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Molecular control of macroscopic forces drives formation of the vertebrate hindgut

Nature volume 565, pages 480–484 (2019)Cite this article

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Abstract

The embryonic gut tube is a cylindrical structure from which the respiratory and gastrointestinal tracts develop1. Although the early emergence of the endoderm as an epithelial sheet2,3 and later morphogenesis of the definitive digestive and respiratory organs4,5,6 have been investigated, the intervening process of gut tube formation remains relatively understudied7,8. Here we investigate the molecular control of macroscopic forces underlying early morphogenesis of the gut tube in the chick embryo. The gut tube has been described as forming from two endodermal invaginations—the anterior intestinal portal (AIP) towards the rostral end of the embryo and the caudal intestinal portal (CIP) at the caudal end—that migrate towards one another, internalizing the endoderm until they meet at the yolk stalk (umbilicus in mammals)1,6. Migration of the AIP to form foregut has been descriptively characterized8,9, but the hindgut is likely to form by a distinct mechanism that has not been fully explained10. We find that the hindgut is formed by collective cell movements through a stationary CIP, rather than by movement of the CIP itself. Further, combining in vivo imaging, biophysics and mathematical modelling with molecular and embryological approaches, we identify a contractile force gradient that drives cell movements in the hindgut-forming endoderm, enabling tissue-scale posterior extension of the forming hindgut tube. The force gradient, in turn, is established in response to a morphogenic gradient of fibroblast growth factor signalling. As a result, we propose that an important positive feedback arises, whereby contracting cells draw passive cells from low to high fibroblast growth factor levels, recruiting them to contract and pull more cells into the elongating hindgut. In addition to providing insight into the early gut development, these findings illustrate how large-scale tissue level forces can be traced to developmental signals during vertebrate morphogenesis.

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Fig. 1: The chick hindgut forms by antero-posterior inversion of endoderm passing through the CIP.
Fig. 2: A contractile gradient drives polarized collective cell movements to form hindgut.
Fig. 3: A posterior FGF8 gradient modulates endoderm tensional gradient to control collective cell movements.
Fig. 4: A one-dimensional quantitative mechanochemical model for an FGF8-mediated contractile gradient mechanism that causes collective cell movements.

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Data availability

Source data for Figs. 1–4 and Extended Data Figs. 1–10 are available in the online version of the paper.

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Acknowledgements

We thank M. Hutson for the dnFGFR1-IRES-GFP plasmid, P. Tschopp and O. Pourquié for valuable discussions, microscopy core MicRoN (Microscopy Resources on the North Quad) at Harvard Medical School, and Sigrid Knemeyer of SciStories LLC for artwork. We acknowledge funding support from the NIH (F32 HD069074, N.L.N.; F32 DK103563, C.L.; R01 HD089934, C.J.T.) and the MacArthur Foundation (L.M.).

Reviewer information

Nature thanks D. Gilmour, N. Nascone-Yoder, K. Weijer and the other anonymous reviewer(s) for their contribution to the peer review of this work.

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Author notes

  1. Nandan L. Nerurkar

    Present address: Department of Biomedical Engineering, Columbia University, New York, NY, USA

  2. Nandan L. Nerurkar

    Present address: Department of Genetics and Development, Columbia University Medical Center, New York, NY, USA

Authors and Affiliations

  1. Department of Genetics, Harvard Medical School, Boston, MA, USA

    Nandan L. Nerurkar, ChangHee Lee & Clifford J. Tabin

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

    L. Mahadevan

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

    L. Mahadevan

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

    L. Mahadevan

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

    L. Mahadevan

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  1. Nandan L. Nerurkar
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Contributions

N.L.N. and C.J.T. conceived and designed experiments. C.L. cloned the DUSP6 reporter and performed sprouty1 in situ hybridizations with assistance from N.L.N. All other experiments (including vital dye injections, electroporations, live imaging, immunofluorescence and tension measurements) and analysis of those experiments were carried out by N.L.N. under the supervision of C.J.T. The mathematical model was formulated, solved and interpreted by N.L.N. and L.M. The manuscript was prepared by N.L.N. and revised by N.L.N., C.L., L.M. and C.J.T.

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Correspondence to Nandan L. Nerurkar or Clifford J. Tabin.

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Extended data figures and tables

Extended Data Fig. 1 Electroporation-based transfection of the chick endoderm.

a, Dorsal view of HH18 embryo (left) stained with PHK26 to visualize hindgut (HG), and quantification (right) of gut length and allantois displacement from HH14 (t = 0 h, n = 4 embryos). Data are mean ± s.d.; unpaired t-test. Scale bar, 100 µm. b, DiO (green arrow), DiI (red arrow) and DiA (yellow arrow) injected into endoderm (E), notochord (N) and paraxial mesoderm (PM), respectively (image representative of 5 embryos from 5 independent experiments) upon injection at HH14 (t = 0 h) and at t = 12 h (left), and quantification of antero-posterior velocity (right, n = 8 embryos (endoderm, paraxial mesoderm) and n = 7 embryos (notochord)). Data are mean ± s.d.; one-way ANOVA with Sidak corrections. *P < 0.05 versus endoderm. Scale bar, 100 µm. c, Ventral view of posterior HH14 embryo following endoderm-specific electroporation with GFP reporter plasmid at HH12 (image representative of ten embryos from ten independent experiments). White arrowhead, CIP; *, ventral lip of the allantois. Scale bar, 100 µm. d, HH18 embryo following endoderm-specific electroporation with GFP reporter plasmid; image representative of eight embryos from eight independent experiments. GFP expression can be seen extending from presumptive midgut endoderm, which remains ventrally open (1), into the hindgut, which has been internalized (2) and extends to the tip of the tail bud. Scale bar, 500 µm. e, Transverse sections through the posterior HH18 embryo of GFP-electroporated embryo as indicated in d; representative images from five chick embryos in five individual experiments. Scale bar, 100 µm. f, To test whether cell proliferation could provide a contributing force to collective cell movements, endoderm proliferation was quantified by colocalization of phospho-histone H3 immunofluorescence (red) with electroporation-based endodermal expression of H2B–GFP (green). Boxes, enlarged at right, correspond to presumptive hindgut (PHG) endoderm (1), and node-adjacent endoderm (N) (2). Quantification indicates that ~1% of PHG cells are mitotic, suggesting that cell division is not likely to be a major contributor to collective movements. Cell numbers are as indicated from n = 4 embryos. Data are mean ± s.d.; unpaired two-tailed t-test; *P < 0.05 versus PHG. Scale bar, 100 µm. g, h, To test whether cell movements are required for hindgut formation, we physically blocked these movements by insertion of a tantalum foil barrier into the endoderm of GFP-electroporated embryos at HH14. In control embryos, in which barrier insertion at an anterior location permitted posterior cell movements, the hindgut formed normally: GFP-expressing cells formed a hollow epithelial tube extending into the tail bud, as indicated in sections from anterior to posterior (left to right) through the HH18 tailbud (g; 3 of 3 embryos formed hindgut). However, when the barrier was inserted posteriorly, blocking endoderm movements through the CIP, hindgut formation was disrupted, despite continued outgrowth of the tail bud (h; 1 of 4 embryos formed hindgut) Scale bar, 100 µm.

Source data

Extended Data Fig. 2 Immunofluorescent detection of polarized epithelial markers in the definitive endoderm.

a–d, Preceding gut tube formation, the endoderm exhibits several hallmarks of a polarized epithelium, including a laminin-rich basement membrane (a), basolateral E-cadherin (b), and apical localization of aPKC (c) and ZO-1 (d). Endoderm was visualized by electroporation with GFP, and staining was performed in anterior (left) and posterior (right) endoderm. White arrowheads, apical or junctional localization (image representative of 3 embryos from 3 independent experiments). Scale bar, 10 µm.

Extended Data Fig. 3 An endodermal contraction gradient based on relative tension measurements.

a, Schematic of strain calculation from cell movements (left) and physical interpretation of the two-dimensional strain components εxy, εxx and εyy (right). x and y axes coincide with medio-lateral and antero-posterior embryonic axes, respectively. At the cellular level, stretching (positive strains) and compaction (negative strains) in a continuous epithelial sheet may be achieved by changes in cell shape or cell–cell contacts that result in an increase or decrease in the distance between centroids of neighbouring cells, respectively. See Supplementary Methods for details of the computation and interpretation of strains in the context of active materials. b, Linear cell density in posterior HH15 endoderm (n = 871 cells from 4 embryos). Data are mean ± s.d.; one-way ANOVA with Dunnett’s correction; *P < 0.05 versus posterior-most bin. c, Correlation of cell area with antero-posterior position; Pearson’s two-tailed coefficient, r = 0.55; n = 616 cells from 3 embryos; each circle and colour indicates one cell and embryo, respectively. d, Dorso-ventral (baso-apical) view of GFP-electroporated cells in the anterior (top left) and posterior (bottom left) endoderm, and negative correlation of cell height with position (right); Pearson’s two-tailed coefficient r = 0.48; n = 522 cells from 5 embryos; each circle and colour indicates one cell and embryo, respectively. Scale bar, 10 µm. e, HH15 posterior embryo in contact with Gastromaster tip, diameter D (left), and following local endoderm ablation (right), which creates a cut of size L (dashed line). Relative tension was measured as the ratio L/D. Scale bar, 100 µm. f, Severed notochord visible following Gastromaster ablation. Samples in which the notochord was cut in addition to the endoderm were not included in the analysis. g, Relative tension measured over time after cutting at t = 0 (n = 3 embryos). Wound-healing response was initiated within 1–2 min, decreasing cut size (not shown). Data are mean ± s.d.; one-way ANOVA with Tukey’s correction; NS, not significant versus t = 0. h, Negative correlation of relative tension in wild-type HH15 posterior endoderm with position. Pearson’s two-tailed correlation, r = 0.94, n = 22 embryos. i, Relative posterior tension at HH15 is significantly reduced following treatment with 100 µM Y-27632 (Rho kinase inhibitor) or 100 µM blebbistatin (non-muscle myosin inhibitor), when compared to treatment with 0.1% DMSO (n = 10 embryos each). Data are mean ± s.d.; one-way ANOVA with Tukey’s correction; *P < 0.001 versus 0.1% DMSO. j, Comparison between relative tension along antero-posterior (A-P, n = 7 embryos) and medio-lateral (M-L, n = 4 embryos) axes. Data are mean ± s.d.; unpaired two-tailed t-test.

Source data

Extended Data Fig. 4 Antero-posterior morphogenetic strains in the hindgut-forming endoderm.

a–c, Average antero-posterior strain εyy (n = 3 embryos, mean ± s.d.) versus position for embryos electroporated with GFP (a), dnFGFR1-IRES-GFP (b) or FGF8-IRES-GFP (c). Large negative (compaction) strains observed in embryos electroporated with GFP were eliminated upon misexpression of dnFGFR1 or FGF8.

Source data

Extended Data Fig. 5 Endoderm cell movements and strains are disrupted with inhibition and activation of actomyosin contractility.

a–c, Representative cell tracks (HH14 to HH16) in embryos electroporated with GFP and exposed to 0.1% DMSO (a; 0 of 3 embryos affected), 2.5 µM cytochalasin D to disrupt actomyosin activity (b; 3 of 3 embryos affected), or 20 nM calyculin A to uniformly increase myosin activity (c; 3 of 3 embryos affected). Scale bar, 100 µm. d–f, Average antero-posterior strain εyy (n = 3 embryos, mean ± s.d.) versus position from time-lapse experiments for embryos electroporated with GFP and treated with 0.1% DMSO (d), 2.5 µM cytochalasin D (e) or 20 nM calyculin A (f).

Source data

Extended Data Fig 6 FGF signalling in the posterior endoderm.

a, b, Fluorescent-section in situ hybridization for FGF8 (a) and sprouty1 (also known as SPRY1, b) in sagittal sections through the posterior HH15 embryo (left). Numbered boxes indicate regions that are magnified on the right; the dashed line demarcates the boundary between endoderm (E) and mesoderm (M). Scale bars: 100 µm (left), 10 µm (right). Gradient expression of both genes observed in three of three embryos. c, Isolated signal from endoderm only (left) and quantification of relative fluorescence (right). d, Immunofluorescent detection of di-phospho ERK (dp ERK1/2) with boxed regions magnified below; the dashed line marks the boundary between endoderm (E) and mesoderm (M). Scale bar, 100 µm (top), 10 µm (bottom). Right, quantification of dp ERK1/2 per cell along the antero-posterior axis (n = 3 embryos). RFU, relative fluorescent units. e, Embryos cultured in the presence of 50 µM SU5402 from HH14 to HH18 (left; outlined region is magnified (top right)) displayed normal anterior development but posterior defects, including axis truncation, allantois malformation and failure of the posterior endoderm to internalize and form hindgut, compared to 0.1% DMSO control (bottom), n = 7 of 9 affected versus 0 of 10 in 0.1% DMSO. Scale bar, 1 mm. f, Somitogenesis is unaffected by exposure to 50 µM SU5402 in ex ovo culture, despite dependence of this process on FGF signalling, suggesting that effects of SU5402 were primarily restricted to the endoderm. HH13 embryos were cultured for 9 h in the presence of either 0.1% DMSO or 50 µM SU5402 following DiI injection into the last-formed somite (red) at the time of exposure (n = 3). Scale bar, 100 µm. g, dp ERK1/2 staining in the posterior endoderm following incubation of embryos from HH13 to HH15 in the presence of either 0.1% DMSO (top, n = 3 embryos) or 50 µM SU5402 (bottom, n = 3 embryos). Effects of SU5402 were most pronounced in endoderm, with some subtle reduction in dp ERK1/2 extending into the subadjacent mesodermal cells. The white dashed line indicates the boundary between endoderm (E) and mesoderm (M). Scale bar, 10 µm. h, Quantification of order parameter for embryos electroporated with GFP, dnFGFR1-IRES-GFP, or FGF8-IRES-GFP. n = 4 embryos per condition; data are mean ± s.d.; one-way ANOVA with Tukey’s correction; NS, not significant versus GFP.

Source data

Extended Data Fig. 7 Role of FGF in modulating endodermal tension via direct effects on actomyosin activity.

a, To detect the timescale across which inhibition of FGF signalling induces a reduction in endoderm tension, embryos were exposed to either 0.1% DMSO or 50 µM SU5402 for up to 120 min after onset of exposure, and relative tension was measured over time (n = 5 embryos per group per time point). A significant decrease in tension was observed within 30 min of exposure to SU5402. Data are mean ± s.d.; two-way ANOVA with Tukey’s correction; *P < 0.05 versus DMSO at equivalent time point. b, To test efficacy of cycloheximide (CHX) treatment for inhibition of protein translation, embryos were electroporated with GFP and then incubated for 6 h in the presence of either 0.1% DMSO or 20 µM CHX. In contrast to DMSO-treated controls (3 of 3 embryos were GFP+), GFP signal was undetectable in CHX-treated embryos (0 of 3 embryos were GFP+). Scale bar, 100 µm. c, Relative tension at HH15 following incubation in 0.1% DMSO or 20 µM CHX in the presence or absence of 500 ng µl−1 HsFGF8 (n = 9 embryos per condition). Data are mean ± s.d.; one-way ANOVA with Tukey’s correction; overbars indicate comparison for stated P value.

Source data

Extended Data Fig. 8 FGF signalling controls tissue strain and contractility in the posterior endoderm.

a, b, Sagittal sections (n = 3) from HH15 embryos stained with RhoA–GTP and electroporated with GFP or FGF8-IRES-GFP (a) in the anterior endoderm, or with GFP or dnFGFR1-IRES-GFP (b) in the posterior endoderm. Scale bar, 10 µm. c, Quantification of RhoA–GTP immunofluorescence per cell in anterior and posterior endoderm (n = 3 embryos per condition). Data are mean ± s.d.; two-way ANOVA with Tukey’s correction; *P < 0.05 versus posterior. d, e, Immunofluorescent detection of total RhoA in HH15 sagittal sections following electroporation with GFP or FGF8-IRES-GFP (d) in the anterior endoderm (n = 4 embryos), or with GFP or dnFGFR1-IRES-GFP (e) in the posterior endoderm (n = 4 embryos); the dashed line demarcates the boundary between endoderm (E) and mesenchyme (M). Scale bar, 10 µm. f, RhoA enrichment in posterior versus anterior endoderm quantified on a per-cell basis from immunofluorescent detection at HH15 (n = 4). Data are mean ± s.d.; one-way ANOVA with Tukey’s correction; *P < 0.05 versus GFP. g, h, Immunofluorescent detection (g) and quantification (h) of phospho-myosin light chain (pMLC) in the posterior endoderm for embryos cultured from HH13 to HH15 in the presence of either 0.1% DMSO (n = 3) or 50 µM SU5402 (n = 3); dashed white lines indicate the boundary between endoderm (E) and mesoderm (M). Data are mean ± s.d.; unpaired two-tailed t-test. Scale bar, 10 µm. i, To determine whether cell–cell adhesions are altered by FGF signalling, immunofluorescent detection of E-cadherin was carried out in anterior endoderm (left), comparing embryos electroporated with GFP or FGF8-IRES-GFP, and in the posterior endoderm (right), comparing embryos electroporated with GFP or dnFGFR1-IRES-GFP. No overt differences in E-cadherin were observed, with localization along the basolateral boundaries of cells in the posterior and anterior endoderm, independent of FGF activity (in 3 of 3 embryos per condition). Changes in cell morphology are apparent, with FGF8-expressing cells in the anterior endoderm adopting a columnar epithelial morphology similar to that in the wild-type posterior endoderm, and dnFGFR1-expressing posterior endoderm cells becoming flattened, similar to cells in the wild-type anterior endoderm. Scale bar, 10 µm. j–m, Owing to redundancy between FGF4 and FG8 in other contexts (see Supplementary Discussion), we tested whether FGF4 and FGF8 act similarly in control of the hindgut-forming endoderm. j, Fluorescent section in situ hybridization for FGF4 in sagittal sections through the posterior HH15 embryo shows a posterior-to-anterior gradient similar to that of FGF8 (in 3 of 3 embryos). Scale bar, 100 µm. Whole-mount HH18 embryos following electroporation of endoderm at HH12 with GFP (k; 0 of 4 affected) or FGF4-IRES-GFP (l; 5 of 7 affected) reveals failure of endoderm to internalize and form hindgut upon misexpression of FGF4, phenocopying effects of FGF8 misexpression. Arrowhead, CIP. Scale bar, 100 µm. m, Relative tension in HH15 endoderm following electroporation of endoderm with GFP or FGF4-IRES-GFP (n = 5 embryos per group) reveals that FGF4, much like FGF8, modulates tension in the posterior endoderm: misexpression eliminates the tensional gradient by elevating tension anteriorly. Data are mean ± s.d.; two-way ANOVA with Tukey’s correction; *P < 0.05 versus GFP at same antero-posterior level.

Source data

Extended Data Fig. 9 DUSP6 reporter reveals FGF signalling gradient in posterior endoderm.

a–c, DUSP6–mScarlet (mSca) reporter co-electroporated into endoderm with nTagBFP control plasmid in embryos treated with 0.1% DMSO (a) or SU5402 (b) reveals loss of reporter activity with SU5402 treatment (n = 4 embryos). As a result, the FGF signalling gradient was flattened, indicated by quantification of the model and shape parameter ι (c). Data are mean ± s.d.; unpaired two-tailed t-test. d, Quantification of DUSP6 reporter-driven mSca activity (red, left y axis), electroporation control nTagBFP signal (blue, left y axis), and the normalized mSca/BFP (black, right y axis) from a single representative embryo (replicates in e) as a function of antero-posterior distance from the CIP. e, Normalized reporter activity profiles for six wild-type embryos; each embryo denoted by different colour. f, To test sensitivity of the DUSP6 reporter to subtle changes in FGF activity (as opposed to the marked effects of FGF ligand misexpression), heparinase I treatment was used to experimentally broaden the gradient. Because FGF ligands are tightly bound by heparin sulfate proteoglycans (HSPGs), treatment to degrade HSPGs would be expected to effectively increase the diffusion coefficient of FGF ligands, resulting in a broadening of the gradient. This effect was confirmed by quantification of the dimensionless FGF-gradient shape parameter ι, as measured by the DUSP6 reporter, following treatment with PBS (n = 7 embryos) or heparinase I at 0.1 U ml−1 (n = 4 embryos) or 1 U ml−1 (n = 6 embryos). Heparinase caused a dose-dependent increase in ι. Data are mean ± s.d.;one-way ANOVA with Tukey’s correction; *P < 0.05 versus PBS.

Source data

Extended Data Fig. 10 Parametric evaluation of cell-movement efficiency.

Dependence of cell-movement efficiency \(\left(\frac{{\bar{u}}_{{\rm{\max }}}}{\Lambda }\right)\) on FGF8 transport parameter ι (a) and matrix stiffness parameter κ (b). Green and red circles indicate experimental values for ι following GFP or FGF8 misexpression, respectively.

Source data

Supplementary information

Supplementary Information

This file contains supplementary methods, Supplementary Discussion, supplementary video. 1-7.

Reporting Summary

Video 1

Live imaging of cell movements in the posterior endoderm from HH14 to HH16 during hindgut formation. Live imaging of cell movements in the posterior endoderm from HH14 to HH16 during hindgut formation (Nerurkar Tabin REV Movie 1.mov).

Video 2

Cell movements are intrinsic to the endoderm. (Nerurkar Tabin REV Movie 2.mov).

Video 3

High magnification time lapse reveal collective movement of endoderm cells during hindgut tube formation. (Nerurkar Tabin REV Movie 3.mov).

Video 4

Pharmacologic modulation of actomyosin contractility disrupts hindgut-forming endoderm cell movements. (Nerurkar Tabin REV Movie 4.mov).

Video 5

Pharmacologic inhibition of FGF signaling disrupts posterior cell movements in the endoderm. (Nerurkar Tabin REV Movie 5.mov).

Video 6

Endoderm-specific inhibition and activation of FGF signaling similarly diminish posterior cell movements in the endoderm. (Nerurkar Tabin REV Movie 6.mov).

Video 7

Model simulation of contractile gradient-based cell movements in response to a spatial gradient of FGF8. (Nerurkar Tabin REV Movie 7.mov).

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Nerurkar, N.L., Lee, C., Mahadevan, L. et al. Molecular control of macroscopic forces drives formation of the vertebrate hindgut. Nature 565, 480–484 (2019). https://doi.org/10.1038/s41586-018-0865-9

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  • DOI: https://doi.org/10.1038/s41586-018-0865-9

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