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A monoastral mitotic spindle determines lineage fate and position in the mouse embryo

Nature Cell Biology volume 24, pages 155–167 (2022)Cite this article

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Abstract

During mammalian development, the first asymmetric cell divisions segregate cells into inner and outer positions of the embryo to establish the pluripotent and trophectoderm lineages. Typically, polarity components differentially regulate the mitotic spindle via astral microtubule arrays to trigger asymmetric division patterns. However, early mouse embryos lack centrosomes, the microtubule-organizing centres (MTOCs) that usually generate microtubule asters. Thus, it remains unknown whether spindle organization regulates lineage segregation. Here we find that heterogeneities in cell polarity in the early 8-cell-stage mouse embryo trigger the assembly of a highly asymmetric spindle organization. This spindle arises in an unusual modular manner, forming a single microtubule aster from an apically localized, non-centrosomal MTOC, before joining it to the rest of the spindle apparatus. When fully assembled, this ‘monoastral’ spindle triggers spatially asymmetric division patterns to segregate cells into inner and outer positions. Moreover, the asymmetric inheritance of spindle components causes differential cell polarization to determine pluripotent versus trophectoderm lineage fate.

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Fig. 1: A monoastral mitotic spindle in the mouse embryo.
Fig. 2: The monoastral spindle is assembled by two separate MTOCs in different subcellular regions.
Fig. 3: Asymmetric organization of the monoastral mitotic spindle.
Fig. 4: The monoastral spindle drives asymmetric cell division.
Fig. 5: Asymmetric inheritance of microtubules and microtubule regulators from the monoastral spindle during cell division.
Fig. 6: Spindle manipulations disrupt division patterns and inner–outer cell segregation.
Fig. 7: Heterogeneities in keratin filament assembly differentially stabilize the cell cortex to trigger monoastral spindle formation in a subset of cells.

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Data availability

Source data are provided with this paper. All data supporting the findings of this study are available from the corresponding author on reasonable request. Source data are provided with this paper.

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Acknowledgements

This work was supported by grants from the National Institutes of Health: National Institute of General Medical Sciences (GM139970-01) and Eunice Kennedy Shriver National Institute of Child Health and Human Development (HD102013-01A1).

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Authors and Affiliations

  1. Department of Cell and Developmental Biology, Institute for Regenerative Medicine, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA

    Oz Pomp, Robin M. Skory, Adam A. Moverley, Piotr Tetlak, Stephanie Bissiere & Nicolas Plachta

  2. Institute of Molecular and Cell Biology, ASTAR, Singapore, Singapore

    Hui Yi Grace Lim

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Contributions

O.P. conceived the project, performed the experiments and data analysis and wrote the manuscript with contributions from all other authors. H.Y.G.L., R.M.S. and A.A.M. assisted with experiments and data analysis. P.T. and S.B. performed mouse work and embryo microinjection experiments. N.P. supervised the project.

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Correspondence to Nicolas Plachta.

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Nature Cell Biology thanks William Bement, Marta Shahbazi and the other, anonymous reviewer(s) for their contribution to the peer review of this work.

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Extended data

Extended Data Fig. 1 Additional characterization of the monoastral mitotic spindle.

a, Additional examples of 8- to 16-cell stage mouse embryos displaying monoastral and anastral mitotic spindles. Dashed lines mark cortical microtubules. Data are from seven independent experiments. b, Light-scattering measurements in embryos injected with GFP mRNA at the 2-cell stage and imaged during the 8- to 16-cell stage. Quantification of GFP fluorescence along the axial (z) axis reveals no light scattering along the entire depth of a cell. Error bars represent s.e.m. Data are from three independent experiments. c, Monoastral spindles frequency is similar in different genetic backgrounds (p = 0.99 by Kolmogorov-Smirnov test). Scale bars, 5 µm in xyz.

Source data

Extended Data Fig. 2 Additional characterization of PLK1 and spindle assembly dynamics.

a,b, Live embryos show localization of PLK1-Emerald to the bridges, connecting sister cells, during interphase (b), and disappearance from the bridge prior to mitosis (b). c, Consistent with its localization on kinetochores from late prophase, PLK1-Emerald puncta are visualized at chromosomes (inset). Bona fide apical and perinuclear PLK1-Emerald-positive MTOCs are also visualized (arrows). d, Live-imaging shows the dynamics of PLK1-localization at the intercellular bridge, cell nucleus and forming spindle. e, Live imaging and graph show that PLK1-Emerald is enriched in the nucleus before NEBD. f, Live-imaging shows that the onset of microtubule growth from the apical and perinuclear MTOCs of the monoastral spindle corresponds with the timing of NEBD. The apical MTOC starts to assemble the astral array (arrows), while the perinuclear MTOC starts to assemble the first spindle half of the future interpolar parts of the spindle. g, Following NEBD, the two main MTOCs that form the monoastral mitotic spindle are moving closer. h, i, Immunofluorescence in non-injected 8-cell embryos. At prophase, microtubules are projected between the apical and perinuclear MTOCs (h) to assemble the monoastral spindle (i). j, Localization pattern of endogenous PLK1 at the monoastral mitotic spindle. k, Live-imaging. In these focal planes, only the apical MTOC is initially visible. Note how the apical MTOC joins the rest of the mitotic spindle. l, Live-imaging during the assembly of an anastral mitotic spindle. In some cases of anastral spindle assembly, an apical EB3-Tomato-positive structure is initially detected, but this fails to display significant microtubule growth and to join the rest mitotic spindle apparatus. Graph shows the proportion of cells displaying this behavior. m, Quantification of PLK1-Emerald and Cdk5rap2-Emerald fluorescence intensities at the main parts of the monoastral and anastral mitotic spindles. Note the retention of high fluorescence levels in the apical parts of the monoastral spindle. n, Immunostaining and live imaging of PLK1 and microtubules and quantification of PLK1 intensity ratio between the apical and the basal poles (arrow shows apical pole). In box plot, centre line is the median, box edges show upper and lower quartiles and whiskers represent the range. **p = 0.0025; Two-tailed Mann-Whitney U-test. o, Quantification of apical MTOC stability in monoastral and anastral spindles. Scale bars, 10 µm in a,b, d-l, n; 2 µm in c. Source numerical data are provided in source data.

Source data

Extended Data Fig. 3 Additional characterization of the asymmetric organization of the monoastral spindle.

a, Selected time-frames from live-imaging experiments performed in whole 8-cell embryos comparing the different division pattern of a cell with an original monoastral or anastral mitotic spindle. Data are from six independent experiments. b, Scheme and quantification of the variability in α-tubulin fluorescence intensity from pole-to-pole along monoastral and anastral spindles. Scale bars, 5 µm. Source numerical data are provided in source data.

Source data

Extended Data Fig. 4 Additional characterization of asymmetric cell division pattern in cells with monoastral spindles.

a, Live-imaging of a cell undergoing asymmetric cell division. Data are from seven independent experiments. b, Disruption of E-cadherin trans interactions using acute treatment with the DECMA1 antibody pushes cells to adopt more outer positions displaying a larger apical than basal surface area. *p = 0.0357; Unpaired two-tailed Mann Whitney U-test. In box plot, the centre line is the median, box edges show upper and lower quartiles and whiskers represent the range. Data are from three independent experiments. c, Live-imaging of GFP-myosin II together with RFP-Utr and H2B-RFP marker reveal two main modes of cell in the intact 8-cell embryo. In some cases, GFP-myosin II accumulates asymmetrically at the cell cortex, closer to the basal region. This corresponds with a more orthogonal division angle and differences in outer-inner cell volumes, similar to cells with monoastral mitotic spindles. Conversely, in some cases, GFP-myosin II accumulates more symmetrically at the cell equator. This is followed by more symmetric division patterns and outer-outer daughter volumes, similar to cells with anastral spindles. Data are from five independent experiments. d, Example of computer segmentation analysis used to track spindle types, division patterns and cell position. Monoastral spindles are detectable by their larger apical than basal regions and their asymmetry in microtubule intensities and orthogonal orientations. Note that with this resolution, the thinner astral microtubule array is not computationally segmented and is thus not visible. Cell tracking permits to follow the more asymmetric division pattern of this cell. Note the asymmetries between the two daughter cells in the inheritance of microtubules associated with the spindle of the parental cell and their final position. Data are from six independent experiments. Scale bars, 10 µm. Source numerical data are provided in source data.

Source data

Extended Data Fig. 5 Additional characterization of spindle manipulations.

a, Protein sequence comparison between mouse katanin, Xenopus tropicalis, and Xenopus laevis sequences. Phosphorylation site serine 131, which inhibits katanin activity, is found in X. laevis but missing from X. tropicalis. Mouse katanin has this phosphorylation site at serine 133 (marked in red). In line with Xenopus experiments64, elimination of the phosphorylation site (katanin-S133G) decreases spindle length. Only a portion of the protein sequences is shown. b, Immunofluorescence for PLK1 in an 8-cell embryo injected into one cell of the 2-cell stage with PLK1 siRNAs and EB3-Tomato. Cyan fluorescence shows injected cells. The image shows a thin confocal section (10 µm thickness) scanned with high laser power to reveal differences in PLK1 levels between non-injected and injected cells at the same focal planes. Graph shows reduction of PLK1 intensity in the injected cells (Mann Whitney U-test). c, Microinjection of Katna1 siRNAs reduces the expression of katanin p60 mRNA, quantified by qPCR in 16-cell embryo homogenates. d,e, Embryos treated with PLK1-inhibitor volasertib produce fewer inner-cells (Mann Whitney U-test) (d), and produce fewer monoastral spindles. f, Correlation between spindle angles (metaphase) and relative volumes of daughter cells (telophase). g, Step-by-step protocol used to segment microtubule asters and interpolar regions of the mitotic spindles (related to Figs. 1d and 4g,l). h, Computer segmentation of inner and outer daughter cells after division shows the effect of PLK1 and katanin manipulations on cell volume. i, Relative daughter cell volumes after manipulations of PLK1 (Mann Whitney U-test and Kruskal–Wallis). j, n, Spindle length after PLK1 and katanin manipulations. Mann Whitney U-test (j) and Kruskal–Wallis test (n). NS, not significant. k, o, Length ratio between the two interpolar parts of the spindle after PLK1 and katanin manipulations. Mann Whitney U-test (k) and Kruskal–Wallis test (o). l, p, Quantifications of the variability in α-tubulin fluorescence intensity from pole-to-pole along spindles after PLK1 and katanin manipulations (see scheme in Extended Data Fig. 3b). m, Examples of mitotic spindles in live embryos following katanin manipulations. Scale bars, 10 µm in b,h,m; 3 µm in g. Source numerical data are provided in source data.

Source data

Extended Data Fig. 6 Additional characterization of F-actin ring formation by asymmetric inheritance of microtubules.

a, Live-imaging of EB3-Tomato, Ezrin-GFP and H2B-GFP shows F-actin clearance by microtubules in an outer cell after division. b, Staining for α-tubulin, Phalloidin-Rhodamine and DAPI in non-injected 8-cell embryos. Left panel shows additional example of a cell undergoing division. A more extensive microtubule network is present on the apical side. c, Live-imaging shows retention of microtubules at the apical cortex of the outer cell during interphase. d, Live-imaging for RFP-MAP2C and Utr-GFP shows examples of F-actin ring formation in outer cells derived from monoastral (left panels) and anastral spindles (right panels). The outer cell derived from the monoastral spindle forms a ring which is larger than those formed by the two outer daughters derived from the anastral spindle. e, Microtubule density under the apical cortex in daughter cells derived from parental cells with monoastral or anastral spindles, quantified two hours after division. Right panels show the effect of PLK1 and katanin manipulations. In box plots, the centre line is the median, box edges show upper and lower quartiles and whiskers represent the range. *p = 0.0499, **p = 0.0059 for volasertib, **p = 0.007 for katanin; Two-tailed Mann Whitney U-test and Kruskal–Wallis test were used to test significance. f, Scheme and quantification of apical PLK1 stability after keratin manipulations. Scale bars, 10 µm. Source numerical data are provided in source data.

Source data

Extended Data Fig. 7 Spindle manipulations affect F-actin ring formation, apical polarization and Cdx2 levels.

a, Embryos treated with Latrunculin A or microinjected at the 1-cell stage with RhoA-T19N display smaller F-actin rings assessed by Phalloidin-Rhodamine staining, lower apical polarization assessed by antibodies against phospho-Ezrin and lower Cdx2 levels. In the segmented 3D views, the F-actin rings have been computationally traced to highlight their size. The embryos were fixed at the 16-cell stage. b, Embryos microinjected into one cell at the 2-cell stage and fixed at the 16-cell stage show the effects of PLK1 and katanin manipulation on F-actin rings, Phospho-Ezrin and Cdx2 levels. Scale bars, 10 µm.

Extended Data Fig. 8 Additional characterization of spindle and ring formation.

a, Immunofluorescence for α-tubulin in embryos treated with Cytochalasin D. Disruption of the cortical actin by drug treatment triggers the formation of multiple ectopic MTOCs. b, Live-imaging of Utr-GFP and RFP-MAP2C in intact embryo shows a lack of correlation between the apical domain’s size and the orientation of the mitotic spindle. Note how the apical spindle pole does not preferentially target the border of the apical domain. Moreover, the spindle shown is aligned relatively orthogonal in a cell displaying a larger apical domain. This contrasts with models suggesting that a small apical domain determines orthogonal spindle positioning. c, Live-imaging of RFP-Utr and H2B-GFP in an intact embryo shows an additional example of apical domains disassembly during mitosis. d, Graph shows the number of inner-cells in the 16-cell stage embryos. Embryos treated with SiR-actin produce more inner-cells than controls. The centre line is the median, box edges show upper and lower quartiles and whiskers represent the range. **p = 0.0019; Two-tailed Mann Whitney U-test was used to test significance. e, Live-imaging of EB3-Tomato injected with Pard6b siRNA. Note that distal-MTOC origin from the cell-cell junctions (lateral MTOC) instead of the apical cortex, compared to Fig. 2c and Extended Data Fig. 2f,g,k. Scale bars, 10 µm. Source numerical data are provided in source data.

Source data

Extended Data Fig. 9 Summary of main findings.

Schematic model shows the central steps by which cells assemble monoastral and anastral spindles, and how spindle organization determines division patterns and inner–outer lineage fate. Smaller insets highlight the main results supporting the model. Scale bars, 5 µm.

Supplementary information

Reporting Summary

Supplementary Table

siRNAs list

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Live imaging of mitotic cells

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Pomp, O., Lim, H.Y.G., Skory, R.M. et al. A monoastral mitotic spindle determines lineage fate and position in the mouse embryo. Nat Cell Biol 24, 155–167 (2022). https://doi.org/10.1038/s41556-021-00826-3

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