https://orcid.org/0000-0001-8760-7865
Abstract
Several mosaic mutations of the mammalian/mechanistic target of rapamycin (mTOR) have recently been found in patients with cortical malformations, such as hemimegalencephaly (HME) and focal cortical dysplasia (FCD). Although all of them should activate mTOR signaling, comparisons of the impact of different mTOR mutations on brain development have been lacking. Also it remains unknown if any potential differences these mutations may have on cortical development are directly related to a degree of mTOR signaling increase. The present study assessed levels of mTORC1 pathway activity in cell lines and rat primary neurons overexpressing several mTOR mutants that were previously found in HME, FCD, cancer patients and in vitro mutagenesis screens. Next we introduced the mutants, enhancing mTORC1 signaling most potently, into developing mouse brains and assessed electroporated cell morphology and migratory phenotype using immunofluorescent staining. We observed the differential inhibition of neuronal progenitor cortical migration, which partly corresponded with a degree of mTORC1 signaling enhancement these mutants induced in cultured cells. The most potent quadruple mutant prevented most of the progenitors from entering the cortical plate. Cells that expressed less potent, single-point, mTOR mutants entered the cortical plate but failed to reach its upper layers and had enlarged soma. Our findings suggest a correlation between the potency of mTOR mutation to activate mTORC1 pathway and disruption of cortical migration.
Introduction
During development, the cerebral cortex arises from the most rostral vesicle of the neural tube (i.e. the telencephalon) in a process that is referred to as corticogenesis. In mice (i.e. the most studied model organism), corticogenesis occurs from embryonic day 10.5 (E10.5) post coitum until the early postnatal period. Most cortical neurons are excitatory and produced in the dorsal telencephalon in the lateral ventricle-adjacent ventricular zone (VZ)/subventricular zone (SVZ). At the onset of corticogenesis, neuroepithelial cells give rise to apical radial glia cells (RGcs) that generate neurons, initially directly through asymmetric divisions and later also indirectly through intermediate progenitors (IPs). RGcs also guide neurons with the aid of basal processes that contact pia to their destination in the cortex during radial cortical migration.
Excitatory neurons are produced in successive waves. They migrate toward the pial surface until they encounter Cajal–Retzius (CR) cells. These cells originate from tangential migration during early neural development and form layer I of the cortex, also called the marginal zone (MZ; (1)). Excitatory neurons first translocate from the neurogenic niche directly to the adjacent subplate (SP). All later-born neurons must pass beyond their predecessors to form five cortex layers (II–VI, numbered from top to bottom) in an inside-out manner. Early-born, SP and layer VI neurons rely solely on somal translocation to integrate into deep layers (DLs). Later-born neurons first adopt a multipolar morphology and immediately switch to a bipolar morphology and migrate to either layer V or the upper layers (ULs). At the ULs, upon contact of their leading process with CR cells, the neurons detach from RGcs basal processes and translocate their soma directly under the MZ (1).
Disturbances in neural stem cell proliferation and migration or differentiation of their progeny can lead to a wide group of malformations of cortical development (MCDs). These MCDs include, among others, focal cortical dysplasia (FCD), hemimegalencephaly (HME) and tuberous sclerosis complex (TSC). MCDs are commonly associated with drug-resistant epilepsy, cognitive impairment, behavioral disorders and autism spectrum disorders. FCD is the most common and heterogeneous MCD, which can be divided into three subtypes (I, II and III). Type II FCDs (IIa and IIb) are characterized by the presence of atypical cells. Type IIa is characterized by dysmorphic/cytomegalic neurons. Type IIb is characterized by the dysmorphic/cytomegalic neurons and also balloon cells (2). Similar cellular alterations are also found in TSC, an autosomal-dominant genetic disorder that is characterized by the presence of cortical tubers, and in HME. HME is a broader MCD that usually encompasses less than half but sometimes even the entire cerebral hemisphere. It also presents with atypical cells, gyrification disturbances, a blurred gray/white matter boundary and the enlargement of deep gray matter structures and ventricles.
Mammalian/mechanistic target of rapamycin (mTOR) is a serine/threonine kinase that is present in the cell in two different multiprotein complexes. mTOR complex 1 (mTORC1) receives signals of nutrient and growth factor availability and regulates cell proliferation and growth by controlling various cellular process, among which translation plays a prominent role (3). Studies of TSC knockout mice showed that mTOR hyperactivation results in neurodevelopmental abnormalities, including brain hypertrophy, heterotopia, an increase in cortical thickness, lamination defects, enlarged ectopic cells and dysplastic neurons (2,4). The transgenic overexpression of an mTOR quadruple hyperactive mutant (SL1+IT) during corticogenesis led to the improper positioning of neurons in the cortex and a greater neuronal soma size (5). De novo mutations in mTOR have also been described in FCD and HME (3,6,7). However, only in the case of a few of them (e.g. L2427P, A1459D and C1483Y), both the excessive activity of the mTORC1 pathway and disturbances in the migration of neurons in the developing cerebral cortex have been demonstrated (8–10). Other FCD and HME-related MTOR mutations as well as numerous gain-of-function mTOR mutants that have been found in mutagenesis screens in yeast (11–13) and extracted from cancer mutation databases (e.g. Catalogue of somatic mutations in cancer, COSMIC [https://cancer.sanger.ac.uk/cosmic]; 14,15), are not yet characterized in this respect so far. Since there has not been a parallel comparison of the effects of overproduction of mTOR mutants in animal models, it is also not clear whether the various activating mutations identified in MTOR will have a different effect on brain development. If yes, another question is rising whether these differences result from the level of mTORC1 activation. Such a possibility was suggested (16), based on clinical cases analysis. In our work, in order to answer the above questions, we comparatively investigated mTOR mutant potential to activate mTORC1 pathway, to next perform, using in utero electroporation, thorough side-by-side comparisons of the effects of most potent mTOR mutants on morphology, identity marker expression and migration of newly born neurons at early stages of cortical neurodevelopment.
Results
Hyperactive mTOR mutant overexpression in cell lines and primary neurons in vitro increases mTORC1 signaling
To choose the most suitable mTOR mutations as a basis for stem cell and animal models of mTORC1 hyperactivation, we browsed the literature for yeast and human mutagenesis screens, patient sequencing efforts and cancer mutation atlases (COSMIC). We screened for mutations that were evolutionarily conserved across eukaryotes and that were preferentially indicated to lead to mTOR kinase hyperactivation. For our initial assays, we chose a total of 12 point mutations that resided within the FAT (L1460P and C1483Y) or kinase (I2017Y, V2198A, S2215Y, L2216H, L2260P, V2403F, E2419K, L2427T and L2431H) domains of mTOR, one triple mutant (V2198A, L2216H and L2260P) and one quadruple mutant (triple plus I2017Y), previously referred to as SL1 and SL1+IT, respectively. We also included two small deletion mutants that encompassed two exons (del1418-90R) or one exon (del2434-56K) in the proximity of mTOR mutational hotspots (Fig. 1A).
We introduced all of the indicated mutations into myc-tagged cDNA of rat mTOR, which has 99% homology at a protein sequence level to either human or mouse mTOR, in the pRK5 plasmid and transfected such constructs into human embryonic kidney 293 (HEK293) cells. We probed mTORC1 signaling pathway activity in lysates of such cells with protein dot blots against the phosphorylated form of ribosomal protein S6 at serine 235/236 (P-S6) in continuously growing and serum-starved cells (Fig. 1B). Although serines 235/236 can be phosphorylated by kinases outside the mTORC1 pathway, it is rapamycin-sensitive and the antibody against these residues worked consistently well in the dot blot assay. Also, we reasoned that tested mutants should activate the mTORC1 pathway directly and therefore in our experimental setting potential contribution of other kinases to the observed P-S6 changes is very unlikely. As controls, we also probed untransfected cells, cells that were transfected with an empty vector, a kinase-dead (KD) mTOR mutant (D2357E/V2364I) and wild-type mTOR. They all exhibited similar levels of S6 phosphorylation, except the KD variant, which expectedly showed a decrease in phosphorylation compared with all of the other controls (Fig. 1B and C), further proving that P-S6 (235/236) can be used as a readout of mTORC1 activity in our experiments. Mutations that repeatedly yielded a higher signal of S6 protein phosphorylation compared with wild-type under the corresponding conditions were the FAT domain mutants L1460P and C1483Y, kinase domain mutants I2017T, S2215Y and R2505P and quadruple mutant SL1+IT. The quadruple mutant SL1+IT, as previously described, exhibited very high level of activity, completely insensitive to starvation (Fig. 1B and C). In our assays, we did not observe an increase in the activation of any other point mutation that was investigated (i.e. triple mutant SL1 or either of the two deletion mutants), but we cannot exclude the possibility that our assay was not sufficiently sensitive to detect minor differences.
To ensure that we chose a mutation that was suitable to model neurodevelopmental disorders, we next evaluated mutants, increasing mTORC1 signaling in HEK293 cells, in cultured rat primary cortical neurons that developed in vitro. We recloned a subset of mTOR mutants into a vector that is routinely used in our laboratory for overexpression studies in neurons that contain the β-actin promoter and transfected such purified plasmids into DIV7 neurons. After 2 day expression, transfected neurons were starved from the B27 supplement and fixed immediately or after 1, 2 or 4 h. In neurons that were positive for co-transfected green fluorescent protein (GFP), we measured the level of P-S6 by immunofluorescent staining using automated wide-field fluorescence microscopy. We observed very high ability of the quadruple mutant SL1+IT to activate mTORC1 signaling, which was insensitive to starvation. In case of mTOR mutants harboring point mutations differences were much less prominent (Fig. 1D and E). At 0 h time point, significantly increased P-S6 signal was noted in cells overexpressing kinase domain, S2215Y and R2505P, mTOR mutants when compared to wild-type mTOR (Fig. 1D and E). The P-S6 immunofluorescence intensities in those cells also differed significantly from the one in neurons overexpressing FAT domain mTOR mutants—L1460P and C1483Y (Fig. 1D and E). Upon starvation, cells expressing L1460P, C1483Y, I2017T and S2215Y mutants displayed a higher potential to sustain mTORC1 pathway activity. On the other hand, neurons with mTOR R2505P were not significantly different from those with wild-type mTOR (Fig. 1D and E). Although, upon starvation differences between mutants were less evident and consistent (Fig. 1D and E), after 4 h of starvation P-S6 immunofluorescence level was higher in S2215Y mutant transfected cells when compared to L1460P and C1483Y. Still, it was much below P-S6 levels in cells overexpressing SL1+IT mTOR mutant. In summary, these experiments validated in developing neurons the hyperactivation of mTORC1 signaling by some of the previously described mTOR mutants, indicating that the transfection of quadruple mutant SL1+IT leads to an incomparably high level of mTORC1 activity with point mutants found in human pathologies. Our experiments also indicated that in the developing neurons the S2215Y mutation was the one that rendered mTORC1 pathway the most active among naturally occurring mutations, at least for codons L1460, C1483, I2017, S2215 and R2505.
mTOR mutant overexpression in the developing mammalian brain results in cortical neuron mispositioning
mTOR hyperactivity in neural progenitor cells leads to their mispositioning in the developing cortex (5). mTOR consists of 2549 amino acids. Within its primary protein structure, several mutational hotspots in FAT and kinase domains have been identified. Mutations arise in such conditions as cancer (most often in the kidney and large intestine) and cortical dysplasia, leading to variable degrees of downstream signaling pathway upregulation, presumably resulting from various mechanisms and leading to various outcomes. We used our panel of preselected mTOR mutants in neuronal cells that were cultured in vitro to investigate whether their potential to activate the mTORC1 pathway in cultured cells correlates with the degree of neural progenitors misposition in the cortex upon overexpression in vivo. We electroporated plasmids that encoded either GFP alone or mTOR with GFP after internal ribosome entry site (IRES) that allowed for the bicistronic expression of our protein of interest and fluorescent tracer into the left hemisphere in half of the embryos of each dam. As controls specifically for the mTOR mutations, a similar construct but that encoded wild-type mTOR was electroporated into the right hemisphere in the other half of the embryos. After 4 days of development, during which superficial layers of the mouse brain are formed, we analyzed the cortical migration of electroporated progenitors by detecting the immunofluorescent staining of co-expressed GFP in coronal sections of embryonic brains. Using counterstaining with Hoechst 33342, we assessed the borders between the neurogenic VZ/SVZ, neuronal projection-dominated intermediate zone (IZ) and cortical plate (CP) that developed into the cortex. The quantitative analysis of GFP-positive cells in each of the aforementioned regions indicated strong inhibition of the cortical migration of progenitors that expressed the SL1+IT mutant. An average of nearly half of these cells were trapped in the IZ, and less than 40% of the cells reached the CP at this stage of development. By comparison, slightly >10% and >80% of mTOR wild-type-expressing control cells were found within the IZ and reached the CP, respectively. The other mTOR mutant variants did not exhibit any significant differences from wild-type mTOR in their distribution between the VZ/SVZ, IZ and CP (Fig. 2A and B).
Hyperactive mTOR mutant screen for neural development studies. (A) Schematic of C-terminal mTOR protein domain structure (UniProtKB P42346) with the location of point mutations that were considered in the study. (B) Example dot blot for phosphorylated ribosomal protein S6 at serines 235 and 236 (P-S6; green) and tubulin (red) as a loading control on lysates of HEK293 cells that were transfected with pRK5 plasmid with the indicated mTOR variants and starved of serum for the indicated times. (C) Quantification of tubulin-normalized P-S6 signal relative to signal in lysates of mTOR wild-type-transfected non-starved cells. Single experiment values are indicated by open circles joined by opaque dashed lines. Average values are indicated by filled dots joined by vivid dashed lines. (D) Example overlay microscopy images of primary neurons transfected with mTOR (wild-type or its mutants as indicated) and EGFP (green) stained for P-S6 (red) and counterstained with Hoechst 33342 (blue), six per mutant and starvation variant. (E) Quantification of average P-S6 immunofluorescence in the cytoplasm of mTOR-transfected primary neurons relative to the average signal in mTOR wild-type-transfected non-starved neurons. Values from single neurons are indicated as small opaque dots. Average values are indicated as large red dots. Underlying boxplots indicate median value, first and third quartiles with whiskers extending no further than 1.5 inter-quartile range. Asterisks indicate P < 0.05 (Bonferroni-corrected unpaired Wilcoxon test) for the indicated mutant under the indicated starvation condition compared with mTOR wild-type (red) L1460P (green) or C1483Y (blue) under the same starvation condition.
Hyperactive mTOR mutants cause mispositioning of migrating neurons in the developing cortex. (A) Immunofluorescent staining of GFP (green) counterstained with Hoechst 33342 (blue) in neuronal progenitors in E17.5 developing brain sections after in utero electroporation on E13.5 with GFP only or mutants of mTOR-IRES-GFP (left hemispheres) or control wild-type mTOR-IRES-GFP (right hemispheres). (B) Quantification of GFP+ cell fraction in ventricular and SVZs, IZ and cortical plate. (C) The 2.5×-magnified insets of images from a, zooming on cortical plate region of respective specimens. (D) Quantification of progenitor migration in the cortical plate as the relative distance between the apical and basal boarders. Boxplots and black dots indicate the mean relative distance of GFP+ progenitors in the cortical plate in each specimen (annotations in capital black letters) and underlying violin plot that indicates the same for each single neuron (annotations in small gray letters). The P-values are based on Bonferroni-corrected, Wilcoxon test (specimen data) and equal variance, two-tailed distribution t-test (single neuron data).
Hyperactive mTOR mutant-expressing neuron cortical layer mispositioning. (A) Immunofluorescent staining of GFP (green) and deep neuronal layer marker Ctip2 (magenta) counterstained with Hoechst 33342 (blue) of selected mutant variants of the same specimens as in Figure 2. (B) Quantification of GFP+ cell fraction in cortical plate deep and superficial layers. The P-values are based on Bonferroni-corrected, two-tailed distribution unpaired Wilcoxon test.
Hyperactive mTOR mutant-expressing neurons retain the identity of their cortical destination layer. (A) Example photographs of immunofluorescent staining of specimens from the same experiment as in Figure 2 for deep (Ctip2) and superficial (Satb2) cortical layer markers. (B) Quantification of Satb2+ and double-positive cell fraction of all cells from all investigated embryos (indicated by light and dark gray bars) and double-positive cell fraction in each investigated embryo (indicated by dots, with red dots indicating embryos where any positive cells were found). The P-values are based on Bonferroni-corrected Fisher-test.
Morphology of neurons that express hyperactive mTOR mutants from human pathologies is perturbed. (A) Example photographs of specimens from the same experiment as in Figure 2 at 40× magnification. Red dashed boxes indicate neurons with measured soma cross-section; red asterisks indicate multipolar cells. Insets underneath the photographs show example processed images with soma cross-section outlines that were used for quantification. (B) Quantification of fraction of bipolar (light gray bar) versus multipolar (dark gray bar) single neurons and fraction of multipolar cells that were found in each embryo (dots). Red shows embryos where any multipolar cells were found. (C) Quantification of neuron soma size as soma cross-section area. The P-values are based on Bonferroni-corrected Fisher-test (cell morphology data) equal variance, two-tailed distribution t-test (cell size data).
We analyzed in more detail the migratory ability of electroporated progenitors by quantifying their relative position between the apical and basal borders of the CP. We performed this quantification of the average neuron migration distance per embryo and the single cell migration distance (Fig. 2C and D). As expected from previous analyses, the most striking phenotype yielded the most potent mutant, SL1+IT, the expression of which caused progenitors that reached the CP to migrate on average only ~40% of the distance from the apical to basal border of the CP compared with ~80% of the distance for wild-type mTOR-expressing cells. Overexpression of kinase domain mutants (S2215Y and R2505P), which were capable of inducing mTORC1 signaling in cultured neurons under basal conditions, considerably inhibited progenitor migration to reach only an average of ~50% of the apical–basal distance. The L2427T mutant and FAT domain L1460P and C1483Y mutants inhibited progenitor migration to a lesser degree when considering the single cell data (S2215Y versus L2427T, L1460P, C1483Y and R2505P versus L2427T, L1460P, C1483Y and R2505P, P < 0.001, two-tailed distribution t-test with Bonferroni correction). Cells that expressed GFP only exhibited a significant increase in the relative apical–basal distance compared with wild-type mTOR-expressing progenitors, based on the single neuron data (Fig. 2C and D).
To determine precisely where cells that overexpressed the mTOR mutants were located in the cortex, we co-stained sections of the brain after the electroporation of variants that expressed the two FAT domain mutants and two kinase domain mutants for Ctip2 and Tbr2 (data not shown), apart from GFP staining, for electroporated cell visualization. Ctip2 is a marker of deep neuronal layers in the CP, with higher expression in layer V and weaker expression in layer VI, which enables discrimination between deep and superficial cortical layers. Tbr2 is expressed in IPs, allowing for division of the neurogenic niche into the VZ and SVZ. Presumably because of the very low number of GFP-positive cells within the neurogenic niche, we did not observe any meaningful differences in cell percentages between the mutants and corresponding wild-type controls in the VZ and SVZ. Within the CP, however, cells that expressed mTOR mutants clearly failed to reach upper cortical layers II–IV with the efficiency of wild-type mTOR-expressing cells (Fig. 3A and B). Only <20% of the cells that expressed the kinase domain mutants reached superficial cortex layers, and most of them accumulated in layers V and VI, whereas FAT domain mutants caused neuron mispositioning to a lesser extent, allowing ~40% of them to reach superficial layers. By comparison, ~70% of wild-type mTOR-expressing cells reached superficial cortex layers in all of the mutant variant controls (Fig. 3A and B). Statistical analysis confirmed that the percentages of neurons that remained in CP6 or reached CP2–4 were significantly different between kinase domain mutants and C1483Y (for CP6, S2215Y versus C1483Y P < 0.05, R2505P versus C1483Y P < 0.05; for CP2–4, S2215Y versus C1483Y P < 0.05, R2505P versus C1483Y P < 0.01; Wilcoxon test with Bonferroni correction). The difference between kinase domain and L1460P mutants did not, however, reach statistical significance.
To further determine whether neurons that were electroporated with mTOR mutants acquired the identity of DL neurons or remained committed to UL fate, we co-stained sections from the same experiment as shown in Figure 2 with UL and DL markers (Satb2 and Ctip2, respectively; Fig. 4). We scanned the section areas with the most GFP-positive cells using a confocal microscope with a 40× objective and analyzed a minimum of 20 cells per embryo. All GFP-positive cells in all of the variant mutants and wild-type controls were positive for Satb2, with only sporadic cells that co-expressed both markers. No single GFP-positive cell was found to be negative for Satb2.
These results indicate that mTOR mutations that activate the mTORC1 pathway cause neuron mispositioning in the cortex, but mispositioned neurons remained committed to their UL program. The magnitude of this effect to a certain degree reflected the strength of mTORC1 pathway activation that resulted from the corresponding mTOR mutation or the mutation location at the FAT or kinase domain.
Morphological changes in mutant mTOR-overexpressing neurons in the developing brain
During corticogenesis, excitatory neurons undergo morphological changes first from multipolar progenitors to bipolar neurons that undergo radial migration. Upon reaching target layers, neurons develop morphology that is characteristic of the neuron type that they are destined to become and the layer where they reside. Thus, an increase in the appearance of multipolar cells upon mTOR mutant electroporation could suggest a delay in the transition from multipolar to bipolar morphology or acceleration of neuron maturation. Both, in turn, should result in electroporated cell mispositioning. We investigated morphology in higher-magnification images of two FAT domain and two kinase domain mutants that are found in human pathologies. We found that a vast majority of both wild-type and mutant mTOR-overexpressing cells in all four variants manifested bipolar morphology (Fig. 5A and B) and in many embryos, we did not find any single multipolar cell. When comparing single cells that were pooled from all embryos for each variant we noticed significantly more multipolar cells among L1460P, S2215Y and R2505P mutant-expressing embryos compared with wild-type controls. This effect was rather small. Therefore, we concluded that this weak phenotype does not underlie mispositioning of most mutant-expressing cells.
The mTOR pathway controls cell size in neurons. We compared the soma size of mutant and wild-type mTOR-overexpressing neurons and noticed that the former neurons were on average distinctly larger, an effect that was observed in all of the mutant variants that were analyzed (Fig. 5A and B). Comparison of the effects of kinase and FAT domain mutants revealed a significantly stronger impact of R2505P than L1460P and C1483Y mutations on the cell soma size (R2505 versus L1460P, P < 0.001; R2505 versus C1483Y, P < 0.05; Student’s t-test with Bonferroni correction). mTOR S2215Y mutant effect on this parameter differed significantly however only when compared with the L1460P mutant (S2215Y versus L1460P, P < 0.01; S2215Y versus C1483Y, not significant; Student’s t-test with Bonferroni correction). Therefore, we concluded that mutant mTOR overexpression in neurons in the developing brain impacts neuron morphology in terms of neuron soma size but this effect did not fully correlate with a degree to which those mutants potentiated mTORC1 pathway in cultured neurons.
Discussion
Somatic mTOR mutations occur in MCD, a neuropathology that is characterized by disorganization of the cerebral cortex. The few studies that analyzed the effects of selected mTOR mutations on cortex organization in animal models indicated that mTORC1 pathway hyperactivity leads to the improper positioning of neurons in the developing cerebral cortex. However, still unknown is whether different mutations of mTOR result in a different range of cortical disorganization. Also unknown is whether the degree of mTORC1 pathway hyperactivity, potentially resulting from different mutations or mutation locations, correlates with the observed changes in laminar organization of the cortex. Resolving these issues was the goal of the present study. We performed in utero brain electroporation of various mTOR mutants that occur clinically, resulting in the mispositioning of migrating neurons more apically without changing their cell identity. The present results revealed a relationship between mutations that strongly activate mTOR or that are located at a kinase domain and a greater potential of UL neurons to be mispositioned to the DL.
In the present study, we first selected four mTOR mutants (L1460P, C1483Y, S2215Y and R2505P) based on their greatest potential to increase activity upon overexpression in HEK293 cells and developing neurons and the fact that their effects on neuronal migration were unknown at the time the study began. One of the mutants (S2215Y) activated the mTORC1 pathway more strongly than the others, but none of them was as efficient as the artificially created SL1+IT mutant. Upon electroporation in the developing cerebral cortex, only the mTOR SL1+IT mutant caused the significant mispositioning of migrating neurons to the IZ. All of the other mutants and the mTOR L2427T mutant, similar to previously published L2427P one, resulted in rather subtle changes in electroporated neuron positioning within the CP. More specifically, the electroporated cells traveled shorter distances toward the pial surface. This phenotype was more visible in cells that were electroporated with kinase domain mutants (R2505P and S2215Y), one of which was also characterized by higher activity than FAT domain mTOR mutants. Notably, the R2505 somatic mutation has not been described in HME or FCD, but this is not apparently because of its inability to affect neuronal migration but rather because of the relatively low mutation frequency in cancer (COSMIC). Our data are in agreement with Kassai et al. (5), who reported that overexpression of the mTOR SL1+IT mutant led to the accumulation of electroporated cells in the IZ and DL. In the present study, we found similar but much less pronounced effects of the overexpression of mTOR that bore pathologically occurring somatic mutations on neuronal mispositioning relative to previous studies on mTOR L2427P, A1459D and C1483Y, which revealed the accumulation of cells not only in the DL but also in the IZ (8–10). Our observations are more similar to those in Tsc1 and Tsc2 conditional knockout mice (17,18). These previous studies reported more subtle changes in the CP. The reason for the small discrepancy in strength of the observed phenotype after electroporation of the similar mTOR mutants in different laboratories is unclear but could be the result of the different mutant protein expression levels that were achieved or slight differences in actual codon changes studied (L2427 to P versus T and 1459 versus 1460).
mTORC1 pathway hyperactivation in neural stem cells and their progeny leads to various abnormalities that can result in neuron mispositioning. Among the affected processes are neural stem cell proliferative potential, fate determination, radial migration, premature differentiation and the improper interaction of the neuron leading edge with the MZ (7,19–25). These observations come from different models, in which mTORC1 hyperactivation was induced by various means (e.g. Tsc1 or Tsc2 knockout, Pten knockout, Rheb overexpression, mTOR SL1+IT mutation and A1459D or C1483Y overexpression) and often appear to be contradictory. PTEN, Tsc1-Tsc2 complex and Rheb act upstream of mTOR and are known to activate other signaling pathways beyond mTOR (3,4,23). To date, data that have been obtained with the use of mTOR hyperactive mutants, including in the present study, rule out changes in the cell cycle or fate specification as a cause of mislamination. In fact, the overexpression of mTOR SL1+IT as early as E12 did not increase the number of actively proliferating neural stem cells (5). The present study and Park et al. (10) found that active mTOR mutant overexpression did not alter the fate determination of UL neurons, even if they were stopped in the DL. A similar observation was recently reported in migrating neurons that lacked Tsc2 (18). These observations suggest that mTOR hyperactivation delays radial migration or soma translocation when migrating cells reach the MZ. Several recent studies support such a possibility and provide insights into the molecular mechanism that is impaired. For example, mTOR hyperactivation that is caused by the C1483Y mutation impairs autophagy-dependent ciliogenesis (10). As a result, migrating neurons cannot respond properly to Wnt, thus delaying the switch from multipolar to bipolar morphology and causing the mispositioning of UL neurons to the DL. The present study does not fully support such a scenario because we did not observe substantial blockade of migration in areas of the multipolar to bipolar morphology transition, with no increase in the number of cells that overexpressed C1483Y or more active mTOR variant (S2215Y) with multipolar morphology. Nonetheless, we observed, albeit rarely, cells with multipolar morphology only in the case of mTOR mutant electroporation. Moon et al. (18) showed that abnormal neuronal migration that was caused by the loss of Tsc2 was likely attributable to changes in the morphology of a leading process. At the molecular level, Tsc2 knockdown or Rheb overexpression led to the mTOR-driven excessive production of Cullin-5, an E3 ligase that is critical for the degradation of phosphorylated disabled homolog 1, resulting in improper responses to reelin. The postulated pathological mechanism resulted in subtle changes in neuronal positioning that were quite similar to the one reported herein. Such subtle changes were also reported when the intrinsic activity of migrating neurons was induced using designer receptors exclusively activated by designer drugs, which led to an increase in their pausing (20). Migrating neurons express mRNAs for various neurotransmitter receptors (e.g. glutamate and γ-aminobutyric acid). Neurotransmitters are known to regulate the process of migration. Neurotransmitters are also known to upregulate mTOR activity (3). Thus, a reasonable possibility is that mTOR upregulation mimics part of the response to neurotransmitters. Recent observations suggest that mTORC1 hyperactivity extends the time of neuronal migration, but the induction of premature cell maturation could also potentially account for the blockade of migration, which has been described in Pten and Tsc1 conditional knockout mice (24,25).
Recent clinical observations led to two opposite conclusions with regard to the relationship between mTOR signaling hyperactivation and the trajectory of MCD. Mirzaa et al. (16), based on an analysis of several cases, concluded that mTOR mutations that lead to moderate increases in mTORC1 activity lead to HME, whereas those that strongly upregulate mTORC1 result in FCD. In contrast, it was proposed that the type of mTOR mutation has a relatively weak impact, whereas the timing of its occurrence decides whether it leads to HME or FCD (26). The present data showed that the degree of mTOR hyperactivation or mutation in the kinase domain in cases when mTOR mutations occur at the same stage may result in subtle differences in cell mispositioning in the UL. With UL, DL and mispositioned neurons that innervate different parts of the brain, stronger mTOR activation could lead to a different severity of neurological symptoms that are associated with improper cortex wiring (e.g. autism spectrum disorders or mood disorders).
Plasmid vectors used in the study
| Name | Description | Addgene # | Source |
|---|---|---|---|
| pEGFP-N1 | expression of EGFP from CMV promoter | - | Clontech |
| pRK5-myc-mTOR | expression of rat mTOR cDNA from CMV promoter | 1861 | gift from David Sabattini (30) |
| pRK5-myc-mTOR-KD | expression of rat mTOR KD cDNA from CMV promoter | 8482 | gift from David Sabiattini (unpublished) |
| pβ-actin-16pl | expression from beta-actin promoter | - | (31) |
| pCAG-Cre-IRES2-GFP | bicistronic expression from CAG promoter | 26646 | gift from Anjen Chenn (32) |
| Name | Description | Addgene # | Source |
|---|---|---|---|
| pEGFP-N1 | expression of EGFP from CMV promoter | - | Clontech |
| pRK5-myc-mTOR | expression of rat mTOR cDNA from CMV promoter | 1861 | gift from David Sabattini (30) |
| pRK5-myc-mTOR-KD | expression of rat mTOR KD cDNA from CMV promoter | 8482 | gift from David Sabiattini (unpublished) |
| pβ-actin-16pl | expression from beta-actin promoter | - | (31) |
| pCAG-Cre-IRES2-GFP | bicistronic expression from CAG promoter | 26646 | gift from Anjen Chenn (32) |
Plasmid vectors used in the study
| Name | Description | Addgene # | Source |
|---|---|---|---|
| pEGFP-N1 | expression of EGFP from CMV promoter | - | Clontech |
| pRK5-myc-mTOR | expression of rat mTOR cDNA from CMV promoter | 1861 | gift from David Sabattini (30) |
| pRK5-myc-mTOR-KD | expression of rat mTOR KD cDNA from CMV promoter | 8482 | gift from David Sabiattini (unpublished) |
| pβ-actin-16pl | expression from beta-actin promoter | - | (31) |
| pCAG-Cre-IRES2-GFP | bicistronic expression from CAG promoter | 26646 | gift from Anjen Chenn (32) |
| Name | Description | Addgene # | Source |
|---|---|---|---|
| pEGFP-N1 | expression of EGFP from CMV promoter | - | Clontech |
| pRK5-myc-mTOR | expression of rat mTOR cDNA from CMV promoter | 1861 | gift from David Sabattini (30) |
| pRK5-myc-mTOR-KD | expression of rat mTOR KD cDNA from CMV promoter | 8482 | gift from David Sabiattini (unpublished) |
| pβ-actin-16pl | expression from beta-actin promoter | - | (31) |
| pCAG-Cre-IRES2-GFP | bicistronic expression from CAG promoter | 26646 | gift from Anjen Chenn (32) |
Antibodies used in the study
| Species, antigen | Dilution | Conditions (antigen retrieval, blocking) | Company | Cat # |
|---|---|---|---|---|
| rabbit anti-mTOR | 1:1000 | - | Cell Signaling | 2983 |
| rabbit anti-P-S6 S235/6 | 1:500 (DB) 1:200 (IF) | heat-mediated antigen retrieval (IF) | Cell Signaling | 4858 |
| mouse anti-tubulin | 1:1000 (DB) | - | Sigma | T5168 |
| chicken anti-GFP | 1:200–1:1000 | none or heat-mediated antigen retrieval | Abcam | ab13970 |
| rat anti-Ctip2 | 1:200–1:500 | heat-mediated antigen retrieval | Abcam | ab18465 |
| rabbit anti-Satb2 | 1:300 | heat-mediated antigen retrieval | Abcam | ab92446 |
| rabbit anti-Tbr2 | 1:500 | heat-mediated antigen retrieval | Abcam | ab23345 |
| Species, antigen | Dilution | Conditions (antigen retrieval, blocking) | Company | Cat # |
|---|---|---|---|---|
| rabbit anti-mTOR | 1:1000 | - | Cell Signaling | 2983 |
| rabbit anti-P-S6 S235/6 | 1:500 (DB) 1:200 (IF) | heat-mediated antigen retrieval (IF) | Cell Signaling | 4858 |
| mouse anti-tubulin | 1:1000 (DB) | - | Sigma | T5168 |
| chicken anti-GFP | 1:200–1:1000 | none or heat-mediated antigen retrieval | Abcam | ab13970 |
| rat anti-Ctip2 | 1:200–1:500 | heat-mediated antigen retrieval | Abcam | ab18465 |
| rabbit anti-Satb2 | 1:300 | heat-mediated antigen retrieval | Abcam | ab92446 |
| rabbit anti-Tbr2 | 1:500 | heat-mediated antigen retrieval | Abcam | ab23345 |
DB, dot blot; IF, immunofluorescence
Antibodies used in the study
| Species, antigen | Dilution | Conditions (antigen retrieval, blocking) | Company | Cat # |
|---|---|---|---|---|
| rabbit anti-mTOR | 1:1000 | - | Cell Signaling | 2983 |
| rabbit anti-P-S6 S235/6 | 1:500 (DB) 1:200 (IF) | heat-mediated antigen retrieval (IF) | Cell Signaling | 4858 |
| mouse anti-tubulin | 1:1000 (DB) | - | Sigma | T5168 |
| chicken anti-GFP | 1:200–1:1000 | none or heat-mediated antigen retrieval | Abcam | ab13970 |
| rat anti-Ctip2 | 1:200–1:500 | heat-mediated antigen retrieval | Abcam | ab18465 |
| rabbit anti-Satb2 | 1:300 | heat-mediated antigen retrieval | Abcam | ab92446 |
| rabbit anti-Tbr2 | 1:500 | heat-mediated antigen retrieval | Abcam | ab23345 |
| Species, antigen | Dilution | Conditions (antigen retrieval, blocking) | Company | Cat # |
|---|---|---|---|---|
| rabbit anti-mTOR | 1:1000 | - | Cell Signaling | 2983 |
| rabbit anti-P-S6 S235/6 | 1:500 (DB) 1:200 (IF) | heat-mediated antigen retrieval (IF) | Cell Signaling | 4858 |
| mouse anti-tubulin | 1:1000 (DB) | - | Sigma | T5168 |
| chicken anti-GFP | 1:200–1:1000 | none or heat-mediated antigen retrieval | Abcam | ab13970 |
| rat anti-Ctip2 | 1:200–1:500 | heat-mediated antigen retrieval | Abcam | ab18465 |
| rabbit anti-Satb2 | 1:300 | heat-mediated antigen retrieval | Abcam | ab92446 |
| rabbit anti-Tbr2 | 1:500 | heat-mediated antigen retrieval | Abcam | ab23345 |
DB, dot blot; IF, immunofluorescence
Materials and Methods
DNA constructs, oligonucleotides and antibodies
The previously described DNA constructs, primary antibodies and oligonucleotides that were used in this study are listed in Table 1, Table 2 and Supplementary Material, Table S1, respectively. Point mutations and deletions were introduced to mTOR cDNA using QuikChange Mutagenesis kit (Stratagene/Agilent, Santa Clara, CA, USA) or Kapa HiFi Site-Directed Mutagenesis kit (Kapa Biosystems/Roche, Basel, CH) polymerases, using vendor-provided protocols and accordingly designed DNA primers (Supplementary Material, Table S1). cDNAs encoding rat wild-type mTOR and its mutants were sub-cloned from pRK5 to pβ-actin-16pl using traditional restriction-enzyme cloning exploiting ClaI and XbaI. mTOR cDNA (wild-type and mutant variants) was sub-cloned into pCAX-Cre-IRES-EGFP, replacing Cre cDNA, using modified sequence- and ligase-independent cloning (27). In short, both vector and insert DNA fragments were amplified with PCR to introduce 25 bp-long overlaps. The 100 ng vector and 500 ng insert were incubated in chew-back reaction with 0.5 U T4 Polymerase (NEB) in 1× Green Buffer (Fermentas/ThermoFisher, Waltham, MA, USA). After 30 min at room temperature reaction was stopped by adding dCTP to 1 mm concentration and placed on ice, then fragments were assembled for 15 min at 37°C and transformed into MH1 Escherichia coli strain. Correct vector assemblies were validated with analytic restriction enzyme digests targeting both insert and vector and seamless DNA ligations were confirmed using Sanger sequencing across junction sites.
Cell line culture and dot blots
HEK293 cells (American Type Culture Collection, Manassas, VA, USA) were cultured under standard mammalian tissue culture conditions in high-glucose Dulbecco’s Modified Eagle Medium with L-glutamine (4 mm), sodium pyruvate (1.25 mm), penicillin (100 U/ml), streptomycin (0.1 mg/ml) and 10% fetal calf serum (FCS). For protein dot blots, 1 × 103 cells per well were seeded in 96 well plates and transfected the next day using Lipofectamine 2000 (Invitrogen/Thermo Fisher Scientific, Waltham, MA, USA) with plasmids that encoded wild-type mTOR, its mutants or an empty construct together with pEGFP-N1 at a mass ratio of 4:1 to monitor transfection efficiency. The day after transfection, a portion of the cells was starved in medium without FCS. All of the cells were then washed with ice-cold phosphate-buffered saline (PBS) and lysed in Leammli buffer without sodium dodecyl sulfate and bromophenol blue for 10 min at room temperature. The lysates were then diluted 40× with transfer buffer (192 mm glycine and 25 mm Tris–HCl, pH 8.3) and transferred to nitrocellulose membranes using a dot blot apparatus (Bio-Rad, Hercules, CA, USA) with a vacuum pump. Membranes were blocked with 5% non-fat milk in Tris-buffered saline with 0.1% Tween (0.5 mm Tris–HCl [pH 7.5], 1.5 mm NaCl and 0.1% Tween-20 [TBST]), probed overnight at 4°C with primary antibodies that were diluted in 5% bovine serum albumin in TBST and developed with fluorescently labeled secondary antibodies diluted 1:500–1:1000 in blocking solution using the Odyssey system (LI-COR Biosciences, Lincoln, NE, USA). The plasmid and starvation variants were treated and probed in quadruplicate. The experiment was repeated four times. Each time, the dilution series of lysate from control untreated cells was processed in parallel to generate relative protein abundance standard curves.
Primary neuronal cell culture, transfection and immunofluorescent staining
Primary neurons were obtained from embryonic day 18 (E18) Wistar rat embryos from dams that were sacrificed according to a protocol that was approved by the 1st Ethical Committee in Warsaw, Poland (decision no. 288/2012), which was in compliance with European Community Council Directive 2010/63/EU. Cortical primary neuronal cultures were prepared and transfected with plasmid DNA on day 7 in vitro (DIV7) using Lipofectamine 2000 as previously described (28). After 2 days, a portion of the cells was starved of B27, and all of the cells were washed with PBS and fixed with 4% paraformaldehyde (PFA) in phosphate buffer. Fixed neurons were blocked and permeabilized with 5% serum and 0.3% TritonX-100 in PBS, immunostained overnight at 4°C with primary antibodies and developed with Alexa Fluor-labeled secondary antibodies diluted 1:200 (Invitrogen/Thermo Fisher Scientific). GFP-positive cells were scanned at 20× magnification using a CellR wide-field fluorescence high-content screening station (Olympus, Shinjuku, Tokyo, Japan) and analyzed using ScanR software.
In utero electroporation and histology
Fetal mouse brain electroporation was performed according to a protocol that was approved by the 1st Ethical Committee in Warsaw, Poland (decision no. 643/2014, 105/2016 and 277/2017). The procedure was performed with E13.5 FVB embryos according to an established protocol (29) with modification. The dams were anesthetized with a ketamine/xylazine mixture. The uterus with embryos was exposed through an abdominal transverse incision through the skin and peritoneum. Approximately 3 μg of plasmid DNA (1 μg/μl) mixed with FastGreen was injected through a 32 gauge mesotherapy needle using a FemtoJet microinjector (Eppendorf, Hamburg, Germany) in three 0.1 s pulses at 500 hPa. Immediately afterward, the brain was electroporated with 4× 30 V, 50 ms pulses at 950 ms intervals using a NEPA21 electroporator that was connected to 3 mm wide electrodes. The electrodes were oriented at ~45° to the transverse embryo head axis with the cathode brought close to the somatosensory cortex. After electroporation, the uterus was returned to the abdominal cavity. The peritoneum and skin were sutured. Antibiotic (0.2 mg enrofloxacin) and analgesic (0.1 mg meloxicam) were subcutaneously administered, and the dams were returned to their home cages for recovery.
Four days after electroporation, the dams were anaesthetized with CO2 and euthanized by cervical dislocation. Embryos were extracted through a cesarean incision. Their forebrains were fixed in 4% PFA in phosphate buffer overnight at 4°C. The fixed forebrains were suffused with two changes of 30% sucrose in phosphate buffer, embedded in Killik medium (Bio-Optica, Milano, Italy), cut using a cryostat into 20 μm-thick coronal sections, transferred to poly-L-lysine-coated support slides and stored at −20°C. For better attachment to slides during immunofluorescent staining, the sections were dried and straighten on slides 10 min at 55°. Optionally antigen retrieval was performed by incubating the slides in 10 mm citrate buffer (pH 6.0) with 0.05% Tween-20 for 30 min at 95°C. The slides were then washed three times with TBS for 5 min, blocked with blocking buffer at room temperature for 1 h and incubated with primary antibody solution overnight at 4°C. The slides were then washed three times with TBST for 5 min, developed with secondary antibody diluted 1:500 for 30 min at room temperature, washed again and sealed with a coverslip using ProlonGold (Invitrogen/Thermo Fisher Scientific). Cell nuclei were counterstained by adding Hoechst 33342 (Invitrogen/Thermo Fisher Scientific) to the secondary antibody solution or first subsequent TBST wash.
Image acquisition and analysis
For histological section scanning for all of the described analyses, an LSM800 confocal microscope (Carl Zeiss, Oberkochen, Germany) was used. Whole areas of immunofluorescently stained brain sections that were GFP-positive cells were scanned with a 10× objective in four Z-stacks for subsequent maximum projection. Hoechst 33342 staining was used to demarcate borders between the VZ/SVZ, IZ and CP. Tbr2 and Ctip2 immunofluorescent staining was used to distinguish the VZ from the SVZ and to distinguish the CP DLs (V and VI) from the CP ULs (II/III and IV), respectively. The regions of interest (ROIs) and GFP-positive neuron coordinates were manually annotated using the ImageJ 1.51n Fiji bundle (specifically the PointPicker plugin for the latter) and used as inputs for python scripts. Script counting fractions of cells found in specific layer used function from path module of matplotlib library to assign neuron with specific coordinates to enclosing ROI of specific layer. The script-calculating apico-basal migration distance of GFP-positive neurons was used to fit second-degree polynomial curves to the apical and basal borders of the CP-enclosing ROI point set. The closest point on each of these curves to the neuron was then determined using the minimizing function by applying the COBYLA method from the scipy.optimize package. The migration distance is expressed as the distance to the apical border divided by the sum of the distances to the apical and basal borders. The script code is available at https://github.com/btarkowski/CP_migration_distance.
For neuronal morphology assessment, areas with the highest concentration of electroporated neurons on the aforementioned brain sections were scanned using a 40× objective in 36 Z-stacks for subsequent maximum projection. A minimum of 20 GFP-positive cells per embryo were analyzed. Cell polarity was manually assessed using the ImageJ Fiji Cell Counter plugin. For cell soma size analysis, rectangular areas around isolated cell somas were manually selected. These areas were thresholded. Dark and bright outliers were removed, and gray morphology and fill hole functions were used to close the resulting masks. The total area of white pixels in the processed masks served as approximate areas of neuron soma cross sections that were used for quantification.
Statistical analysis
Experiments with HEK293 cells were repeated four times. The hyperactivity of a specific mutant was indicated when it exhibited higher activity than the wild-type control at most timepoints in all four experiments. For the other assays generating measurement variables, we performed unpaired statistical tests. The non-parametric Wilcoxon test was applied when the sample size was small or when the data did not have a normal distribution (i.e. ribosomal protein S6 in neurons in vitro, neuron distribution into VZ/SVZ, IZ and CP and cortical layer staining after in utero electroporation). In the other cases, we performed Student’s t-test for samples with equal variance (i.e. neuron migration within the CP and neuron size). For the assays with nominal variables, Fisher’s test was applied (i.e. cell polarity and neuronal layer marker expression). Statistical tests and plots were programmed with R in RStudio IDE.
Acknowledgements
We thank Dr Andrzej Cwetsch, Dr. Katarzyna Kowalska and Dr. Artur Czupryn for protocols and support implementing techniques, Dr Anna Kosson for providing experimental animals for in utero electroporation and Alina Zielińska for technical support in the laboratory and animal facility.
Conflict of Interest statement. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Funding
Polish National Science Centre (2012/04/S/NZ3/00264 to B.T. and 2017/27/B/NZ3/01358 to J.J.); 7FP [602391, ‘EPISTOP’ and Polish Ministerial funds for science (years 2014–2018) to J.J.]; Polish National Center for Research and Development (EPIMARKER no. STRATEGMED3/306306/4/2016 to J.J.); Foundation for Polish Science ‘Mistrz’ Professorial Subsidy (to J.J.).
