Myelination is a biosynthetically demanding process in which mTORC1, the gatekeeper of anabolism, occupies a privileged regulatory position. We have shown previously that loss of mTORC1 function in Schwann cells (SCs) hampers myelination. Here, we genetically disrupted key inhibitory components upstream of mTORC1, TSC1 or PTEN, in mouse SC development, adult homeostasis, and nerve injury. Surprisingly, the resulting mTORC1 hyperactivity led to markedly delayed onset of both developmental myelination and remyelination after injury. However, if mTORC1 was hyperactivated after myelination onset, radial hypermyelination was observed. At early developmental stages, physiologically high PI3K-Akt-mTORC1 signaling suppresses expression of Krox20 (Egr2), the master regulator of PNS myelination. This effect is mediated by S6K and contributes to control mechanisms that keep SCs in a not-fully differentiated state to ensure proper timing of myelination initiation. An ensuing decline in mTORC1 activity is crucial to allow myelination to start, while remaining mTORC1 activity drives myelin growth.https://doi.org/10.7554/eLife.29241.001
Neurons transmit electrical impulses throughout our body along cable-like structures called axons. Similar to electric cables, the axons are enveloped in an insulating sheath called myelin, which makes the impulses travel faster down the axon. If myelin does not form correctly or gets lost later in life, it can lead to muscle weakness and numbness.
Myelin is produced in nerves by specialized cells called the Schwann cells, which wrap around the axons several times to create a thick myelin sheath. The signaling complex called mTORC1 plays an important role in this process. A study in 2014 showed that when mTORC1 was inactive, the myelin sheath was abnormally thin. However, it was not known whether increased mTORC1 would force Schwann cells to produce more myelin.
Now, Figlia et al. – including some of the researchers involved in the 2014 study – used genetically modified mice to manipulate two proteins known to control the activity of mTORC1. When both proteins were removed, either individually or in combination, mTORC1 activity was higher than normal. However, in developing nerve cells, high levels of mTORC1 did not cause the young Schwann cells to produce more myelin, but rather stopped them to become the specialized cells that wrap around axons. Figlia et al. then increased mTORC1 levels after Schwann cells had already started wrapping around axons. In this case, a high activity of mTORC1 resulted in thicker myelin. This suggests that a normal development of a nerve and ultimately the thickness of the myelin sheath depend on when and how much of mTORC1 is available.
This discovery could help to develop new therapies for myelin diseases. Increasing the activity of mTORC1 in Schwann cells after they have started wrapping may boost myelin production in diseases in which the myelin sheath is too thin. Conversely, inhibiting mTORC1 could help in situations when the Schwann cells cannot develop properly.https://doi.org/10.7554/eLife.29241.002
Myelin is a lipid-rich membrane specialization of Schwann cells (SCs) and oligodendrocytes (OLs) winding around axons to accelerate the propagation of action potentials (Hartline and Colman, 2007). The geometrical properties of each myelin segment, such as its thickness and length, are essential to achieve fast and accurate conduction along myelinated fibers (Waxman, 1980). Conversely, myelin geometry is deranged in numerous diseases of the nervous system, including neuropathies and multiple sclerosis. To produce myelin segments with correct thickness and length, the plasma membrane of myelinating cells undergoes a remarkable, highly regulated expansion. It was estimated that the SC membrane has to expand several thousand fold during myelination (Chrast et al., 2011; Webster, 1971) and that OLs produce myelin equal to three times their own weight every day (Norton and Poduslo, 1973). Consequently, myelination is an anabolically demanding task.
The mechanistic target-of-rapamycin (mTOR), a serine-threonine protein kinase, is the core component of mTOR complex 1 (mTORC1) and mTOR complex 2 (mTORC2). Among them, mTORC1 is the gatekeeper of multiple anabolic reactions, including mRNA translation and biosynthesis of lipids, purines, and pyrimidines (Ben-Sahra et al., 2013; Ben-Sahra et al., 2016; Porstmann et al., 2008; Thoreen et al., 2012). The mTORC1 pathway is tightly connected to PI3K-Akt signaling downstream of growth factors. Upon growth factor stimulation, Akt activates mTORC1 by phosphorylating inhibitors of mTORC1, the TSC complex and PRAS40 (Inoki et al., 2002; Manning et al., 2002; Sancak et al., 2007; Vander Haar et al., 2007). The TSC complex is composed of the subunits TSC1, TSC2, and TBC1D7 (Dibble et al., 2012), and functions as a GTPase-activating unit towards the small GTPase Rheb, a potent activator of mTORC1 when loaded with GTP (Tee et al., 2003). Accordingly, disruption of the TSC complex causes mTORC1 hyperactivation in a wide range of tissues and cell types (Byles et al., 2013; Castets et al., 2013; Kwiatkowski et al., 2002).
Consistent with the intense anabolic challenge posed by myelination, the PI3K-Akt-mTORC1 axis has emerged as a fundamental player in PNS and CNS myelination (Taveggia, 2016; Wood et al., 2013). Genetic disruption of mTORC1 in SCs or OLs impaired myelination, demonstrating that mTORC1 function is required for this process (Bercury et al., 2014; Lebrun-Julien et al., 2014; Norrmén et al., 2014; Sherman et al., 2012; Wahl et al., 2014; Zou et al., 2014). However, studies aimed at examining the consequences of increased mTORC1 activity, predicted to augment myelin growth, have yielded conflicting results. Overexpression of constitutively active Akt or hyperactivation of the PI3K-Akt pathway by deletion of PTEN in OLs caused hypermyelination (Flores et al., 2008; Goebbels et al., 2010), while deleting TSC1 was detrimental to OL myelination (Jiang et al., 2016; Lebrun-Julien et al., 2014). In SCs, hyperactivation of the PI3K-Akt pathway with various approaches did not lead to univocal results (Cotter et al., 2010; Domènech-Estévez et al., 2016; Flores et al., 2008; Goebbels et al., 2012). Thus, we aimed here at elucidating the functional roles of mTORC1 activation in SCs during various steps of development, in homeostasis, and after injury. Based on our results and reconciling also previous reports, we propose a model suggesting that mTORC1 signaling exerts multiple distinct roles at different stages of SC differentiation.
Using a loss-of-function approach, we have previously discovered that mTORC1 – but not mTORC2 – promotes myelin growth in the PNS (Norrmén et al., 2014). To further define the role of mTORC1 in myelination, we have now pursued the converse approach, hyperactivating mTORC1 by conditional ablation of TSC1, a strategy that destabilizes the TSC complex (Dibble et al., 2012). SC-specific TSC1 mutants were generated by crossing mice harboring a floxed allele of Tsc1 (Kwiatkowski et al., 2002) with mice expressing a Cre transgene under control of regulatory sequences of the Dhh gene (Jaegle et al., 2003) (DhhCre:Tsc1KO). We first confirmed, by western blot analysis of postnatal day 5 (P5) sciatic nerves, that TSC1 was successfully depleted (Figure 1a, Figure 1—figure supplement 1a). We then assessed mTORC1 activity by analyzing the phosphorylation levels of its downstream effectors. As expected, phosphorylation of two well-established mTORC1 targets, S6K and 4EBP1 (Hay and Sonenberg, 2004), was increased in TSC1-mutant nerves, together with phospho-S6S235/236 levels, a target of S6K (Figure 1a,b, Figure 1—figure supplement 1a). As additional evidence of mTORC1 hyperactivation, we found that cultured mutant SCs isolated from either dorsal root ganglia (DRG) or postnatal nerves were enlarged, consistent with the prominent role of this pathway in cell size control (Lloyd, 2013) (Figure 1c, Figure 1—figure supplement 2a,b).
Next, we assessed the extent of myelination by electron microscopy (EM). Surprisingly, P5 DhhCre:Tsc1KO nerves exhibited a strong reduction in myelinated fibers due to an arrest of most SCs at the promyelinating stage (Figure 1d,e). No overt defect in radial sorting was evident, thus indicating a bona fide impairment in the onset of myelination. The percentage of myelinated fibers progressively increased with time, almost doubling by P14. By P60, most fibers were ultimately myelinated, although occasional promyelinating SCs were still present (Figure 1d,e). Additionally, the myelinated nerve fibers were hypomyelinated, presumably as a consequence of delayed onset of myelination (Figure 1d; for quantification, see Figure 6l). Impaired SC differentiation was reflected in reduced levels of myelin protein P0, while cJun and Oct6 – both highly expressed in promyelinating SCs – were upregulated (Figure 1—figure supplement 2c,d). Consistent with a failure of mutant cells to promptly differentiate, we also detected an increase in proliferating Sox10-positive SCs and, consequently, we found overall more SCs (P3; Figure 1f–h).
Non-mTORC1 related functions of the TSC complex have been reported (Neuman and Henske, 2011). Thus, we assessed whether the phenotype of DhhCre:Tsc1KO mice was genuinely due to high mTORC1 activity by treating mice with the mTORC1-inhibiting drug rapamycin. Rapamycin was administered at P3 and P4 and the mice analyzed at P5. Despite the short course of treatment, we found a robust increase in myelinated fibers in rapamycin-treated, compared to vehicle-only treated, mutant nerves (Figure 1i,j). The morphological rescue was paralleled by a decrease in cJun and an increase in P0, roughly back to the levels of control nerves, together with the expected suppression of S6K phosphorylation at the mTORC1-sensitive site T389 (Figure 1k, Figure 1—figure supplement 1b). In line with the in vivo results, we found that in vitro myelination of TSC1 mutant-derived DRG explant cultures was strongly defective, but could be remarkably improved by acute treatment with rapamycin, consistent with a PNS-autonomous origin of the in vivo rescue (Figure 1—figure supplement 2e,f).
Collectively, our data show that high mTORC1 activity following deletion of TSC1 in SCs has, paradoxically, a detrimental effect on PNS myelination by delaying the transition from promyelinating to myelinating SCs.
The PI3K-Akt pathway is a key upstream driver of mTORC1 and, in turn, mTORC1 activation dampens the PI3K-Akt pathway through multiple inhibitory feedback loops (Efeyan and Sabatini, 2010). To explain the discrepancy between the effects of high mTORC1 signaling in SCs lacking TSC1 and the positive role generally attributed to PI3K-Akt signaling in PNS myelination (Taveggia, 2016), we reasoned that overactive mTORC1 may suppress the PI3K-Akt pathway, and consequently SC differentiation, via the aforementioned feedback loops. To test this hypothesis, we assessed phosphorylation of Akt and the upstream receptor ErbB2 in DhhCre:Tsc1KO nerves. Since the MAPK pathway can also be subjected to feedback inhibition by mTORC1 (Carracedo et al., 2008), and considering the crucial functions of this pathway in SC biology (Ishii et al., 2013; Newbern et al., 2011; Sheean et al., 2014), we also examined the phosphorylation status of Erk1/2. No major changes in ErbB2 or Erk1/2 phosphorylation could be detected (Figure 2a, Figure 2—figure supplement 1a). By contrast, we observed a strong reduction in Akt phosphorylation at both T308 and S473, together with a global decrease of Akt substrates phosphorylation (Figure 2a, Figure 2—figure supplement 2a and Figure 2—figure supplement 1a). Moreover, pharmacological inhibition of the PI3K-Akt axis, but not the Mek-Erk axis, abolished S6 phosphorylation upon neureugulin-1 stimulation of primary SCs, although some minor contribution of the Mek-Erk axis to mTORC1 activation was also apparent in this experimental setting (Figure 2—figure supplement 2b). Together, these results demonstrate that in SCs mTORC1 is mainly activated by the PI3K-Akt pathway, and that, in turn, high mTORC1 activity suppresses Akt activation without significantly affecting the MAPK pathway or activation of ErbB2-ErbB3 receptors.
To address whether the observed Akt suppression is responsible for the defective myelination due to TSC1 deletion in SCs, we sought to restore Akt activity in TSC1 mutants by simultaneously deleting the phospholipid phosphatase PTEN which limits activation of the PI3K-Akt pathway (Song et al., 2012) (Figure 2b). Given the known tumor suppressor role of both TSC1 and PTEN (Inoki et al., 2005), we used the more restricted MpzCre mouse line (Feltri et al., 1999) to generate suitable single or double mutants, referred to as MpzCre:Tsc1KO, MpzCre:PtenKO, and MpzCre:Tsc1KO:PtenKO. Both proteins could be successfully depleted (Figure 2—figure supplement 2c,d). As anticipated, co-deletion of TSC1 and PTEN released the previously observed suppression on the PI3K-Akt pathway upon single deletion of TSC1, indicated by restored phosphorylation of Akt at T308 (Figure 2c, Figure 2—figure supplement 1b). We also noticed, based on the resulting levels of phospho-S6S235/236, that MpzCre:Tsc1KO:PtenKO nerves contained higher mTORC1 activity compared to MpzCre:Tsc1KO (Figure 2c, Figure 2—figure supplement 1b). To explain this finding, we hypothesized that the high Akt activity in MpzCre:Tsc1KO:PtenKO may further increase mTORC1 activity via phosphorylation and subsequent inhibition of PRAS40, an inhibitory protein associated with mTORC1 (Sancak et al., 2007) (Figure 2b). In agreement, MpzCre:Tsc1KO:PtenKO nerves displayed increased PRAS40 phosphorylation at the Akt-dependent phospho-site T246 compared to MpzCre:Tsc1KO nerves (Figure 2c, Figure 2—figure supplement 1b). On the morphological level, P5 MpzCre:Tsc1KO:PtenKO nerves looked profoundly larger than those of controls or single mutants (Figure 2d). Nevertheless, EM analysis revealed no detectable improvement in SC differentiation compared to single mutants, but rather an aggravation of the phenotype with SCs in contact with sorted axons arrested at the promyelinating stage and virtually no myelinated fibers (Figure 2e,f). Unexpectedly, closer inspection of P5 MpzCre:PtenKO mice revealed also a substantial decrease in myelinated fibers compared to controls due to SCs arrested at the promyelinating stage. Consistent with the morphological findings, cJun levels were comparably increased in both single knockout nerves compared to controls, and significantly more increased in double knockout nerves compared to single mutants (Figure 2g, Figure 2—figure supplement 1c).
These findings indicated that reduced Akt signaling due to mTORC1 hyperactivation in MpzCre:Tsc1KO is likely not responsible for the observed impaired SC differentiation. On the other hand, the data also suggested that hyperactivation of the PI3K-Akt pathway early in development is per se detrimental to SC differentiation, potentially via mTORC1. In line with these conclusions, we found that SC-specific expression of constitutively active (myristoylated) Akt1, Akt2, or Akt3 in DRG-explant cultures inhibited myelination (Figure 2h, Figure 2—figure supplement 2e). To corroborate our findings and to determine the aforementioned potential role of high mTORC1 signaling in PTEN mutants, we generated PTEN knockout mice using DhhCre-mediated recombination (DhhCre:PtenKO) and treated the resulting animals with vehicle or rapamycin (treatments at P3 and P4, analyses at P5). As observed in MpzCre:PtenKO mice previously (Figure 2f), also vehicle-treated DhhCre:PtenKO animals showed a decrease in myelinated fibers compared to controls, and rapamycin treatment effectively improved the DhhCre:PtenKO phenotype (Figure 2i,j). In line with the morphological rescue, rapamycin also reduced expression of Oct6 and cJun (Figure 2k, Figure 2—figure supplement 1d).
To elucidate the molecular basis of how hyperactive mTORC1 impairs SC differentiation, we compared the transcriptomes of P5 DhhCre:Tsc1KO and DhhCre:PtenKO nerves, both displaying high mTORC1 activity, with age-matched nerves in which mTORC1 activity has been eliminated by conditional ablation of Raptor in SCs (DhhCre:RptorKO) (Norrmén et al., 2014), together with controls. Unbiased hierarchical clustering showed that the replicates within each genotype group closely clustered together, thus validating comparisons. Notably, DhhCre:Tsc1KO and DhhCre:PtenKO samples formed a distinct cluster compared to the other groups, confirming also at a global transcriptional level the previously described similarities (Figure 3a). Gene ontology (GO) analyses of DhhCre:Tsc1KO and DhhCre:PtenKO data sets revealed a particular upregulation of transcripts associated with cell proliferation compared to controls (Figure 3—figure supplement 1a,b), consistent with our previous observations (Figure 1f,g) and with other reports (Goebbels et al., 2010). In addition, downregulated lipid biosynthesis-associated mRNAs were overrepresented in DhhCre:RptorKO compared to controls (Figure 3—figure supplement 1c), in line with our previous findings (Norrmén et al., 2014).
Given our focus on cell differentiation, we scrutinized our datasets to identify transcription factors (TFs) potentially regulated by mTORC1. To this end, we selected two subsets out of significantly changed mRNAs (fold change >1.2, FDR < 0.05): (1) Upregulated in both DhhCre:Tsc1KO and DhhCre:PtenKO (i.e. high mTORC1 activity), but downregulated in DhhCre:RptorKO (i.e. no mTORC1 activity) or (2) Downregulated in DhhCre:Tsc1KO and DhhCre:PtenKO, but upregulated in DhhCre:RptorKO. Applying these criteria, we identified 110 putative mTORC1-induced genes (subset 1) and 131 potential mTORC1-repressed genes (subset 2), among which 7 and 16 encoded TFs, respectively (Figure 3b,c). Among the mTORC1-repressed TFs emerged Krox20 (also known as Egr2), a crucial TF for the differentiation of myelinating SCs (Svaren and Meijer, 2008; Topilko et al., 1994). Thus, we reasoned that Krox20 may link mTORC1 activity to the onset of SC myelination. First, we confirmed the RNA-sequencing results with qRT-PCRs for Krox20 and other TFs with well-established roles in PNS myelination (cJun, Oct6, Brn2, Sox10, Sox2, Id2). Only Krox20 displayed a pattern consistent with stringent mTORC1-dependent regulation, being downregulated in both DhhCre:Tsc1KO and DhhCre:PtenKO nerves and mildly, but significantly upregulated in DhhCre:RptorKO nerves at P5 and P8 (Figure 3d, Figure 3—figure supplement 1d). Furthermore, protein levels of Krox20 were also increased in Raptor mutants and decreased in TSC1 mutants (P5; Figure 3e, Figure 3—figure supplement 2a). Krox20 downregulation correlated with the extent of mTORC1 hyperactivation: Krox20 levels were comparably reduced in mice with single deletion of TSC1 or PTEN compared to controls and barely detectable in double knockout mice at P5 (Figure 3f, Figure 3—figure supplement 2b). In further support of the critical relationship between Krox20 and the PI3K-Akt-mTORC1 axis, we found that acute inhibition of mTORC1 either with rapamycin or torin-1, an ATP-competitive inhibitor of mTOR, strikingly increased basal Krox20 expression in cultured SCs, as did pharmacological inhibition of PI3K with LY294002 (Figure 3g,h). Suppression of mTORC1 or PI3K also enhanced cyclic-AMP induced upregulation of Krox20 (Figure 3i). We then examined the roles of the classical mTORC1 targets, 4EBPs and S6K, in regulating Krox20. Overexpression of a constitutively active version of 4EBP1 (i.e. cannot be phosphorylated by mTORC1) did not significantly affect Krox20 expression (Figure 3j). In contrast, inhibition of S6K with LYS6K2 moderately increased basal Krox20 expression and partially rescued the defective myelination in DRG-explant cultures from DhhCre:Tsc1KO animals (Figure 3k,l, Figure 3—figure supplement 1e).
In conclusion, this set of data provides converging evidence that Krox20 is an mTORC1-regulated transcript, and that S6K is part of the mechanisms linking mTORC1 activity to Krox20 expression. Our results indicate that high mTORC1 activity suppresses expression of Krox20, the main driver of SC myelination, thereby inhibiting onset of myelination.
Our results so far show a peculiar inhibitory role of high mTORC1 signaling in SC differentiation. In this light, mTORC1 activity in physiological conditions would be expected to be high before onset of myelination and to decrease as SCs differentiate into myelinating SCs. Thus, we evaluated mTORC1 and Akt activities throughout mouse nerve development. Consistently, we found high activity of mTORC1, reflected by the phosphorylation status of S6, from embryonic day 17.5 (E17.5) to P1, while Akt activation peaked at P1. In both cases, highest levels preceded detection of P0 and MBP (Figure 4a,b), in line with recent data obtained from postnatal rats (Heller et al., 2014). Conversely, as mTORC1 activity declined between E17.5 and P5, the levels of myelin proteins increased, as did the levels of Krox20 mRNA (Figure 4c), consistent with negative regulation of Krox20 expression by mTORC1. To examine mTORC1 activity in early nerve development at cellular resolution, we performed immunohistochemistry at P1. The majority of SCs highly expressing phospho-S6 were not yet myelinating (80.09 ± 2.35%, mean ±s.e.m., n = 4 mice) (Figure 4d, inset 1), while weaker phospho-S6 staining was found in association with myelinated fibers (Figure 4d, inset 2), consistent with previous cell culture data (Heller et al., 2014). High mTORC1 activity was often observed in arrangements reminiscent of axon bundles. These assemblies consist of SCs surrounding multiple axons and extending cytoplasmic processes to sort large caliber axons prior to myelination in a process called radial sorting. Consistently, the temporal span of high mTORC1 activity coincides with the period of intense radial sorting (Figure 4a,b). Radial sorting was impaired when mTORC1 was disrupted in DhhCre:RptorKO nerves (Norrmén et al., 2014), indicating that the high mTORC1 activity observed in early nerve development is required in this process. Thus, we examined P5 MpzCre:Tsc1KO:PtenKO nerves in which we had observed the highest mTORC1 activity (Figure 2c) for radial sorting alterations. We found that bundles often contained fewer axons than in control nerves (Figure 4e). Coherent with this finding, the number of sorted axons was significantly higher compared to controls (Figure 4f), indicative of increased radial sorting.
Collectively, we conclude that: (1) In normal nerve development, high mTORC1 activity is present before the onset of myelination and declines as SCs start myelinating; (2) The decrease in mTORC1 activity is physiologically required to allow SCs to differentiate into myelinating SCs, based on the findings that SC differentiation is impaired by sustained activation of mTORC1 in TSC1 and/or PTEN mutants; (3) High mTORC1 activity inhibits the differentiation of myelinating SCs, but promotes radial sorting, possibly to ensure that the differentiation program of myelination is activated only after radial sorting is completed.
According to our timeline analysis, the activity of the PI3K-Akt-mTORC1 axis in adult nerves is substantially lower than in early development (Figure 4a,b). Thus, we analyzed and compared systematically the effects of differently increased mTORC1 and/or PI3K-Akt signaling in adult SCs. To this end, we crossed our floxed mice with mice carrying a MpzCreERT2 transgene, thus allowing inducible SC-specific ablation of TSC1 and/or PTEN (referred to as MpzCreERT2:Tsc1KO, MpzCreERT2:PtenKO, and MpzCreERT2:Tsc1KO:PtenKO). Tamoxifen was administered to young adult mice and three months later (months post-tamoxifen, mpt) TSC1 and/or PTEN protein levels were substantially reduced in the corresponding nerves (Figure 5a–c, Figure 5—figure supplement 1a–c). Western blot analyses for phospho-S6KT389 and phospho-AktT308 showed that, like in development, deletion of TSC1 in adult nerves resulted also in hyperactivation of mTORC1 and suppression of Akt activation, while deletion of PTEN or combined deletion of TSC1 and PTEN hyperactivated both PI3K-Akt and mTORC1 signaling. Notably, MpzCreERT2:Tsc1KO:PtenKO mice displayed higher mTORC1 levels than either single knockout animals, reminiscent of development (Figure 5d, Figure 5—figure supplement 1d). Macroscopically, MpzCreERT2:PtenKO and MpzCreERT2:Tsc1KO:PtenKO nerves were enlarged compared to controls and MpzCreERT2:Tsc1KO nerves at 3 mpt (Figure 5e). At the ultrastructural level, we found no signs of demyelination in controls or single mutants, while MpzCreERT2:Tsc1KO:PtenKO nerves contained small numbers of demyelinated fibers (174 ± 45 per sciatic nerve cross section, mean ±s.e.m., n = 3 mice per genotype). Next, we evaluated myelin thickness expressed as g-ratio, i.e. the ratio between the axon diameter and the fiber diameter (axon plus myelin). All mutant nerves showed radial hypermyelination (i.e. increased myelin thickness without visible structural abnormalities) as indicated by lower g-ratios, with MpzCreERT2:Tsc1KO:PtenKO animals being significantly more hypermyelinated than single knockouts (Figure 5f,g). In TSC1 mutants, radial hypermyelination was more pronounced in fibers with an axonal diameter smaller than 2 μm, whereas in MpzCreERT2:PtenKO and MpzCreERT2:Tsc1KO:PtenKO nerves, axons of all sizes were significantly affected (Figure 5h). We did not observe substantial major shifts in axonal size distributions, pointing to a genuine increase in myelin thickness as reason for reduced g-ratios (Figure 5i). The observed radial hypermyelination was persistent at 6 mpt in MpzCreERT2:Tsc1KO and MpzCreERT2:PtenKO nerves (Figure 5—figure supplement 2a–d). Intriguingly, at 6 mpt, PTEN mutants were more hypermyelinated than TSC1 mutants and showed a stronger reduction in g-ratio compared to 3 mpt (Figure 5—figure supplement 2b,f).
In addition to radial hypermyelination, MpzCreERT2:PtenKO nerves exhibited various myelin abnormalities, including infoldings, outfoldings, and tomacula, both at 3 mpt and 6 mpt (Figure 5f,j, Figure 5—figure supplement 2e), consistent with a previous study (Goebbels et al., 2012). In MpzCreERT2:Tsc1KO mice such alterations were slightly, but not significantly, increased, while MpzCreERT2:Tsc1KO:PtenKO nerves contained more myelin abnormalities than single mutants.
Finally, many Remak bundles in PTEN and double mutants displayed redundant SC membrane wrapping around non-myelinated axons, while such features were undetectable in controls or TSC1 knockouts (Figure 5k, Figure 5—figure supplement 3a). These findings are in line with previous reports in PTEN mutants (Goebbels et al., 2010), as is our confirmatory observation of occasional redundant membranes wrapped around collagen fibers in these mice (Figure 5—figure supplement 3b).
Together, our data show that high mTORC1 signaling following deletion of TSC1 and/or PTEN in adult differentiated SCs is capable of reactivating radial myelin growth. This effect is proportional to the levels of mTORC1 hyperactivation. Additionally, the subtly different phenotypes of TSC1 and PTEN mutants, including the presence of abnormal wrapping of Remak fibers in PTEN and double mutants, but not in TSC1 mutants, are consistent with potential mTORC1-independent functions of the PI3K-Akt pathway in membrane wrapping and myelin growth.
In marked contrast to the outcome in early PNS development, hyperactivation of the PI3K-Akt-mTORC1 axis in adulthood promotes myelin growth. These two opposing effects point to a dual role of mTORC1 signaling in normal SC myelination: Inhibition of SC differentiation before the onset of myelination, but promotion of myelin production after myelination has started. This dual role model predicts that the response of SCs to mTORC1 hyperactivation depends on the SC differentiation status. Thus, we performed two additional sets of experiments, in which either SCs in adult nerves with hyperactive mTORC1 were forced to dedifferentiate and redifferentiate, or mTORC1 was hyperactivated in SCs that had just differentiated into myelinating SCs.
First, we induced SC dedifferentiation by carrying out nerve crush injuries in TSC1- or PTEN-conditional knockout backgrounds. Upon nerve injury, SCs dedifferentiate, engage with regrowing axons, and remyelinate them in a highly regulated sequence of events that recapitulates many aspects of developmental myelination (Chen et al., 2007; Scherer et al., 1994; Zorick et al., 1996), but also involves a special repair program (Jessen and Mirsky, 2016). We reasoned that, if our model is valid, inducible deletion of TSC1 or PTEN should lead to impaired remyelination when mice undergo a nerve crush injury. Nerve surgeries were performed in MpzCreERT2:Tsc1KO and MpzCreERT2:PtenKO animals at 2 mpt and nerves were analyzed morphologically at 5, 12, and 30 days post-crush (dpc) (Figure 6a). Quantification of intact-appearing myelin profiles (defined as non-discontinuous and non-collapsed myelin rings) at 5 dpc revealed no major differences in demyelination between mutants and controls (Figure 6b,c) allowing comparative analysis of subsequent remyelination (although a subtle influence of the maintenance phenotypes of MpzCreERT2:Tsc1KO and MpzCreERT2:PtenKO mice cannot be excluded). At 12 dpc, when initial remyelination occurs, both MpzCreERT2:Tsc1KO and MpzCreERT2:PtenKO nerves showed a marked reduction in remyelinated fibers. In TSC1 knockouts, a minor reduction was persistent at 30 dpc, when remyelination initiation in controls was virtually completed (Figure 6b,d). In line with defective remyelination initiation, the percentage of SCs expressing Krox20 was reduced at 12 dpc in MpzCreERT2:PtenKO mice, while the overall proportions of SCs were not changed (Figure 6e–g). Consistently, Oct6 levels were increased in MpzCreERT2:Tsc1KO nerves at the same time point (Figure 6h, Figure 6—figure supplement 1a).
In the second set of experiments, we genetically hyperactivated mTORC1 during developmental myelination, but only after most SCs have started myelinating. According to our model, constitutive activation of mTORC1 in cells that have just differentiated to myelinating SCs should lead to radial hypermyelination. To achieve this goal, we crossed Tsc1-floxed mice with mice carrying a Plp1CreERT2 transgene (yielding Plp1CreERT2:Tsc1KO). We induced recombination by administering 4-hydroxytamoxifen (4OH-TMX) (Zuchero et al., 2015) from P8 to P12 since most SCs have started myelinating at this time (Arroyo et al., 1998), and compared these inducible mutant mice with the previously described DhhCre:Tsc1KO mice, in which recombination occurs before onset of myelination (Figure 6i), together with controls. Plp1CreERT2-driven deletion of TSC1 was successful and led to constitutive mTORC1 activation, as indicated by increased phosphorylation of S6K at T389 (Figure 6j, Figure 6—figure supplement 1b). Consistent with our model, P60 Plp1CreERT2:Tsc1KO nerves were radially hypermyelinated, in contrast to the hypomyelinated DhhCre:Tsc1KO nerves (Figure 6k,l).
Our results reveal that PI3K-Akt-mTORC1 signaling directs PNS myelination at multiple critical levels, including onset of myelination and myelin growth. We started our studies by conditionally deleting TSC1 to explore the effects of hyperactivating mTORC1 in developing SCs. Unexpectedly, the resulting constitutively elevated mTORC1 activity profoundly delayed SC differentiation to myelinating cells. Searching for an explanation, we investigated whether inhibitory feedback of mTORC1 on PI3K-Akt signaling, shown in other cases to account for paradoxical results upon hyperactivation of mTORC1 (Byles et al., 2013; Tang et al., 2014; Yecies et al., 2011), might be responsible. However, the phenotype of TSC1 mutants was even aggravated after restoring Akt activity in SCs via simultaneous PTEN deletion. Moreover, we found that single deletion of PTEN leading to high Akt activity in vivo, or expression of constitutively active Akt in cell cultures, inhibited the onset of myelination per se. Notably as in TSC1 mutants, inhibition of SC differentiation by hyperactive PI3K-Akt due to loss of PTEN was mediated by mTORC1 since rapamycin treatment effectively rescued this effect.
These data unveiled a direct myelination-inhibitory function of the PI3K-Akt-mTORC1 axis. In support of this paradigm, we discovered that mTORC1 negatively regulates Krox20, the main TF required for onset of SC myelination, via S6K. Coherently, we found that mTORC1 activity is physiologically high in not-yet myelinating SCs and declines with their commitment to myelination. In this light, the impaired onset of myelination in TSC1 and/or PTEN mutants can be viewed as a failure of mutant SCs to promptly downregulate mTORC1 activity when myelination has to start. What is then the physiological function of high mTORC1 during early nerve development? We have shown previously that genetic inhibition of mTORC1 impairs radial sorting (Norrmén et al., 2014) and describe now that strong hyperactivation of mTORC1 can accelerate this crucial process in myelination. We infer that high mTORC1 activity during early nerve development may drive radial sorting, concomitant with transiently inhibiting further SC differentiation to myelinating cells, thereby serving as a key component of the regulatory program that coordinates completion of radial sorting and onset of myelination (Feltri et al., 2016).
Intriguingly, recent studies on melanocytes, which share a common developmental origin with SCs, revealed parallels between mTORC1 functions in these cell types. Analogous to the differentiation-inhibiting role of mTORC1 in SCs and the negative regulation of Krox20 by mTORC1, mTORC1 hyperactivation in melanocytes impaired melanogenesis, while inhibition of mTORC1 increased levels of MITF, a TF essential for melanin production (Cao et al., 2017; Ho et al., 2011). Hence, it will be appealing to explore whether an inhibitory role of mTORC1 activity on cell differentiation is a frequent feature of neural crest-derived cells.
Our results show that physiologically high activity of the PI3K-Akt-mTORC1 axis before the ordinary developmental onset of myelination has a myelination-inhibitory function. However, it has been previously reported that conditional ablation of PTEN in the Schwann cell lineage increased the number of myelinated fibers in 3 months-old mice, most likely due to aberrant myelination of normally non-myelinated C fibers (Goebbels et al., 2010). Thus, chronic hyperactivation of PI3K-Akt signaling may be able to, in the long term, drive Remak cells to a myelinating-like phenotype (see also below).
In contrast to the inhibitory effect on SC differentiation, we found that deletion of TSC1 and/or PTEN in adult SCs was able to reactivate myelin growth, proportionally to the levels of mTORC1 activity. However, in spite of comparable mTORC1 hyperactivity, we noticed subtle phenotypical differences between MpzCreERT2:Tsc1KO and MpzCreERT2:PtenKO nerves. First, MpzCreERT2:PtenKO, but not MpzCreERT2:Tsc1KO, displayed increased radial hypermyelination at 6 mpt compared to 3 mpt, indicating that specifically PTEN ablation led to continuous radial myelin growth. Second, in MpzCreERT2:PtenKO and MpzCreERT2:Tsc1KO:PtenKO, but not in MpzCreERT2:Tsc1KO, redundant processes of non-myelinating SCs repeatedly wrapped around axons in Remak bundles, analogous to previous observations (Domènech-Estévez et al., 2016; Goebbels et al., 2010). These findings indicate that the PI3K-Akt pathway is likely to serve also mTORC1-independent functions in controlling SC myelination. In particular, the redundant wrapping of SC membranes upon loss of PTEN, but not of TSC1, prompts us to speculate that, also during physiological myelination, driving of SC membrane wrapping might involve PI3K-Akt-dependent/mTORC1-independent mechanisms. We envisage that the PI3K-Akt pathway serves converging scopes during myelin growth, recruiting (1) mTORC1 to activate the synthesis of myelin building blocks, and (2) mTORC1-independent targets for membrane wrapping. The existence of such a PI3K-Akt-dependent/mTORC1-independent ‘wrapping force’ in myelination is supported by recent reports (Domènech-Estévez et al., 2016; Mathews and Appel, 2016) and may underlie the regulation of the mechanistically required cytoskeletal dynamics. In agreement, recent work demonstrated that constitutively active Akt induces Rac1 activity in SCs (Domènech-Estévez et al., 2016), and remodeling of the actin cytoskeleton is an essential prerequisite for membrane expansion during myelination (Nawaz et al., 2015; Novak et al., 2011; Zuchero et al., 2015).
The formation of myelin abnormalities – a hallmark of numerous neuropathies (Dyck and Thomas, 2005) – may also involve mTORC1-independent targets of the PI3K-Akt pathway, given that such alterations accumulated substantially in MpzCreERT2:PtenKO, but less so in MpzCreERT2:Tsc1KO. Nevertheless, high mTORC1 signaling probably supports their growth, since myelin abnormalities were more abundant in MpzCreERT2:Tsc1KO:PtenKO with higher mTORC1 activity compared to single MpzCreERT2:PtenKO. In accord, treatment with rapamycin has been shown to diminish the load of myelin abnormalities in PTEN mutant mice (Goebbels et al., 2012).
Taken together, also supported by data from others (Beirowski et al., 2017), our results support a model in which the PI3K-Akt-mTORC1 axis fulfills multiple major roles in SC myelination (Figure 7). In early SC development, high mTORC1 signaling maintains SCs in a non-differentiated state by suppressing Krox20 expression and, at the same time, promotes radial sorting. After axonal sorting is completed, a physiological decline in mTORC1 activity releases the suppression on Krox20 expression and allows myelination to start. When this turning point has been passed, the residual mTORC1 activity drives myelin production in concert with mTORC1-independent targets of PI3K-Akt. Downstream of mTORC1, both lipid synthesis via activation of the RXRγ-SREBP1c axis (Norrmén et al., 2014) and protein synthesis (Sheean et al., 2014) are key processes supporting myelin growth.
Our model is especially supported by the opposing outcomes of mTORC1 hyperactivation in developing non-differentiated SCs versus in adult myelinating SCs, revealing that SCs reacted differently depending on their differentiation status. To confirm this notion, we forced adult differentiated SCs to dedifferentiate due to nerve crush injuries, followed by axonal regeneration and SC redifferentiation. As expected in this experimental setting, redifferentiation and remyelination were delayed in MpzCreERT2:Tsc1KO and MpzCreERT2:PtenKO mutants. Conversely, deletion of TSC1 when most SCs had just started myelinating enhanced radial myelin growth.
Our conceptual model raises a number of questions for future investigations including: Are different upstream receptors mediating the differentiation-inhibiting and myelin-growth promoting roles of mTORC1? Is a single upstream receptor system accounting for both, but through different ligands (e.g. neuregulin-1 isoforms), or through different concentrations of the same ligand? What is responsible for the physiological decline in mTORC1 activity that allows SCs to start myelinating? What are the postulated mTORC1-independent targets of PI3K-Akt? We can only speculate in this context, but we have shown here that neuregulin-1 signaling is a major activator of the PI3K-Akt-mTORC1 axis in SCs, in line with previously reported findings of strongly reduced phospho-S6 levels in SCs co-cultured with DRG neurons missing neuregulin-1 (Heller et al., 2014). Neuregulin-1 isoforms are known to exert different effects on SCs in various experimental settings involving several signaling pathways (Mei and Nave, 2014). Thus, signaling through different neuregulin-1 isoforms might contribute to the different functions of mTORC1 in SC biology, potentially in concert with additional signals integrating other adaxonal and also abaxonal cues (Ghidinelli et al., 2017; Heller et al., 2014; Herbert and Monk, 2017; Monk et al., 2015; Pereira et al., 2012).
Modulation of the pathways upstream of mTORC1 has been recently explored as a promising therapeutic approach in animal models of hereditary peripheral neuropathies (Bolino et al., 2016; Fledrich et al., 2014; Goebbels et al., 2012; Nicks et al., 2014). Based on our ‘dual-role’ model, we expect that future studies will benefit from monitoring mTORC1 activity in conjunction with the differentiation status of SCs to enhance efficacy. Moreover, inhibition of mTORC1 should be considered as a valuable therapeutic strategy in hereditary or acquired peripheral neuropathies in which SC differentiation is defective.
Mice harboring floxed alleles of Tsc1 (STOCK Tsc1<tm1Djk>/J, RRID:IMSR_JAX:005680) and Pten (C;129S4-Pten<tm1Hwu>/J, RRID:IMSR_JAX:004597) were obtained from The Jackson Laboratory. Mice harboring floxed alleles of Rptor (Bentzinger et al., 2008; Polak et al., 2008) and mice carrying a Cre transgene under control of the Dhh (RRID:IMSR_JAX:012929) or Mpz promoter (RRID:IMSR_JAX:017927), or a CreERT2 transgene under control of the Plp1 or Mpz promoters have been described (Feltri et al., 1999; Jaegle et al., 2003; Leone et al., 2003). To generate non-inducible conditional deletion of TSC1, PTEN, or Raptor, floxed mice were crossed with DhhCre- or MpzCre-positive mice. To generate inducible conditional deletion of TSC1 or PTEN, floxed mice were crossed with MpzCreERT2-positive mice, and at 8–10 weeks of age 2 mg of tamoxifen (Sigma-Aldrich, Saint Louis, MO, USA) in 10% ethanol/sunflower seed oil (Sigma-Aldrich) were injected intraperitoneally once a day on five consecutive days in both mutant and control animals. Developmental inducible deletion of TSC1 was achieved crossing floxed mice with Plp1CreERT2-positive mice and administering intraperitoneally to both mutant and control pups 75 μg per gram of body weight of 4-hydroxytamoxifen (Sigma-Aldrich) dissolved in Kolliphor EL (Sigma-Aldrich) once a day on five consecutive days from P8 to P12. Experimental animals were on a hybrid background between C57B6 and the background of origin of the floxed mice (2–4 backcrosses with C57B6 for all TSC1, PTEN, or TSC1/PTEN mutants and their controls; 7–8 backcrosses with C57B6 for Raptor mutants and their controls). Cre-negative animals (floxed homozygous or heterozygous) were used as controls in the experiments. To control for variations in background, littermate controls were used. Wild-type mice were on a C57B6 background. Mice of either sex were used in the experiments. For in vivo rescues, rapamycin (Millipore, Billerica, MA, USA) was dissolved in a solution of 5% PEG-400, 5% Tween-80, and 4% ethanol, and 5 μg rapamycin per gram of body weight were administered intraperitoneally once a day at P3 and P4. Genotypes were determined through genomic PCR using the following primers: Cre forward 5’-accaggttcgttcactcatgg-3’, reverse 5’-aggctaagtgcctcttctaca-3’; TSC1 forward 5’-gtcacgaccgtaggagaagc-3’, reverse 5’-gaatcaaccccacagagcat-3’; PTEN forward 5’-caagcactctgcgaactgag-3’, reverse 5’-aagtttttgaaggcaagatgc-3’, Raptor forward 5’-atggtagcaggcacactcttcatg-3’, reverse 5’-gctaaacattcagtccctaatc-3’. Mice were housed with a maximum number of five animals per cage, kept in a 12 hr light-dark cycle, and fed standard chow ad libitum.
Mice were subjected to unilateral sciatic nerve crush injury two months after tamoxifen administration. After inducing anesthesia with isofluorane inhalation, the sciatic nerve was exposed through blunt dissection of the thigh muscles and compressed for 30 s. For analgesia 0.1 mg kg−1 buprenorphine (Temgesic, Reckitt Benckiser, UK) were injected intraperitoneally once prior to the surgery, and thereafter every 12 hr for two days. Morphological analysis and immunostaining of crushed nerves were performed on sections 3 mm distal to the injury site. For biochemical analysis, the whole nerve distal to the injury site was used.
Immediately after dissection, sciatic nerves were fixed with 3% glutaraldehyde and 4% paraformaldehyde in 0.1 M phosphate buffer. Sciatic nerves were further treated with 2% osmium tetroxide (EMS, Hatfield, PA, USA), dehydrated over a series of acetone gradients and embedded in Spurrs resin (EMS). Semithin sections (650 nm) were stained with 1% toluidine blue and used for qualitative analysis and for quantifying intact myelin profiles after nerve crush. Ultrathin sections (65 nm) were imaged with a FEI Morgagni 268 TEM, and random 5 × 5 MIA (multiple image alignment) fields were acquired for qualitative analysis and for quantifying the percentage of myelinating SCs. G-ratio measurements and quantification of myelin abnormalities were performed on EM reconstructions of the entire sciatic nerve, obtained from additional sections (99 nm) collected on ITO coverslips (Optics Balzers, Germany) and imaged with either a Zeiss Gemini Leo 1530 FEG or Zeiss Merlin FEG scanning electron microscopes attached to ATLAS modules (Zeiss, Germany). To calculate the g-ratio, the axon diameter was derived from the axon area, while the fiber diameter was calculated adding to the axon diameter twice the average of the myelin thickness measured at two different locations of the myelin ring. At least 200 fibers per nerve were analyzed.
The following primary antibodies were used: TSC1 (CST, #6935, 1:1,000), phospho-S6KT389 (Cell Signaling Technology Cat# 9234 also 9234L, 9234P, 9234S RRID:AB_2269803, 1:1,000), S6K (Cell Signaling Technology Cat# 9202S RRID:AB_331676, 1:1,000), phospho-4EBP1T37/46 (Cell Signaling Technology Cat# 2855 also 2855L, 2855S, 2855P RRID:AB_560835, 1:1,000), 4EBP1 (Cell Signaling Technology Cat# 9452S RRID:AB_10693791, 1:1,000), phospho-S6S235/236 (Cell Signaling Technology Cat# 4857S RRID:AB_2181035, 1:1,000), S6 (Cell Signaling Technology Cat# 2317S RRID:AB_10694551, 1:1,000), cJun (Cell Signaling Technology Cat# 9165 RRID:AB_2130165, 1:1,000), phospho-ErbB2Y1248 (Cell Signaling Technology Cat# 2247S RRID:AB_331725, 1:1,000), ErbB2 (Cell Signaling Technology Cat# 2165S RRID:AB_10692490, 1:1,000), phospho-AktT308 (Cell Signaling Technology Cat# 9275 also 9275S, 9275L RRID:AB_329828 only for the western blot in Figure 4a, Cell Signaling Technology Cat# 4056 also 4056S, 4056L RRID:AB_331163 for the other western blots, both used 1:1,000), phospho-AktS473 (Cell Signaling Technology Cat# 4051 also 4051S, 4051L RRID:AB_331158, 1:1,000), Akt (Cell Signaling Technology Cat# 9272 also 9272S RRID:AB_329827, 1:1,000), phospho-ERK1/2 T202/Y204 (Cell Signaling Technology Cat# 9106 also 9106L, 9106S RRID:AB_331768, 1:1,000), ERK1/2 (Cell Signaling Technology Cat# 9102 also 9102L, 9102S RRID:AB_330744, 1:1,000), phospho-PRAS40T246 (CST, #13175, 1:1,000), PRAS40 (Cell Signaling Technology Cat# 2691S RRID:AB_2225033, 1:1,000), PTEN (Cell Signaling Technology Cat# 9559S RRID:AB_10695541, 1:1,000), Akt phospho-substrates (Cell Signaling Technology Cat# 10001S RRID:AB_10950819, 1:1,000), α-tubulin (Sigma-Aldrich Cat# T5168 RRID:AB_477579, 1:1,000), β-actin (Sigma-Aldrich Cat# A5316 RRID:AB_476743, 1:1,000), GAPDH (Hytest Cat# 5G4-9B3 RRID:AB_1616725, 1:10,000), NF (Sigma-Aldrich Cat# N5264 RRID:AB_477278, used for immunocytochemistry, 1:200), NF (Aves Labs Cat# NF-M RRID:AB_2313554, used for immunohistochemistry, 1:1,000), MBP (Bio-Rad/AbD Serotec Cat# MCA409S RRID:AB_325004, 1:200), Sox10 (R&D Systems, AF2864, 1:200), S100 (Dako Cat# Z0311 RRID:AB_10013383, 1:200), laminin-α2 (Sigma-Aldrich Cat# L0663 RRID:AB_477153, 1:200). Primary antibodies against Oct6 and Krox20 (both used 1:1,000) were generous gifts from Dr. Dies Mejer. HRP-, AP-, and fluorophore-conjugated secondary antibodies were purchased from Jackson ImmunoResearch and used 1:200 for immunostainings or 1:10,000 for western blots. Alexa488-coupled phalloidin (used 1:100) was from Life Technologies (Carlsbad, CA, USA) (RRID:AB_2315147).
The following chemical inhibitors were used in in vitro experiments: rapamycin (Sigma-Aldrich), LY294002 (Sigma-Aldrich), CI-1040 (Sigma-Aldrich), torin-1 (Tocris, UK), LYS6K2 (Focus Biomolecules, Plymouth Meeting, PA, USA).
Immediately after dissection, sciatic nerves were placed in ice-cold PBS, the epineurium was removed with fine forceps, and the nerves were snap-frozen in liquid nitrogen and stored at −80°C until further processing. To prepare lysates, nerves were ground on dry ice, mixed with PN2 lysis buffer (25 mM Tris-HCl pH 7.4, 95 mM NaCl, 10 mM EDTA, 2% SDS, protease and phosphatase inhibitors (Roche, Switzerland)), boiled, and spun for 15 min. 10–30 μg of proteins per sample were mixed 1:4 with sample buffer (200 mM Tris-HCl pH6.8, 40% glycerol, 8% SDS, 20% β-mercaptoethanol, 0.4% bromophenol blue), run on 4–15% polyacrylamide gradient gels (Biorad, Hercules, CA, USA), and blotted onto PVDF membranes (Millipore). After blocking with 5% milk in TBS-T, membranes were incubated overnight with primary antibodies diluted in 5% BSA in TBS-T. Chemiluminescent signals were generated using HRP- or AP-conjugated secondary antibodies and ECL (GE Healthcare), ECL prime (GE Healthcare), or CDP-Star (Roche), and detected using Fusion FX7 (Vilber Lourmat, Germany). If required, after signal detection membranes were stripped with a buffer containing β-mercaptoethanol (1.52 g Tris, 2 g SDS, 700 μl β-mercaptoethanol in 100 ml water, pH 6.8) for 15 min at 55°C and reprobed with antibodies. Quantification of band intensities was performed with ImageJ (version 1.50i). α-tubulin, β-actin, or GAPDH were used as normalization controls. For cropped blots, the control bands are displayed immediately below the other bands that belong to the same membrane. Full-length blots are shown in Supplementary files 1–8. Size markers refer to All Blue Precision Protein Standards (Biorad). Protein size is expressed as apparent molecular weight in kDa.
Immediately after dissection, sciatic nerves were fixed for one hour in 4% paraformaldehyde, cryopreserved in 10% sucrose for one hour, 20% sucrose overnight, and then embedded in OCT (TissueTek, Torrance, CA, USA). 8 μm thick cryosections were cut, blocked with blocking buffer (1% BSA, 10% goat or donkey serum, 0.1% triton X-100 in PBS) for one hour, incubated overnight with primary antibodies, incubated for one hour with fluorophore-conjugated secondary antibodies, counterstained with DAPI (Life Technologies), and mounted with Vectashield (Vector Laboratories, Burlingame, CA, USA). The same procedure was used for cell culture stainings. For cell proliferation experiments, mouse pups were injected with 50 μg EdU (Life Technologies) per gram of body weight and sacrificed one hour later. EdU staining was performed using the Click-iT EdU Alexa647 kit (Life Technologies) as per manufacturer’s instructions. Immunostainings were imaged using an epifluorescence microscope (Zeiss Axio Imager.M2) equipped with a monochromatic CCD camera (sCMOS, pco.edge), or with a confocal microscope (Leica TCS SP8). One representative section per sample was imaged and analyzed.
Immediately after dissection, sciatic nerves were placed in ice-cold PBS, the epineurium was removed with fine forceps, and the nerves were snap-frozen in liquid nitrogen and stored at −80°C until needed. Total RNA was extracted using RNeasy kit (Qiagen, Germany) for developmental samples or Qiazol (Qiagen) for adult samples and cells as per manufacturer’s instructions. 50–200 ng of total RNA were reverse transcribed using Maxima First Strand cDNA Synthesis Kit (Thermo Fisher Scientific, Waltham, MA, USA) as per manufacturer’s instructions. qPCR reactions were performed using FastStart Essential DNA Green Master (Roche) and Light Cycler 480 II (Roche). The sequences of the primers used and their specificity are the following: Krox20 (mouse and rat) forward 5’-acagcctctacccggtggaagac-3’, reverse 5’-cagagatgggagcgaagctactcggata-3’; cJun (mouse and rat) forward 5’-gccaagaactcggaccttctcacgtc-3’, reverse 5’-tgatgtgcccattgctggactggatg-3’; Oct6 (mouse) forward 5’-gagcactcggacgaggatg-3’, reverse 5’-cacgttaccgtagagggtgc-3’; Brn2 (mouse) forward 5’-tcaaatgccctaagccctcg-3’, reverse 5’-cgggaggggtcatccttttc-3’; Sox10 (mouse) forward 5’-ccgaccagtaccctcacct-3’, reverse 5’-tcaatgaaggggcgcttgt-3’; Sox2 (mouse) forward 5’-ggaaagggttcttgctgggt-3’, reverse 5’-acgaaaacggtcttgccagt-3’; Id2 (mouse) forward 5’-catcagcatcctgtccttgc-3’, reverse 5’-ttctcctggtgaaatggctgat-3’; GAPDH (mouse and rat) forward 5’-ggtgaaggtcggtgtgaacggatttgg-3’, reverse 5’-ggtcaatgaaggggtcgttgatggcaac-3’; β-actin (mouse) forward 5’-gtccacacccgccacc-3’, reverse 5’-ggcctcgtcacccacatag-3’; α-tubulin (mouse) forward 5’-tcttagttgtcgggaacggt-3’, reverse 5’-ggagatgcactcacgcatgata-3’. Relative mRNA fold changes for each gene were obtained by using the 2-ΔΔCt method after normalization to GAPDH, β-actin, or α-tubulin.
To generate lentiviral vectors for overexpression in SCs, the hPGK promoter of the pCCLsin.PPT.hPGK.PRE lentiviral backbone was replaced by a 1.1 kb fragment of the rat P0 promoter, as previously described (Norrmén et al., 2014). myrAkt constructs were obtained from Addgene (#9008, #9016, #9017) (Ramaswamy et al., 1999), PCR-amplified, and inserted between the AgeI and SalI restriction sites of the modified pCCLsin.PPT.hPGK.PRE vector. The 4EBP1-4xA construct was obtained from Addgene (#38240) (Thoreen et al., 2012) and subcloned between the BamHI and NheI restriction sites of the pcDNA3.1 vector. As control, the eGFP coding sequence from the pCCLsin.PPT.hPGK.PRE vector was subcloned between the NheI and EcoRI restriction sites of the pcDNA3.1 vector. All constructs were sequence-verified prior to usage.
To prepare embryonic SCs, E13.5 mouse DRGs were isolated, digested with trypsin-EDTA 0.25% (Life Technologies) for 30 min, resuspended in N2-medium (Advanced DMEM:F12 plus N2 supplement (Life Technologies)) with 50 ng ml−1 7S NGF (Millipore), and plated on uncoated dishes. After one week, the SC-neuron network was mechanically detached from the underlying fibroblast layer, digested with 0.25% trypsin (Sigma-Aldrich, T9201) and 0.1% collagenase (Sigma-Aldrich, C0130), and replated on PLL-coated dishes in N2-medium devoid of NGF. After reaching confluency, cells were trypsinized, centrifuged, and resuspended in flow buffer (PBS plus 2% FCS (Life Technologies)). Flow cytometry experiments were performed with LSR Fortessa (BD Biosciences, Franklin Lakes, NJ, USA) using between 105 and 106 cells per genotype, and the data were analyzed with the FlowJo software (RRID:SCR_008520, version 10.0.7). To account for differences in the number of recorded events for each sample, the flow cytometry profiles were normalized to their modes.
To prepare neonatal mouse SCs, sciatic nerves were dissected from P2 mouse pups, the epineurium was removed, and the nerves were digested with 1.25 mg ml−1 trypsin (Sigma-Aldrich, T9201) and 2 mg ml−1 collagenase (Sigma-Aldrich, C0130) for one hour. Cells were then centrifuged, resuspended in D-medium (DMEM-Glutamax plus 10% FCS (Life Technologies)), and seeded on PLL-coated 12 mm glass coverslips. After overnight incubation, cells were fixed with 4% paraformaldehyde and stained.
To prepare rat SCs, sciatic nerves were dissected from P2 Sprague-Dawley rat pups, the epineurium was removed, and nerves were digested with 1.25 mg ml−1 trypsin (Sigma-Aldrich, T9201) and 2 mg ml−1 collagenase (Sigma-Aldrich, C0130) for one hour. After centrifugation, the cells were resuspended in D-medium (DMEM-Glutamax plus 10% FCS (Life Technologies)) and plated on PLL-coated dishes. Contaminating fibroblasts were eliminated with 10 μM Ara-C treatment (Sigma-Aldrich) for 48 hr, followed by complement-mediated removal of fibroblasts using an anti-Thy1.1 antibody (Bio-Rad/AbD Serotec Cat# MCA04G RRID:AB_322809, 1:50). After a 10 min incubation with the anti-Thy1.1 antibody, rabbit complement (Millipore) was added for 40 min. SCs were then further expanded in SC growth medium (DMEM-Glutamax (Life Technologies), 10% FCS (Life Technologies), 5 μg ml−1 bovine pituitary extract (Life Technologies), 2 μM forskolin (Sigma-Aldrich)), tested for mycoplasma contamination, aliquoted, and stored in liquid nitrogen. For differentiation experiments, SCs were transferred to defined medium (100 μg ml−1 of holotransferrin (Sigma-Aldrich), 60 ng ml−1 progesterone (Sigma-Aldrich), 1 μg ml−1 insulin (Sigma-Aldrich), 16 μg ml−1 putrescine (Sigma-Aldrich), 400 ng ml−1 L-thyroxin (Sigma-Aldrich), 160 ng ml−1 sodium selenite (Sigma-Aldrich), 10 ng ml−1 triiodothyronine (Sigma-Aldrich), 38 ng ml−1 dexamethasone (Sigma-Aldrich), and 300 μg ml−1 BSA (Sigma-Aldrich) in DMEM:F12-Glutamax (Life Technologies)), and differentiation was induced with 1 mM dbcAMP (Sigma-Aldrich) for 24 hr. For experiments with neuregulin stimulation, SCs were serum-starved for 6.5–8 hr in DMEM-Glutamax with 0.05% FCS (Life Technologies) and subsequently treated with 10 ng ml−1 recombinant human EGF domain of neuregulin-1 β1 (R&D Systems, Minneapolis, MN, USA). For all other experiments, SCs were kept in SC growth medium. Transfection of rat SCs was performed using Lipofectamine 3000 (Life Technologies) as per manufacturer’s instructions. Only SCs lower than passage five were used for experiments.
DRGs were isolated from E13.5 embryos, digested for 45 min with trypsin-EDTA 0.25% (Life Technologies), and seeded on 12 mm coverslips coated with 10% Matrigel (Corning, NY, USA) at a density of 1.2 DRGs per coverslip. DRGs from embryos with the same genotype were pooled before seeding. Cultures were kept in supplemented NB-medium (Neurobasal (Life Technologies), B27 supplement (Life Technologies), 4 g l−1 D-glucose (Sigma-Aldrich), 2 mM L-glutamine (Life Technologies), 50 ng ml−1 7S NGF (Millipore)) for the first five days and then in C-medium (MEM-Glutamax (Life Technologies), 10% FCS (Life Technologies), 4 g l−1 D-glucose (Sigma-Aldrich), 50 ng ml−1 7S NGF (Millipore)) supplemented with 50 μg ml−1 ascorbic acid (Sigma-Aldrich) to induce myelination for an additional 8–10 days. Both supplemented NB- and C-medium were changed every other day. Before staining, DRG-cultures were fixed in 4% paraformaldehyde for 15 min and subsequently permeabilized with ice-cold methanol for 10 min. Images of whole coverslips were acquired using an epifluorescence microscope (Zeiss Axio Imager.M2) equipped with a monochromatic CCD camera (sCMOS, pco.edge) and an automated stage. Between 3 and 6 coverslips per condition were imaged and analyzed. To quantify the extent of myelination, four random fields per coverslip were selected, the MBP-positive and the NF-positive areas per field were measured with ImageJ (version 1.50i) after thresholding, and the MBP area/NF area ratio per coverslip was calculated as average of the MBP area/NF area ratios of the different fields. The same threshold was applied to all samples and conditions.
HEK293T cells were from ATCC (Manassas, VA, USA), were not further authenticated, and were regularly monitored to assure lack of mycoplasma contamination. Subconfluent HEK293T cells were transfected in 10 cm dishes with lentiviral vectors and the packaging plasmids psPAX2 and pCMV-VSV-G using Lipofectamine 2000 as per manufacturer’s instructions (Life Technologies). Non-concentrated viruses were collected 48 hr after transfection, aliquoted, and stored at −80°C until needed. For infection of DRG-explant cultures, non-concentrated viruses were mixed 1:1 with supplemented NB-medium and added to the cultures for 24 hr from DIV 3 to 4.
The quantity and quality of isolated RNA was determined with a Qubit (1.0) Fluorometer (Life Technologies) and a Bioanalyzer 2100 (Agilent, Santa Clara, CA, USA). The TruSeq Stranded mRNA Sample Prep Kit (Illumina, San Diego, CA, USA) was used in the succeeding steps. Briefly, total RNA samples (150 ng) were ribosome depleted and then reverse-transcribed into double-stranded cDNA with actinomycin added during first-strand synthesis. The cDNA samples were fragmented, end-repaired and polyadenylated before ligation of TruSeq adapters. The adapters contain the index for multiplexing. Fragments containing TruSeq adapters on both ends were selectively enriched with PCR. The quality and quantity of the enriched libraries were validated using Qubit (1.0) Fluorometer and the Bioanalyzer 2100 (Agilent). The product is a smear with an average fragment size of approximately 360 bp. The libraries were normalized to 10 nM in Tris-Cl 10 mM, pH 8.5 with 0.1% Tween-20. The TruSeq SR Cluster Kit v4-cBot-HS or TruSeq PE Cluster Kit v4-cBot-HS (Illumina) was used for cluster generation using 8 pM of pooled normalized libraries on the cBOT. Sequencing was performed on the Illumina HiSeq 2500 paired-end at 2 × 126 bp or single-end 126 bp using the TruSeq SBS Kit v4-HS (Illumina).
The raw reads were first cleaned by removing adapter sequences, trimming low quality ends, and filtering reads with low quality (phred quality <20) using Trimmomatic (Bolger et al., 2014). Sequence alignment of the resulting high-quality reads to the Mus musculus reference genome (build GRCm38) and quantification of gene level expression was carried out using RSEM (version 1.2.22) (Li and Dewey, 2011). To detect differentially expressed genes we applied count based negative binomial model implemented in the software package EdgeR (R version: 3.2.2, edgeR_3.12.0) (Robinson et al., 2010). The differential expression was assessed using an exact test adapted for over-dispersed data. Genes showing altered expression (fold change >1.2) with adjusted (Benjamini and Hochberg method) p-value<0.05 (indicated as false discovery rate, FDR) were considered differentially expressed. Within this set of genes, downregulated and upregulated genes were separately subjected to gene ontology analysis of biological processes using the online tool Enrichr (http://amp.pharm.mssm.edu/Enrichr/).
Data processing and statistical analyses were performed using GraphPad Prism (RRID:SCR_002798, version 7.0a) and Microsoft Excel (version 15.27). Data distribution was assumed to be normal and variances were assumed to be equal, although this was not formally tested due to low n number. Sample sizes were chosen according to sample sizes generally employed in the field. The investigators were blinded to the genotypes during analysis of morphological and immunohistochemical data, except for those cases in which mutant mice exhibited an obvious phenotype. No randomization methods were used. Two-tailed unpaired Student’s t-test was used if only two conditions or genotypes were compared. In all other cases, one- or two-way ANOVAs followed by Tukey’s, Dunnett’s, or Sidak’s multiple comparisons tests were employed, as indicated in the figure legends. p<0.05 was considered to be statistically significant. No samples or data were omitted during the analyses.
RNA-sequencing data have been deposited in the ENA database under accession number PRJEB20661.
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Beth StevensReviewing Editor; Boston Children's Hospital, Harvard Medical School, United States
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 "Dual Function of the PI3K-Akt-mTORC1 Axis in Myelination of the Peripheral Nervous System" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Gary Westbrook as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Robert H Miller (Reviewer #1); Wendy B. Macklin (Reviewer #2). 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 of reviews and reviewer/editor discussion:
The reviewers find that this is a compelling paper that investigates the functional roles of mTOR1 activation in Schwann cell myelination during different stages of development, homeostasis, and in the context of remyelination after injury. Using multiple genetic manipulations to disrupt key inhibitory components of mTORC1 (TSC1 or PTEN), the authors convincingly demonstrate, that hyperactivation of mTORC during early development blocks myelination through inhibition of Schwann cell differentiation and generation of radial sorting. By contrast, hyperactivation of mTORC in the adult animal after peripheral nerve crush results in hypermyelination. The studies also provide evidence that high mTORC1 activity inhibits Krox20 expression, a known master regulator of Schwann cell myelination, and demonstrate Krox20 as an effector of mTORC signaling in PNS myelination that is mediated by SK6.
Strengths of the study are the careful use of multiple genetic approaches to delineate the mechanisms by which mTORC modulates Schwann cell development and myelination in different contexts. The study also helps explain conflicting results raised by the series of earlier studies and defines a model that is important for future investigations on Schwann cell differentiation and remyelination. The results are well communicated, and data is of high quality and supports the interpretations. However there were a few points that require clarification and expanded discussion, as outlined below. (The full reviews are also appended below).
1) P0-PTEN-KO mice have fewer myelinated axons at P5, and Dhh-PTEN cKO mice are comparable, indicating that at either stage of differentiation, PTEN cKO/Akt overactivation induces decreased numbers of myelinated axons. However CNP-PTEN cKO mice have significantly more myelinated axons at 3 M (Goebbels et al. J Neurosci 2010), and that promoter should have deleted PTEN in this same time frame of Schwann cell differentiation. Some discussion of this is needed.
2) In the nerve injury experiments, according to their own data, the mice would have had increased myelin, altered Remak bundles and increased numbers of myelinated axons before the nerve injury. How does this impact the eventual myelin loss/recovery after nerve injury? How do they define "intact appearing myelin profiles" (Figure 6) in the environment of altered myelin?
This is an excellent manuscript that clearly defines a complex phenomena in the mTORC modulation of PNS development. The authors have been careful to rule out other potential signaling paths and have, in many cases, provided multiple studies to support their interpretation. The manuscript represents one of the most detailed and careful studies of the role of mTORC signaling in peripheral myelination to date. There are few concerns with the data which is of high quality and supports the interpretation of the authors.
Overall, this is a very comprehensive and extremely well done manuscript that will be of interest to a wide range of developmental neurobiologists.
This study by Figlia et al. focuses on an important research problem that has had significant, apparently contradictory data for a number of years. It has been clearly shown that mTORC1 activity is required for myelination. Yet, Dr. Suter's group and others have shown that deleting TSC1 from myelinating cells, which clearly upregulates mTORC1 activity since TSC1 normally inhibits mTORC1 activity, actually results in hypomyelination. The current manuscript is a detailed, complex analysis of why these unexpected data have been obtained by multiple investigators. This study is important and thorough.
This study demonstrates yet again that deleting TSC in Dhh-expressing Schwann cells, i.e., in early Schwann cell lineage cells, results in increased mTORC1 activity yet hypomyelination, and that rapamycin treatment induces partial recovery of myelination. They then demonstrate that deletion of PTEN, which increases mTORC1 activity by way of activated Akt signaling, in either Dhh or P0-expressing cells results in a partial reduction in myelination. The double TSC/PTEN conditional knockout eliminates essentially all myelination. Rapamycin can recover some of this loss of myelination. Thus, these data are convincing and consistent with some of the data in the literature that increasing mTOR signaling by deletion of either endogenous inhibitor can reduce myelination in Schwann cells.
The studies also show convincingly that high mTORC1 activity likely inhibits Krox20 expression, consistent with its apparent inhibition of initial Schwann cell differentiation into myelinating Schwann cells.
They demonstrate, as has Goebbels et al., 2010, that deleting PTEN and other mTORC1 activators in adults induces hypermyelination. This is a far more in-depth analysis of sciatic nerve hypermyelination than in the Goebbels et al. study, and they go on to study remyelination after injury in these mTORC1-activated mice. Importantly, they show that, as in development, remyelination is impaired in mice with excess mTORC1 signaling in remyelinating Schwann cells.
This study is important as it answers many of the questions raised by the series of earlier studies, and defines a model that is important for future investigations on Schwann cell differentiation.
Nevertheless, there are some important concerns:
1) P0-PTEN-KO mice have fewer myelinated axons at P5, and Dhh-PTEN cKO mice are comparable, indicating that at either stage of differentiation, PTEN cKO/Akt overactivation induces decreased numbers of myelinated axons. However CNP-PTEN cKO mice have significantly more myelinated axons at 3 M (Goebbels et al. J Neurosci 2010), and that promoter should have deleted PTEN in this same time frame of Schwann cell differentiation. Some discussion of this is needed.
2) In the nerve injury experiments, according to their own data, the mice would have had increased myelin, altered Remak bundles and increased numbers of myelinated axons before the nerve injury. How does this impact the eventual myelin loss/recovery after nerve injury? How do they define "intact appearing myelin profiles" (Figure 6) in the environment of altered myelin?https://doi.org/10.7554/eLife.29241.034
- Ueli Suter
- Camilla Norrmén
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank all members of the Suter lab, especially Dr. Deniz Gökbuget, for discussions, Drs. Monica Ghidinelli and Ned Mantei for critically reading the manuscript, and Joanne Jeker, Francesco Santarella, the Functional Genomic Center Zürich (FGCZ), and the ScopeM imaging facility of ETH Zürich for excellent technical support. We also thank Dr. Dies Meijer (University of Edinburgh) for DhhCre mice and antibodies, Drs. Laura Feltri and Lawrence Wrabetz (University of Buffalo) for MpzCre mice, and Drs. Michael Hall and Markus Rüegg (University of Basel) for floxed-Rptor mice.
Animal experimentation: Mice were kept at 12 hr light/dark cycle. All experiments were approved by the Veterinary Office of the Canton of Zurich, Switzerland (reference No. ZH161/2014, ZH44/2013)
- Beth Stevens, Reviewing Editor, Boston Children's Hospital, Harvard Medical School, United States
- Received: June 2, 2017
- Accepted: July 31, 2017
- Version of Record published: September 7, 2017 (version 1)
© 2017, Figlia et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.