs (Fig. 5B). Interestingly, exposure of ASA+/+ cells to sulfatides led to reductions of PDGFRα in DRMs (Fig. 5A). The total level of cholesterol (another DRM compound) and its buoyancy were not significantly different between ASA−/− and ASA+/+ (Fig. 5C). Sulfatides were significantly increased in DRM fractions but decreased in non-DRM fractions from ASA−/− cells (Fig. 5D). Together, these results suggest that changes in DRM composition occur in ASA−/− cells, contributing to a reduction of PDGFRα in the plasma membrane.
To evaluate whether gliogenic and receptor deficits can be corrected in ASA−/− cells, enzyme replacement was done by treating ASA−/− cells in culture medium conditioned from ASA prepared from ASA-overexpressing cells (ASACM) (36). This cross-correction experiment takes advantage of the general property of lysosomal enzymes to be taken up by cells, primarily via the endocytic mechanism involving a mannose 6-phosphate receptor (53). Cross-correction using this approach has been well established with a significant number of lysosomal enzymes, including ASA (54). ASA−/− NP spheres were incubated with 5, 10, or 25% ASACM before analysis. Immunocytochemical detection of ASA protein by confocal microscopy showed intracellular presence of ASA in corrected ASA−/− cells (Fig. 6A). ASA enzyme activity was substantially recovered (data not shown). Although sulfatide content was not normalized, there was a significant recovery of the normal ratio of long versus short fatty acid sulfatides in cells cultured with 25% ASACM (Fig. 6B).
Levels of PDGFRα were significantly normalized in corrected ASA−/− cells (Fig. 6C). Recovery of PDGFRα led to significant relocalization of the receptor in DRM fractions and less exosomal shedding (Fig. 6D). A significant recovery of AKT phosphorylation at serine residue 473 was observed but not in ERK1/2 phosphorylation (Fig. 6, E and F). Upon differentiation, production of O4+ OLs from corrected ASA−/− cells was significantly normalized (Fig. 6G).
Our experiments of exosomal shedding are relevant because this may be a regulatory mechanism used by OLs to locally and rapidly control levels of receptor available at the cell surface. To study this process in more detail, we measured receptor levels in exosomes isolated from primary glial cultures, CG4 cells (central glial-4 cells) and in the murine cortex. First, mixed glial cultures were prepared from newborn ASA+/+ and ASA−/− pups and maintained for 7, 14, and 21 DIV, to enrich immature A2B5+/NG2+ OPCs (7 DIV) or intermediate O4+, O1+, and mature MBP+ OLs (14 and 21 DIV), respectively (28, 55). Quality control of OL differentiation in these conditions was confirmed by Western blot analysis of MBP expression (Fig. 7A). Whereas ASA+/+ glial cultures showed the expected increase of MBP during culture differentiation, OL differentiation was markedly reduced in ASA−/− glial cultures (Fig. 7A). Exosomes were isolated from conditioned medium at each time point and analyzed for PDGFRα content by immunoblotting. PDGFRα was found in low but significant levels in exosomes from ASA+/+ glia (Fig. 7, B and C). As expected, PDGFRα was increased in exosomes from ASA−/− glia (Fig. 7, B and C). A similar analysis was done with conditioned medium from cultures of oligodendroglial CG4 cells. Cells were grown in proliferating conditions for 5 DIV before analysis of exosomes. Immunodetectable levels of PDGFRα were present in the conditioned medium (Fig. 7D), indicating that this process occurs in OPCs independently of astrocytes being present.
Finally, we examined the extent to which exosomal shedding of PDGFRα occurs during myelination in the mouse brain. For this, we adapted a previously described technique (34). Transmission electron microscopy imaging confirmed the exosomal ultrastructure of these preparations from ASA+/+ and ASA−/− brain material (Fig. 7E). Subsequent immunoblotting analysis showed the presence of PDGFRα (Fig. 7F) in brain exosomes from both genotypes. Exosomal shedding of PLP was used as a quality control for our experiments (Fig. 7G). This analysis showed a peak of the receptor in exosomes isolated from the cortex of P14 ASA+/+ brain (Fig. 7F), a trend that was also observed in primary glial cell cultures (Fig. 7C). In contrast, PDGFRα levels were higher in exosomes from the ASA−/− cortex. Altogether, these results provide evidence that PDGFRα is exosomally secreted in vivo during the peak of myelination. Contextual changes, such as sulfatide accumulation, that alter this process may be relevant to understand possible defects in myelination in the brains of metachromatic leukodystrophy patients and in other demyelinating conditions.
This study provides insight into the interaction between sulfatides and PDGFRα activity in NPs. Experimental conditions that genetically or exogenously lead to increased sulfatide content revealed 1) defects in the association of PDGFRα with DRMs, 2) impaired translation of PDGF-AA signaling via repression of PDGFRα/AKT activation, 3) increased shedding of PDGFRα via exosomes, and 4) decreased formation of NG2+ OPCs and O4+ OLs.
Various studies have shown that OL differentiation could either be promoted by a deficient synthesis of sulfatides (56, 57) or inhibited by anti-sulfatide antibodies (58, 59), suggesting that sulfatides negatively regulate OL differentiation (20, 21). However, the mechanism of this repression has remained unaddressed. Our results using ASA−/− NPs with intrinsically high levels of sulfatides support the idea that these lipids are associated with reduced capacity to generate OLs, in line with previous in vivo analyses (60).
Although sulfatides are major components of mature myelin, they are synthesized during neural development in two distinct waves, first of short and then long fatty acid sulfatides, coinciding with the appearance of immature OPCs and more differentiated OLs, respectively (61). The physiological relevance of this change remains unclear. Recently published studies from our laboratory showed that short (C16:0 and C18:0) and long (C24:0 and C24:1) fatty acid sulfatides are components of DRMs isolated from ASA+/+ and ASA−/− brains and that their relative abundance is developmentally regulated (32). We propose that variations in short/long sulfatide isoforms may affect the fluidity of DRMs and induce changes in PDGFRα signaling (60, 62, 63). Our findings of a significant reduction of the ratio of long versus short fatty acid sulfatides associated with a normalization of the PDGFRα levels and AKT phosphorylation in ASA-corrected NPs support this hypothesis.
PDGFRα regulates the proliferation, survival, and differentiation of OPCs and OLs (10, 11, 39, 64, 65), a function that is dependent on its association with DRMs (16). Interestingly, PDGFRα membrane localization responds to changes in sphingolipid content (66). For example, ganglioside GM1 induced the relocation of PDGFRα to non-DRM/caveolar membranes, inhibiting PDGFRα signaling (67). Our study identified sulfatides as another group of sphingolipids with the capacity to reduce the association of the PDGFRα with DRMs and, consequently, impair the sensitivity of ASA−/− NPs to PDGF-AA. This defect did not appear to be caused by a reduction in gene transcription, increased degradation, or activation of cellular stress responses to the accumulation of sulfatides but rather to the impaired activity of the PDGFRα/AKT pathway. This most likely stems from lowered association of the receptor with DRMs in ASA−/− NPs. Interestingly, the survival/expansion of OPC mediated by PDGF-AA involves the activation of AKT through phosphorylation of residues Thr308 and Ser473 (68, 69). Our results demonstrate that in the presence of high levels of sulfatides, the relocalization of PDGFRα to non-DRM domains and phosphorylation of Thr308 and Ser473 of AKT are severely compromised. These findings further underline the importance of DRM integrity in AKT signaling activation (70, 71). Although the molecular mechanism by which sulfatides reduce the association of the receptor in DRMs is unclear, the involvement of sulfatides in this process is demonstrated by its normalization observed after correction of the ASA deficiency and the reduction in the ratio of long and short sulfatides in treated ASA−/− NPs. Further experimentation is under way to study the molecular mechanism mediating this effect.
Our study identifies a novel cellular mechanism contributing to the regulation of the PDGFRα in NPs via exosomal shedding. The amount of receptor secreted by exosomes corresponds to about 3–4% of the PDGFRα reduction in ASA−/− NPs, which is similar to the fraction of receptor associated with DRM. These findings suggest that the fraction of receptor associated with DRM is subjected to exosome control. Low levels of PDGFRα in DRMs may be sufficient to interfere with critical survival/proliferation signaling in OPCs. This would weaken the capacity of the ASA−/− cultures to generate normal numbers of OPCs and OLs and explain the reduction on PDGFRα levels.
Exosomes are intralumenal vesicles contained in multivesicular bodies playing important roles in cell-cell communication (72,–75), and their shedding is a physiological relevant process. For example, the transferrin receptor and the myelin protein PLP undergo exosomal secretion during differentiation of erythrocytes and OLs, respectively (26, 51, 76). Sphingolipids contribute to regulating the formation of exosomes (77, 78), and cholesterol and ceramides promote exosome secretion (76, 79,–81). Our experiments suggest a role for sulfatides in exosomal biogenesis. Intrinsic increases in sulfatides, such as in ASA−/− NPs, and, to a lesser extent, acute exposure to exogenous sulfatides promoted the shedding of PDGFRα. Exosomal shedding of PDGFRα in glial cells has not been reported previously, but this secretory mechanism may be a relevant physiological process by which OPCs rapidly and locally regulate the concentration of receptor. In turn, this may have an immediate impact on the receptor-mediated responses to proliferation, survival, and/or differentiation.
How PDGFRα translocates to exosomes and is targeted for removal is still unclear. We hypothesize this process could involve changes in lipid-protein composition in membrane domains where the receptor is assembled. The changes in long versus short fatty acid sulfatides observed in DRMs in our study appear to support this. At this moment, excluding the EGFR, the possibility that other receptors are fated to exosomal secretion cannot be ruled out. Confocal microscopy studies suggested co-localization of PDGFRα with sulfatides in ASA+/+ and to a lesser extent in ASA−/− NPs (data not shown). There is evidence that sulfatides are components of exosomal membranes (82). However, exosomes isolated from NPs appeared to contain little if any sulfatide (data not shown). This last observation suggests that PDGFRα and sulfatides might use different intracellular vesicular trafficking compartments. The reduction of PDGFRα from DRMs upon increased concentrations of sulfatides supports this conclusion.
In vivo studies in the ASA+/+ cortex showed an increase in exosomal secretion of PDGFRα before the peak of myelination. These results provide further evidence that exosomal regulation of PDGFRα might be part of a larger regulatory program involved in myelination. Sulfatides may contribute to the control of the local amount of PDGFRα available in the OPC cell body to respond effectively to PDGF-AA during myelination. In vivo, exosomal shedding may involve other mechanisms, including axonal signals, nerve conduction, and astrocytic and microglial signals (83,–88). In this scenario, one of the events occurring when OPCs stop dividing and start myelination is the production of large quantities of sulfatides to cope with the production of the myelin membrane. Such conditions may activate the exosomal loss of PDGFRα from OPCs to coordinate its postmitotic cycle and myelinating program. Studies by Trajkovic et al. (50) and Bakhti et al. (51) demonstrating a fundamental role of endosomes and exosomes during OL maturation and myelination appear to support this model. Whether this mechanism is also relevant in tethering OL numbers during CNS development and remyelination is currently unknown.
In the context of the ASA deficiency, our study provides an initial understanding of how ASA deficiency and sulfatides may contribute to demyelination and remyelination defects in the human lysosomal storage disease metachromatic leukodystrophy (60). Our observations may also be extrapolated to other demyelinating diseases, such as multiple sclerosis, where demyelination could lead to the availability of myelin debris, transforming myelin-derived sulfatides into environmental toxins (31, 89,–91) that may impair remyelination. In this regard, other myelin components, such as myelin-associated glycoprotein, myelin oligodendrocyte glycoprotein, and Nogo A, are known to exert inhibitory functions, decreasing the regenerative capacity of the brain (92,–94).
Our work may also extend to non-neural situations. For example, sulfatides are also expressed in non-nervous tissues, such as renal and pancreatic tissues, where they have been described as participating in the secretion of insulin (95,–97), and they are elevated in a wide range of cancers, where they are thought to participate in metastasis (98,–100). In some of these diseases, sulfatides may contribute to pathology by altering the regulation of local signals via changes in DRMs and/or exosomal secretion. Because sulfatides comprise a heterogeneous population of lipids, future studies will need to determine the physiological role of the different sulfatide isoforms.
We thank Zhiyuan Sun for technical assistance in mass spectrometry and Dr. Gerardo Morfini for relevant experimental discussions.
*This work was supported, in whole or in part, by National Institutes of Health Ruth L. Kirschstein National Research Service Award F31 (to K. P. C.). This work was also supported by National Multiple Sclerosis Society Grants PP1516 and RG 4439-A-2 and by Department of Defense Contract W81XWH-11-1-0198 (to M. I. G.).
2The abbreviations used are: