The small GTPase RhoA is required to maintain spinal cord neuroepithelium organization and the neural stem cell pool - PubMed (original) (raw)
Comparative Study
The small GTPase RhoA is required to maintain spinal cord neuroepithelium organization and the neural stem cell pool
Dominik Herzog et al. J Neurosci. 2011.
Abstract
The regulation of adherens junctions (AJs) is critical for multiple events during CNS development, including the formation and maintenance of the neuroepithelium. We have addressed the role of the small GTPase RhoA in the developing mouse nervous system using tissue-specific conditional gene ablation. We show that, in the spinal cord neuroepithelium, RhoA is essential to localize N-cadherin and β-catenin to AJs and maintain apical-basal polarity of neural progenitor cells. Ablation of RhoA caused the loss of AJs and severe abnormalities in the organization of cells within the neuroepithelium, including decreased neuroepithelial cell proliferation and premature cell-cycle exit, reduction of the neural stem cell pool size, and the infiltration of neuroepithelial cells into the lumen of the ventricle. We also show that, in the absence of RhoA, its effector, mammalian diaphanous-related formin1 (mDia1), does not localize to apical AJs in which it likely stabilizes intracellular adhesion by promoting local actin polymerization and microtubule organization. Furthermore, expressing a dominant-negative form of mDia1 in neural stem/progenitor cells results in a similar phenotype compared with that of the RhoA conditional knock-out, namely the loss of AJs and apical polarity. Together, our data show that RhoA signaling is necessary for AJ regulation and for the maintenance of mammalian neuroepithelium organization preventing precocious cell-cycle exit and differentiation.
Figures
Figure 1.
Recombination of the conditional RhoA allele in NEPs of the spinal cord. A, Regulatory sequences of the Brn4 promoter drive the expression of the Cre recombinase in NEPs. During _Brn4–Cre-_mediated recombination the genomic region between the LoxP sites is excised; in case of the conditional RhoA allele, exon 3 is excised leading to the inactivation of the RhoA gene. Recombination of the lacZ reporter gene leads to the expression of β-galactosidase in recombined neuroepithelial cells. B, Immunohistochemistry on forelimb spinal cord cross-sections of E10.5 embryos demonstrates efficient loss of RhoA protein in the spinal cord but not in other tissues. Scale bar, 100 μm.
Figure 2.
RhoA is required for the organization of the ventricular zone. A–J, Spinal cord cross-sections stained with hematoxylin and eosin; magnifications of the lumen and the central spinal cord region (A′–J′). Early in development (A, B), the ventricular structure is normal in mutant spinal cords and the lumen is formed correctly. At E11.5, mutant spinal cords show dysplasias. Compared with controls (C, C′), the normal epithelial organization is lost and the ventricular structure is disorganized with formation of rosette-like structures and invasive mesenchymal-like cells present in the spinal cord lumen (D, D′). At E12.5, the epithelial organization of the VZ is progressively lost in mutant spinal cords (F, F′) compared with controls (E, E′). At E13.5, cells from the VZ dispersed throughout the mutant spinal cords and the lumen almost disappeared (H, H′ compared with G, G′). Subsequently, at E14.5, the VZ and the lumen of RhoA mutant spinal cords completely disappeared (J, J′ compared with I, I′). Arrowheads point to rosette-like structures present in mutant spinal cords. Scale bars: A–J, 100 μm; A′–J′, 20 μm.
Figure 3.
RhoA is required for proliferation, survival, and localization of mitotic cells. A–C, Proliferation was assessed in control and mutant spinal cords by expression of Ki67 (A) and mitotic marker phospho-histone-H3 (B). Cell death was examined by TUNEL in control and mutant spinal cords (C). D, The proliferation ratio was determined by the quotient of Ki67-positive cells per total cells of the spinal cord. At E10.5, before apparent morphological defects, the proliferation ratio was significantly decreased from 69.29 ± 0.66% in the control to 54.28 ± 0.62% in the mutant (n = 3, p = 0.0001) and at E11.5 from 35.35 ± 0.7% in the control to 30.96 ± 0.49% in the mutant (n = 3, p = 0.0069), respectively. At E12.5, the proliferation ratio in the control (17.39 ± 0.23%) was not significantly altered compared with the mutant (16.57 ± 0.13%). E, The relative number of mitotic cells in the spinal cord was examined by quantification of phosphor-histone-H3-positive cells to total spinal cord cells. The percentage of mitotic cells was unchanged between control (E10.5, 4.04 ± 0.2%; E11.5, 1.27 ± 0.01%; E12.5, 0.44 ± 0.03%) and mutant (E10.5, 4 ± 0.05%; E11.5, 1.48 ± 0.15%; E12.5, 0.61 ± 0.03%) spinal cords. F, Whereas the relative number of mitotic cells was unchanged, more cells that are not dividing at the ventricular surface can be observed in the mutant spinal cords (arrows in B). The ratio of non-ventricular dividing cells to all mitotic cells was quantified and is significantly increased in the mutant (E10.5, 30.56 ± 2.05%; E11.5, 44.16 ± 0.58%; E12.5, 91.35 ± 0.46%) compared with control (E10.5, 11.67 ± 1.81%; E11.5, 4.68 ± 1.17%; E12.5, 11.63 ± 1.74%) spinal cords at all stages analyzed (E10.5, n = 3, p = 0.0014; E11.5, n = 3, p = 0.0001; E12.5, n = 3, p = 0.0001). G, The ratio of TUNEL-positive cells to total cells present in control and mutant spinal cords was quantified. At E10.5, no significant difference can be observed between control (4.6 ± 1.43%) and mutant (5.6 ± 1.21%). At E11.5, a significant increase in the ratio of TUNEL-positive cells from 5.07 ± 0.33% in the control to 8.56 ± 0.97% in the mutant was found (n = 3, p = 0.0273) and at E12.5 from 1.86 ± 0.13% in the control to 3.75 ± 0.53% in the mutant (n = 3, p = 0.0259). H, The decreased proliferation and increased cell death leads to a significant lower number of cells present in mutant spinal cords from E11.5 onward. The total cell number was determined by quantifying DAPI-positive nuclei, and the numbers of cells at E10.5 in the control spinal cord were set to 100% (control: E10.5, 100 ± 2.59%; E11.5, 264.37 ± 3.4%; E12.5, 365.77 ± 7.35%; mutant: E10.5, 101.02 ± 6.51%; E11.5, 208.80 ± 7.52%; E12.5, 285.64 ± 6.01%). Scale bars, 100 μm.
Figure 4.
RhoA is required to keep NEPs in the cell cycle. BrdU was given 20 h before analysis. Arrowheads indicate that cells that have incorporated BrdU (BrdU+) but were not proliferative anymore (Ki67−). The fraction of cells being BrdU+ Ki67− to all cells being BrdU+ was determined. At E10.5, cell-cycle exit is increased from 28.35 ± 1.34% in the control to 43.89 ± 0.35% (n = 3, p = 0.0004) and at E11.5 from 32.88 ± 0.48 to 39.56 ± 0.85% (n = 3, p = 0.0023). Scale bar, 60 μm.
Figure 5.
RhoA is required to maintain neural stem/progenitor cells. A, Immunohistochemistry for the stem/progenitor marker Sox2 and neuronal marker NeuN. B, Immunohistochemistry for Doublecortin (DCX) showed a qualitative increase of early neurons in RhoA mutant spinal cords. C, D, Quantification of staining showed in B. The percentage of NeuN-positive cells is increased in the mutants (C), and consequently the percentage of Sox2-positive cells is decreased (D). The percentage of NeuN-positive cells increases from 27.81 ± 0.39% in the control to 39.74 ± 0.75% at E10.5 (n = 3, p = 0.0001) and from 63.11 ± 0.84 to 69.87 ± 0.65% at E11.5 (n = 3, p = 0.0035). Meanwhile, the Sox2-positive fraction decreases from 72.19 ± 0.39% in the control to 60.26 ± 0.75% in the mutant at E10.5 (n = 3, p = 0.0001) and from 36.92 ± 0.85% in the control to 30.14 ± 0.65% in the mutant at E11.5 (n = 3, p = 0.0031). Scale bars, 100 μm.
Figure 6.
Normal Notch, Wnt, MSX1, and Shh expression in RhoA mutants. A–D, Expression of Hes5 (A), Wnt-1 (B), MSX1 (C), and Shh (D), examined by in situ hybridization, is not changed at E10.5 and E11.5 in RhoA mutant spinal cords compared with control spinal cords, indicating normal Notch and dorsoventral signaling in mutant RhoA spinal cords. Quantification of the expression domains relative to the total area of the spinal cords are not significantly changed (data not shown). Scale bars, 100 μm.
Figure 7.
RhoA is required for the maintenance of cell–cell adhesions. A–L′ Immunohistochemistry to reveal the structure of cell–cell adhesion on control and mutant spinal cord cross-sections. In RhoA mutants, the staining pattern for the cell–cell adhesion markers N-cadherin and β-catenin is altered at E10.5, and the staining is less intense, less structured, and not continuous at the ventricular surface (arrows in B, F, J and in higher magnification in B′,B″, F′, F″, J′, J″). Cortical actin is less intense and appears to be more diffuse and less well structured (J,J′,J″) in RhoA mutant spinal cords. At E11.5, the gaps in mutant ventricular surface cell–cell adhesions are more prominent and are identified as the areas in which cells invade the spinal cord lumen (D, H, L and in higher magnification in D′, H′, L′). Scale bars: A–L, 20 μm; A′–L′, A″–J″, 10 μm.
Figure 8.
RhoA localizes mDia1 but not ROCK1 to the apical membrane. A, Immunohistochemistry for the Rho downstream effector mDia1 show that this protein is poorly localized to the ventricular surface of mutant RhoA spinal cords (arrows in c, c′ and in higher magnification in d, d′). B, The localization of another important Rho downstream effector, ROCK1, shows no obvious alteration in RhoA mutant spinal cords. Scale bars: a, a′, c, c′, 20 μm; b, b′, d, d′, 10 μm.
Figure 9.
Overexpression of DN mDia1 in neural stem progenitor cells results in the loss of AJs and apical polarity. Forebrain progenitors of wild-type E12.5 embryos were electroporated in utero with a GFP expression vector (control) or coelectroporated with the GFP vector and DN mDia1 expression vector and analyzed 24 h later. Transfected cells were identified by GFP expression. A–F”, Confocal images of immunohistochemistry for N-cadherin (A, A′, B, B′ and _z_-projections of higher magnification in E, E″, F, F″) show a loss of AJs at the ventricular surface in neural stem/progenitor cells transfected with DN mDia1 (areas of lost AJs marked by arrows in B, B′, and F, F″) but not in cells expressing GFP only (A, A′ and E, E″). Apical polarity and disruption of PKCζ localization is lost after DN mDia1 expression (marked by an arrow in D, D′) compared with GFP-expressing control areas (C, C′). Scale bars: A–D′, 50 μm; E–F″, 20 μm.
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