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Hydraulic control of mammalian embryo size and cell fate

Nature volume 571, pages 112–116 (2019)Cite this article

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Abstract

Size control is fundamental in tissue development and homeostasis1,2. Although the role of cell proliferation in these processes has been widely studied, the mechanisms that control embryo size—and how these mechanisms affect cell fate—remain unknown. Here we use the mouse blastocyst as a model to unravel a key role of fluid-filled lumen in the control of embryo size and specification of cell fate. We find that there is a twofold increase in lumenal pressure during blastocyst development, which translates into a concomitant increase in cell cortical tension and tissue stiffness of the trophectoderm that lines the lumen. Increased cortical tension leads to vinculin mechanosensing and maturation of functional tight junctions, which establishes a positive feedback loop to accommodate lumen growth. When the cortical tension reaches a critical threshold, cell–cell adhesion cannot be sustained during mitotic entry, which leads to trophectoderm rupture and blastocyst collapse. A simple theory of hydraulically gated oscillations recapitulates the observed dynamics of size oscillations, and predicts the scaling of embryo size with tissue volume. This theory further predicts that disrupted tight junctions or increased tissue stiffness lead to a smaller embryo size, which we verified by biophysical, embryological, pharmacological and genetic perturbations. Changes in lumenal pressure and size can influence the cell division pattern of the trophectoderm, and thereby affect cell allocation and fate. Our study reveals how lumenal pressure and tissue mechanics control embryo size at the tissue scale, which is coupled to cell position and fate at the cellular scale.

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Fig. 1: Blastocyst development is driven by increased lumenal pressure and cortical tension of trophectoderm.
Fig. 2: Lumenal expansion promotes junctional maturation.
Fig. 3: Stretch-induced cortical stiffening and mitosis collectively lead to blastocyst collapse.
Fig. 4: Physical model predicts the change in blastocyst size and oscillation dynamics with perturbed cortical tension.
Fig. 5: Lumenal pressure couples cell positioning and fate specification.

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Data availability

Source Data for Figs. 1, 3, 5 and Extended Data Figs. 1, 2, 4, 5 are available in the online version of the paper. Any other relevant data are available from the corresponding authors upon reasonable request.

Code availability

Code for theoretical model is available on GitHub at https://github.com/truizherrero/Hydraulic-control-of-embryo-size.

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Acknowledgements

We thank all laboratory members and the EMBL animal facility for support; M. Furuse for GST-C-CPE/CPE313 plasmids; A. Aulehla, D. Gilmour, F. Graner, E. Hannezo, S. Hopyan, P. Liberali, R. Prevedel and J. Solon for feedback; T. Higashi, A. Miller and C. Schwayer for the protocol on fluozin leakage assay. C.J.C. and G.M. are supported by EMBL Interdisciplinary Postdocs (EIPOD) fellowship under Marie Sklodowska-Curie Actions COFUND (664726). T.H. is supported by EMBL, Deutsche Forschungsgemeinschaft (DFG) and the European Research Council (742732). T.R.-H. was supported by the Simons Foundation. L.M. thanks the MacArthur Foundation and the Radcliffe Institute for support. A.D.-M. is supported by EMBL and DFG (DI 2205/2-1).

Author information

Authors and Affiliations

  1. European Molecular Biology Laboratory, Heidelberg, Germany

    Chii Jou Chan, Maria Costanzo, Gregor Mönke, Martin Bergert, Alba Diz-Muñoz & Takashi Hiiragi

  2. Paulson School of Engineering and Applied Sciences, Harvard University, Cambridge, MA, USA

    Teresa Ruiz-Herrero & L. Mahadevan

  3. Department of Biology, Drexel University, Philadelphia, PA, USA

    Ryan J. Petrie

  4. Department of Physics, Harvard University, Cambridge, MA, USA

    L. Mahadevan

  5. Department of Organismic and Evolutionary Biology, Harvard University, Cambridge, MA, USA

    L. Mahadevan

  6. Kavli Institute for Bionano Science and Technology, Harvard University, Cambridge, MA, USA

    L. Mahadevan

  7. Institute for the Advanced Study of Human Biology (WPI-ASHBi), Kyoto University, Kyoto, Japan

    Takashi Hiiragi

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Contributions

C.J.C. conceived the project, designed the experiments and wrote the manuscript with input from all authors. C.J.C. and M.C. performed and analysed all experiments; G.M. performed inter-collapse interval quantification; R.J.P. assisted with the initial setup of the micropressure system; M.B. and A.D.-M. assisted with atomic force microscopy measurements; T.R.-H. and L.M. created the theory, performed simulations and suggested experimental tests of the theory. L.M. contributed to the writing of the manuscript. T.H. supervised the study, helped to design the project, performed oocyte recovery and contributed towards writing the manuscript.

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Correspondence to Chii Jou Chan, L. Mahadevan or Takashi Hiiragi.

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Extended data figures and tables

Extended Data Fig. 1 Blastocyst size is controlled by an embryo-autonomous mechanism.

a, Schematic representing mouse blastocyst development. The blastocyst, surrounded by a glycoprotein coat of the zona pellucida, is partitioned into the epithelial trophectoderm, the ICM and the cavity. The outer trophectoderm is further classified into the mural trophectoderm (which surrounds the blastocyst cavity) and the polar trophectoderm, which is in contact with the ICM. Apicobasal polarity is marked by the outer apical domains (red) and trophectoderm cell–cell contacts with the apically localized tight junctions and adherens junctions. Na+/K+ -ATPase pumps, enriched at the basolateral membrane of the trophectoderm cells, help to establish an osmotic gradient across the trophectoderm to drive fluid influx (green arrows). Together with the maturation of tight junctions that helps to seal the blastocyst against fluid outflow via paracellular junctions, blastocyst expansion then proceeds. b, Images of embryos without zona pellucida undergoing blastocyst development (representative data from three independent experiments). The corresponding cavity diameter as a function of time (bottom) shows cycles of blastocyst collapse and re-expansion similar to those of embryos with intact zona-pellucida. Scale bar, 20 μm. Dotted line denotes the cavity. Time is shown as h:min after E3.5. c, Top, images of E3.5 blastocyst expressing mTmG (left) and after segmentation (right). Bottom, box plot showing trophectoderm basal area normalized to cell volume in embryos of various sizes at the E3.5 (black, n = 11, 20 and 20 embryos for 1/4, 1/2 and whole embryos, respectively) and E4.5 stages (red, n = 16, 21 and 22 embryos for 1/4, 1/2 and whole embryos, respectively). Scale bar, 20 μm. Box plots show median, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points.

Source data

Extended Data Fig. 2 Quantifying cavity pressure in mouse embryos using a micropressure probe.

a, b, Characteristics of successful cavity pressure measurements. Top, images showing two mouse blastocysts before and during pressure measurement. The baseline reading is close to zero when the microelectrode (0.5 μm) is not in contact with the blastocysts. A transient pressure spike is recorded as the tip of the microelectrode penetrates the zona pellucida. This is followed by a stable reading for about 5–10 s, before some potential leaks occur, which may cause a gradual drop in the reading with time. c, d, Proof-of-principle experiments to verify the pressure measurement. c, The addition of Bb(−) (25 μM) leads to a transient reduction in cavity size and pressure within 10 s. d, The accuracy of the device was further demonstrated by comparing the cytoplasmic pressure of mouse oocytes at the germinal vesicle (GV) stage, measured directly by the micropressure probe and indirectly by Laplace’s law though micropipette aspiration (Pc = 2γ/R, in which γ is the cortical tension of the oocyte measured by micropipette aspiration, and R is the radius of the oocyte). Data obtained by both approaches are not significantly different (Mann–Whitney U test). N, number of oocytes. Scale bars, 20 μm. All data show representative examples from three or more independent experiments.

Source data

Extended Data Fig. 3 Vinculin is recruited towards the tight junctions in a tension-dependent manner during blastocyst expansion.

a, b, Immunostaining of wild-type embryos at the late blastocyst stage (E4.25), showing co-localization of ZO1 (green) with ppMRLC (red) (a) and MYH9 (red) (b). Line scans are shown on the right. c, d, Immunostaining of wild-type embryos at the late blastocyst stage (E4.25), showing co-localization of vinculin (red) with occludin (green) (c) and non-localization with α-catenin (green) (d). Line scans are shown on the right. A and B mark the apical and basal sides of trophectoderm cell–cell junctions. Scale bars, 5 μm. e, Left, vinculin is more enriched at the mural trophectoderm junctions than at the polar trophectoderm junctions in E4.25 blastocysts (box plot shown at top right, n = 61 cells from 11 embryos), which correlates with higher cortical tension in mural trophectoderm cells compared to those of polar trophectoderm cells (bottom right, n = 167 cells from 25 embryos). Scale bars, 20 μm. f, Top, immunostaining of blastocysts shows stronger signals of vinculin at trophectoderm junctions of reduced blastocysts (1/4 and 1/2 blastocysts) compared to whole embryos at E3.75, but equally strong signals in all blastocysts at E4.25. Scale bars, 10 μm. Bottom, box plot of vinculin signal intensity in various blastocysts at E3.75 (black, n = 58 cells from 15 embryos) and E4.25 (red, n = 54 cells from 17 embryos). An overall higher intensity of vinculin at E4.25 across all blastocysts is consistent with the trend in trophectoderm cortical tension (Fig. 1e). g, Immunostaining of vinculin in Bb(−)-treated and control embryos at the E4.25 stage. Bottom, box plot of vinculin signal intensity in control (n = 52 cells from 11 embryos) versus Bb(−)-treated blastocysts (n = 28 cells from 5 embryos). Scale bar, 20 μm. h, Schematic depicting tension-dependent vinculin recruitment at tight junctions, possibly through actin binding, as trophectoderm cells are increasingly stretched during cavity expansion. Mann–Whitney U test. All box plots show median, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points. In a-d, representative examples from three independent experiments are shown.

Extended Data Fig. 4 Leakage of blastocoel fluid during trophectoderm cell division leads to the collapse of blastocysts at mature stage.

a, Images of a blastocyst (at E4.25) showing blastocoel loaded with 4-kDa FITC–dextran dye, before (top) and after (bottom) collapse. Dye leakage following blastocyst collapse occurs preferentially at cell–cell junctions between mitotic trophectoderm cells (white arrows). Scale bar, 20 μm. b–d, Cavity diameter as a function of time (after E3.5) for 1/2 embryos treated with 2 μg ml−1 of aphidicolin (n = 11 embryos) (c) and 50 nM of nocodazole (n = 17 embryos) (d) compared to control 1/2 embryos (n = 17 embryos) (b). e, Images showing that nocodazole-treated 1/2 blastocysts that failed to expand upon initial collapse can re-cavitate upon washout. Scale bar, 30 μm. f, Cortical tension of mural trophectoderm cells measured during interphase (black) and metaphase (red) at mid- (E4.0) and late (E4.5) blastocyst stages. Different symbols correspond to the different cells being measured. Inset, image showing tension measurement for a trophectoderm cell undergoing mitosis at E4.5 (green indicates nucleus at M-phase). In a, e, f, representative examples from three independent experiments are shown.

Source data

Extended Data Fig. 5 Expansion rates of the blastocyst cavity scale with the total cell number.

a, Cavity diameter at the plateau stage for whole blastocysts (without zona pellucida) and reduced blastocysts (1/2 blastocysts, blue; 1/4 blastocysts, green). b, Box plot of volume expansion rate for reduced and whole blastocysts shows that blastocysts expand at a rate that scales approximately with the initial number of cells. For a, b, n = 14, 21 and 16 embryos for whole, 1/2 and 1/4 embryos, respectively. c, Time evolution of cavity volume for six whole embryos (representative examples from four independent experiments). The slopes showed a gradual increase with time as the cell number increased. d, Box plot of volume expansion rate for control 1/2 embryos (n = 20 embryos) compared to 1/2 embryos (n = 11 embryos) treated with aphidicolin (2 μg ml−1), which arrests cells at S-phase entry. Mann–Whitney U test. Blastocysts with a reduced number of cells expressed fewer Na+/K+-ATPase pumps overall, which led to a reduced fluid influx and cavity expansion rate. Box plots show median line, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points.

Source data

Extended Data Fig. 6 Measurement of apparent trophectoderm stiffness (Young’s modulus) by atomic force microscopy.

a, Force–distance curve measured on a mature blastocyst (E4.25). Blastocysts were typically indented with a loading force of 5 to 10 nN. Data show representative examples from four independent experiments. Dashed line is the fit of the approach curve to the shell model (see Methods). b, Measured trophectoderm epithelial thickness as a function of cavity radius. Data pooled from n = 31 embryos in 4 independent experiments. The thinning of the trophectoderm shell is a natural consequence of blastocyst expansion, as predicted by the model (see Supplementary Notes). The value of individual trophectoderm shell thickness is used for fitting in a, to extract the apparent trophectoderm stiffness (see Methods).

Extended Data Fig. 7 Regulation of blastocyst size by cortical tension, tight-junction permeability, ion influx and cell–cell adhesion.

a, Dose-dependent volume expansion rate in Bb(−)-treated embryos (red, n = 15, 8 and 4 embryos for 5, 12.5 and 25 μM, respectively) and m−/− Myh9+/− embryos (blue, n = 12 embryos), compared to the wild type (black, n = 23 embryos). b, c, Maximum cavity diameter reached for wild type (black, n = 14 embryos), m−/− Myh9+/− (blue, n = 12 embryos) and embryos treated with Bb(−) (red) in various concentrations (n = 15, 8 and 4 embryos for 5, 12.5 and 25 μM, respectively) (b). These embryos all have similar numbers of cells (c), which shows that the reduced cavity size is not due to a reduced number of cells. n = 28, 24 and 31 embryos for wild-type, m−/− Myh9+/− and Bb(−)-treated embryos, respectively. d, Volume expansion rate in CPE- (red, n = 12 embryos), ouabain- (blue, n = 17 embryos), hypertonic- (pink, n = 28 embryos) and ECCD1-treated (green, n = 23 embryos) embryos compared to controls (black, n = 23 embryos). e, Maximum cavity diameter for CPE- (red, n = 14 embryos), ouabain- (blue, n = 17 embryos), hypertonic- (pink, n = 25 embryos) and ECCD1-treated (green, n = 16 embryos) embryos compared to controls (black, n = 14 embryos). f, ECCD1-treated embryos have similar numbers of cells to those of controls (dark blue, E5.0), whereas CPE-treated embryos have slightly higher numbers of cells than do control embryos (black, E4.25), despite their reduced cavity size. n = 17 and 26 embryos for 313 (CPE131, used as control) and CPE-treated embryos; n = 21, 26 and 28 embryos for control, ouabain- and hypertonic-treated embryos; and n = 18 and 17 embryos for control and ECCD1-treated embryos, respectively. g–i, Embryos treated with Y-27632 (a Rho-kinase inhibitor that targets actomyosin contractility) showed a reduced expansion rate (g) and cavity size (h), despite having cell numbers that were similar to those of controls (i). n = 14, 13, 11 and 10 embryos for controls, 10, 20 and 40 μM Y-27632-treated embryos, respectively, in g and h; n = 19 and 15 for control and Y-27632-treated embryos, respectively, in i. All box plots show median line, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points. Mann–Whitney U test. ****P < 1 × 10−5.

Extended Data Fig. 8 Maternal heterozygous knockout and zygotic heterozygous (m+/− Myh9+/−) embryos exhibit smaller cavity and increased ICM compared to wild type in the same littermate.

a, Immunostaining of late-stage (E4.25) m+/− Myh9+/− and wild-type embryos in the same littermate (m+/+ Myh9+/+), showing trophectoderm (CDX2, magenta), epiblast (SOX2, grey) and primitive endoderm (GATA4, red) fates. Data are a representative example from three independent experiments. Scale bar, 20 μm. b–e, Volume expansion rate for m+/− Myh9+/− (n = 11 embryos) is significantly lower than that of m+/+ Myh9+/+ (n = 11 embryos) embryos (b), which leads to a smaller cavity size (c) and higher ratio of ICM to trophectoderm (e), despite the range in cell number being similar for both genotypes (d). n = 17 and 18 embryos for m+/+ Myh9+/+ and m+/− Myh9+/− embryos, respectively, in d and e. f, Immunostaining of m+/− Myh9+/− embryos (E4.25) showing misallocation of CDX2-expressing cells in the ICM (white arrowhead). Scale bar, 20 μm. Right, quantification of lineage misallocation in m+/− Myh9+/− embryos. n = 35 and 24 embryos for m+/+  Myh9+/+ and m+/− Myh9+/− embryos, respectively. All box plots show median line, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points. Mann–Whitney U test.

Extended Data Fig. 9 Increased cortical tension leads to reduced cavity expansion rate and size, whereas ECCD1 treatment preserves tight junctions.

a, Volume expansion rate in embryos treated with 20 μM lysophosphatidic acid (LPA, n = 16 embryos), and 0.5 nM calyculin A (CalyA, n = 21 embryos) compared to wild type (n = 38). LPA and calyculin A are known to activate cortical contractility. b, Maximum cavity diameter reached for wild type (n = 14 embryos), LPA (n = 16 embryos) and calyculin A (n = 25 embryos). c, Total cell number for wild type (n = 30 embryos) and embryos treated with LPA (n = 15 embryos) and calyculin A (n = 11 embryos) is similar, which indicates that the reduced expansion rate and cavity size is not due to a reduced cell-proliferation rate. Box plots show median line, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points. Mann–Whitney U test. d, e, Tight junctions remain sealed and functional, despite impaired functions of adherens junctions. d, Top, immunostaining of E5.0 embryos treated with 50 μg ml−1 of ECCD1 (representative example from three independent experiments). A and B indicate the apical and basal sides of cell–cell junctions, respectively. Bottom, corresponding line scans show that tight junctions appear intact, as shown by the decoupling of signals of ZO1 and occludin from E-cadherin. A.U., arbitrary units. Scale bar, 5 μm. e, Top, tight-junction leakage assay shows that an ECCD1-treated embryo loaded with fluozinc (100 μM) displayed no increase in fluorescence intensity within the cavity when ZnCl2 (200 μM) was added. Bottom, percentage of blastocysts showing no difference in permeability (green) in ECCD1-treated embryos compared to control embryos. N, number of embryos. Scale bar, 20 μm.

Extended Data Fig. 10 Reduced lumen expansion biases cell allocation towards the ICM through increased asymmetric division of outer cells.

a, Schematic depicting a DiI dye assay that assesses how outer trophectoderm (TE) cells contribute to the ICM through asymmetric divisions. b, Box plot of ratios of DiI-positive cell number within the ICM to the total ICM cell number, in control (n = 22 embryos) and hypertonic-treated embryos (n = 24 embryos). Box plot shows median line, 25th and 75th percentiles, and whiskers extending to maximum and minimum data points. Mann–Whitney U test. c, Time-lapse images of hypertonic-treated CDX2–GFP×H2B–mCherry embryo labelled with DiI dye, during blastocyst expansion. An outer cell (yellow arrowhead) undergoes asymmetric division to generate one trophectoderm-forming and one ICM-forming cell; the latter carries the DiI signal (magenta arrow) as it internalizes into the ICM. CDX2 (but not H2B) expression is downregulated (magenta arrowhead) following internalization, whereas CDX2 expression in the outer cells remains constant or increases throughout blastocyst development. Data show a representative example from three independent experiments. Time is shown as h:min after nuclear envelope breakdown. Scale bar, 20 μm. d, Schematic represents how lumenal expansion affects cell positioning and fate acquisition. Reduced lumenal expansion increases the frequency of asymmetric division of outer cells, which generates a trophectoderm- and an ICM-forming cell; the latter eventually acquires ICM fate.

Supplementary information

Supplementary Information

Supplementary Notes describes the theoretical approach to study the growth and dynamics of mouse blastocysts during development.

Reporting Summary

Video 1

Development of a blastocyst with zona pellucida (ZP). DIC imaging of a ZP-intact embryo from early (E3.5) to late blastocyst stage (E4.5). Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 2

Development of a blastocyst without zona pellucida (ZP). DIC imaging of a ZP-free embryo from early (E3.5) to late blastocyst stage (E4.5). Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 3

Development of 1/2 blastocysts. DIC imaging of 1/2 embryos during blastocyst development. Collapse events typically occurred prior to 12:00, earlier than that for the whole embryos. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 4

Development of 1/2 blastocysts treated with aphidicolin. DIC imaging of 1/2 embryos treated with 2 μg/ml of aphidicolin, during blastocyst development. Blastocysts typically showed continual expansion of cavity without any collapse prior to 18:00. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 5

Development of 1/2 blastocysts upon aphidicolin washout. DIC imaging showed that embryos that were treated with 2 μg/ml of aphidicolin (E3.5 to E4.0) started to show collapse events upon washout (E4.0 to E4.0 + 6 hr). Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 6

Development of 1/2 blastocysts treated with nocodazole. DIC imaging of 1/2 embryos treated with 50 nM of nocodazole, during blastocyst development. Blastocysts showed constant fluid leakage and failed to expand. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 7

Development of blastocysts treated with blebbistatin (Bb(−)). DIC imaging of embryos treated with 5 μM of Bb(−), during blastocyst development. Blastocysts showed a reduced cavity expansion rate and did not exhibit any collapse events. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 8

Development of blastocysts treated with calyculin A (CalyA). DIC imaging of embryos treated with 0.5 nM of CalyA, during blastocyst development. Blastocysts showed more frequent collapse prior to 12:00. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 9

Development of 1/4 blastocysts. DIC imaging of 1/4 embryos during blastocyst expansion. Collapse events typically occurred prior to 12:00, earlier than that for the whole embryos. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 10

Development of maternal homozygous KO and zygotic heterozygous (m−/−Myh9+/−) embryos. DIC imaging of m−/−Myh9+/− embryos during blastocyst expansion, showing reduced expansion rate of the cavity. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 11

Development of blastocysts treated with lysophosphatidic acid (LPA). DIC imaging of embryos treated with 20 μM of LPA, which activates TE cortical tension. Time (h:mm). 00:00 denotes E3.5 stage. Time-lapse imaging consists of an image taken every 15 min and is displayed at 10 fps.

Video 12

Tracking of TE division pattern in hypertonic-treated blastocysts. Fluorescent imaging (top) of a hypertonic-treated, DiI-labeled CDX2-GFP×H2B-mCherry embryo undergoing blastocyst expansion, shown by bright field imaging (bottom). Reduced lumenal expansion leads to more frequent asymmetric division of outer cells, which generates one TE- and another ICM-forming cell (labeled with DiI) that downregulates its CDX2 expression with time. Fluorescent images are compiled from different z-slices to track the dividing TE cell and ICM-forming cell while bright field images are compiled at the same z-slice. Time-lapse imaging consists of an image taken every 20 min and is displayed at 7 fps.

Video 13

Tracking of TE division pattern in WT blastocysts. Fluorescent imaging (top) of a WT, DiI-labeled CDX2-GFP×H2B-mCherry embryo undergoing blastocyst expansion, shown by bright field imaging (bottom). Asymmetric division of outer TE cells to generate an ICM-forming daughter cell is rarely observed. Fluorescent images are compiled from maximum projection of several z-slices to better visualize the ICM while bright field images are compiled at the same z-slice. Time-lapse imaging consists of an image taken every 15 min and is displayed at 7 fps.

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Chan, C.J., Costanzo, M., Ruiz-Herrero, T. et al. Hydraulic control of mammalian embryo size and cell fate. Nature 571, 112–116 (2019). https://doi.org/10.1038/s41586-019-1309-x

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  • DOI: https://doi.org/10.1038/s41586-019-1309-x

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