Optogenetic inhibition of actomyosin reveals mechanical bistability of the mesoderm epithelium during Drosophila mesoderm invagination

  1. Hanqing Guo
  2. Michael Swan
  3. Bing He  Is a corresponding author
  1. Department of Biological Sciences, Dartmouth College, United States
  2. Department of Molecular Biology, Princeton University, United States

Abstract

Apical constriction driven by actin and non-muscle myosin II (actomyosin) provides a well-conserved mechanism to mediate epithelial folding. It remains unclear how contractile forces near the apical surface of a cell sheet drive out-of-the-plane bending of the sheet and whether myosin contractility is required throughout folding. By optogenetic-mediated acute inhibition of actomyosin, we find that during Drosophila mesoderm invagination, actomyosin contractility is critical to prevent tissue relaxation during the early, ‘priming’ stage of folding but is dispensable for the actual folding step after the tissue passes through a stereotyped transitional configuration. This binary response suggests that Drosophila mesoderm is mechanically bistable during gastrulation. Computer modeling analysis demonstrates that the binary tissue response to actomyosin inhibition can be recapitulated in the simulated epithelium that undergoes buckling-like deformation jointly mediated by apical constriction in the mesoderm and in-plane compression generated by apicobasal shrinkage of the surrounding ectoderm. Interestingly, comparison between wild-type and snail mutants that fail to specify the mesoderm demonstrates that the lateral ectoderm undergoes apicobasal shrinkage during gastrulation independently of mesoderm invagination. We propose that Drosophila mesoderm invagination is achieved through an interplay between local apical constriction and mechanical bistability of the epithelium that facilitates epithelial buckling.

Editor's evaluation

The authors examine the process of mesoderm invagination in the Drosphila embryo and found that while myosin contractility is critical to prevent tissue relaxation during the early phase of the process, it is dispensable for the subsequent folding step. Through modeling and experimental analyses, the authors find that folding is likely mediated by a joint action of active cell shape changes in the mesoderm and apico-basal shrinking in the surrounding ectoderm and suggest that the mesoderm behave as a mechanically bistable tissue during gastrulation.

https://doi.org/10.7554/eLife.69082.sa0

Introduction

Contractile forces generated by actin and myosin (actomyosin) networks are widely employed in embryogenesis to drive cell motility and cell shape change that pattern epithelial tissues (Martin and Goldstein, 2014; Munjal and Lecuit, 2014). During epithelial folding mediated by apical constriction, the actomyosin network in epithelial cells constricts cell apices, which leads to bending of the cell sheet (Sawyer et al., 2010). It remains unclear how ‘in-plane’ contractile forces generated at the apical surface drive ‘out-of-the-plane’ folding of the tissue. The invagination of the presumptive mesoderm during Drosophila gastrulation is a well-characterized epithelial folding process mediated by apical constriction (reviewed in Gilmour et al., 2017; Martin, 2020; Gheisari et al., 2020). During gastrulation, ventrally localized mesoderm precursor cells constrict their cell apices and become internalized into a ventral furrow. The folding process occurs in two steps (Leptin and Grunewald, 1990; Sweeton et al., 1991). During the first 10–12 min, the ventral cells undergo apical constriction and elongate in the apical-basal direction, while the cell apices remain near the surface of the embryo (the lengthening phase). In the next 6–8 min, the ventral cells rapidly internalize as they shorten back to a wedge-like morphology (the shortening phase) (Figure 1a). Ventral furrow formation is controlled by the dorsal-ventral (DV) patterning system. The maternally deposited morphogen Dorsal controls the expression of two transcription factors, Twist and Snail, in the presumptive mesoderm. Twist and Snail in turn cause myosin activation at the apical pole of ventral mesodermal cells through a sequential action of G-protein coupled receptor (GPCR) signaling (Leptin, 1991; Parks and Wieschaus, 1991; Costa et al., 1994; Kölsch et al., 2007; Manning et al., 2013; Kerridge et al., 2016) and RhoGEF2-Rho1-Rho associated kinase (Rok) pathway (Barrett et al., 1997; Häcker and Perrimon, 1998; Nikolaidou and Barrett, 2004; Dawes-Hoang et al., 2005; Martin et al., 2009; Mason et al., 2013). Activated myosin forms a supracellular actomyosin network across the apical surface of the prospective mesoderm and drives the constriction of cell apices (Martin et al., 2010; Martin et al., 2009; Martin and Goldstein, 2014).

Figure 1 with 3 supplements see all
Light-dependent membrane recruitment of CRY2-Rho1DN results in rapid myosin inactivation.

(a) Drosophila mesoderm invagination occurs in distinct lengthening and shortening phases. Ventral side is up (same for all cross-section images in this work unless otherwise stated). A single ventral cell undergoing apical constriction is outlined in green. In this study, we sought to test whether myosin contractility is required throughout the folding process by acute, stage-specific inhibition of Rho1 (arrowheads). (b) Cartoon depicting the principle of the optogenetic tool used in this study. Upon blue light stimulation, CRY2-Rho1DN is translocated from the cytosol to the plasma membrane through the interaction between CRY2 and membrane anchored CIBN. (c) Confocal images showing the rapid membrane recruitment of CRY2-Rho1DN upon blue light illumination. (d) Relative abundance of membrane recruited CRY2-Rho1DN over time after blue light stimulation. Error bar: s.d., N=6 embryos. (e) A wild-type embryo expressing Sqh-GFP showing apical myosin accumulation during ventral furrow formation. N=4 embryos. (f) Activation of Opto-Rho1DN results in rapid dissociation of myosin from the ventral cell cortex (arrow) in a gastrulating embryo. N=8 embryos. (g) Confocal images showing the confined membrane recruitment of CRY2-Rho1DN within a region of interest (ROI, red box) that has been scanned by a focused beam of blue laser. N=6 embryos. All scale bars=20 μm.

The essential role of apical constriction in ventral furrow formation has been well demonstrated. Genetic mutations or pharmacological treatments that inhibit myosin activity disrupt apical constriction and result in failure in ventral furrow formation (reviewed in Gheisari et al., 2020; Martin, 2020). Biophysical studies show that cell shape change occurred during the lengthening phase is a direct, viscous response of the tissue interior to apical constriction (Gelbart et al., 2012; He et al., 2014). However, accumulating evidence suggests that apical constriction does not directly drive invagination during the shortening phase. First, it has been observed that the maximal rate of apical constriction (or cell lengthening) and the maximal rate of tissue invagination occur at distinct times (Polyakov et al., 2014; Rauzi et al., 2015), and timed injection of Rok inhibitor indicates that the late stage of ventral furrow invagination is less sensitive to myosin inactivation (Krajcovic and Minden, 2012). Second, it has been previously proposed, and more recently experimentally demonstrated, that myosin accumulated at the lateral membranes of constricting cells (lateral myosin) facilitates furrow invagination by exerting tension along the apical-basal axis of the cell (Brodland et al., 2010; Conte et al., 2012; Gracia et al., 2019; John and Rauzi, 2021). Finally, a number of computational models predict that mesoderm invagination requires additional mechanical input from outside of the mesoderm, such as ‘pushing’ forces from the surrounding ectodermal tissue (Muñoz et al., 2007; Conte et al., 2009; Allena et al., 2010; Brodland et al., 2010). These models are in line with the finding that blocking the movement of the lateral ectoderm by laser cauterization inhibits mesoderm invagination (Rauzi et al., 2015). A similar disruption of ventral furrow formation can also be achieved by increasing actomyosin contractility in the lateral ectoderm (Perez-Mockus et al., 2017). While these pioneer studies highlight the importance of cross-tissue coordination during mesoderm invagination, the actual mechanical mechanism that drives the folding of the mesodermal epithelium and the potential role of the surrounding ectodermal tissue remain to be elucidated.

In this study, we investigate the mechanics of ventral furrow formation by asking whether actomyosin contractility is required throughout the folding process. By developing an optogenetic tool to acutely inhibit actomyosin activity, we find that the dependence of furrow invagination on actomyosin contractility is strongly stage-dependent: Inhibition of actomyosin during apical constriction results in immediate relaxation of the constricted tissue, whereas similar treatment after a stereotyped transitional stage does not impede invagination, suggesting that the mesoderm epithelium has two stable mechanical status during gastrulation. The binary tissue response to actomyosin inhibition can be recapitulated by a 2D vertex model that combines apical constriction in the mesoderm and apicobasal shortening in the neighboring ectoderm, which generates in-plane compression due to volume conservation. Finally, we show evidence the lateral ectoderm undergoes apicobasal shortening around the transitional stage of ventral furrow formation. This ectodermal shortening process, as well as the associated ventrally directed movement of the ectoderm, could occur independently of mesoderm invagination. Taken together, we propose that Drosophila mesoderm epithelium is mechanically bistable during gastrulation. We further hypothesize that the mechanical bistability of the mesoderm is attributed to ectodermal compression and functions together with active cell shape change in the mesoderm to facilitate mesoderm invagination.

Results

Plasma membrane recruitment of Rho1DN results in rapid loss of apical myosin and F-actin network

In order to acutely inhibit myosin activity in live embryos (Figure 1a), we generated an optogenetic tool, ‘Opto-Rho1DN,’ to inhibit Rho1 through light-dependent plasma membrane recruitment of a dominant negative form of Rho1 (Rho1DN). Like other Rho GTPases, Rho1 cycles between an active, GTP-bound state and an inactive, GDP-bound state (Etienne-Manneville and Hall, 2002). Rho1DN bears a T19N mutation, which abolishes the ability of the mutant protein to exchange GDP for GTP and thereby locks the protein in the inactive state (Barrett et al., 1997). A second mutation in Rho1DN, C189Y, abolishes its naive membrane targeting signal (Sebti and Der, 2003; Roberts et al., 2008). When Rho1DN is recruited to the plasma membrane, it binds to and sequesters Rho1 GEFs, thereby preventing activation of endogenous Rho1 as well as Rho1-Rok-mediated activation of myosin (Barrett et al., 1997; Feig and Cooper, 1988).

Opto-Rho1DN is based on the CRY2-CIB light-sensitive dimerization system (Liu et al., 2008; Kennedy et al., 2010; Guglielmi et al., 2015; Figure 1b). The tool contains two components: a GFP-tagged N-terminal of CIB1 protein (CIBN) anchored at the cytoplasmic leaflet of the plasma membrane (CIBN-pmGFP) (Guglielmi et al., 2015), and a fusion protein with Rho1DN, a mCherry tag, and a blue light-reactive protein Cryptochrome 2 (CRY2-Rho1DN). CRY2-Rho1DN remained in the cytoplasm when the sample was kept in the dark or imaged with a yellow-green (561 nm) laser (Figure 1c, T=0). Upon blue (488 nm) laser stimulation, CRY2 undergoes a conformational change and binds to CIBN (Kennedy et al., 2010; Liu et al., 2008), resulting in rapid recruitment of CRY2-Rho1DN to the plasma membrane (Figure 1c and d; Figure 1—video 1). The average time for half-maximal membrane recruitment of CRY2-Rho1DN is 4 s (3.9±2.8 s, mean±s.d., n=5 embryos). Embryos expressing CIBN-pmGFP and CRY2-Rho1DN developed normally in a dark environment and could hatch (100%, n=63 embryos). In contrast, most embryos (>85%, n=70 embryos) kept in ambient room light failed to hatch, reflecting the essential role of Rho1 in Drosophila embryogenesis (Barrett et al., 1997; Magie et al., 1999; Johndrow et al., 2004).

The effectiveness of Opto-Rho1DN on myosin inactivation was validated by stimulating embryos during apical constriction, which resulted in diminishing of apical myosin signal within 60 s (Figure 1e and f; Figure 1—video 2). Rapid disappearance of cortical myosin upon Rho1 inhibition is consistent with the notion that Rho1 and myosin cycle quickly through active and inactive statues due to the activity of GTPase activating proteins (GAPs) for Rho1 and myosin light chain phosphatases, respectively (Munjal et al., 2015; Coravos and Martin, 2016). Our data also provide direct evidence that sustained myosin activity during ventral furrow formation requires persistent activation of Rho1.

In addition to myosin, we found that acute Rho1 inhibition also affects cortical F-actin networks during apical constriction, presumably through inactivation of the Rho1 effector formin Diaphanous (Johndrow et al., 2004; Homem and Peifer, 2008; Mason et al., 2013; Coravos and Martin, 2016). In unstimulated embryo, F-actin is enriched at the apical domain of the constricting cells (Figure 2a, red arrowheads and cyan arrows) and along the lateral membrane in both constricting and non-constricting cells (Figure 2a, yellow arrows and magenta arrows). Apical F-actin disappeared within 1.2 min after Opto-Rho1DN stimulation (Figure 2b). Lateral F-actin in the constricting cells was not immediately affected and only appeared to diminish 4 min after stimulation (Figure 2b). In contrast, lateral F-actin in the ectodermal cells was not significantly affected (Figure 2b). Taken together, our results indicate that Opto-Rho1DN is an effective tool for spatially- and temporally-confined inactivation of apical actomyosin contractility through simultaneous myosin inactivation and actin disassembly.

Rapid disassembly of apical F-actin in the constricting cells after Opto-Rho1DN stimulation.

(a) Cross-sections (left) and en face views (right) of a representative unstimulated embryo showing the localization of the F-actin marker UtrophinABD-Venus (UtrABD-Venus) during ventral furrow formation. At the onset of apical constriction, F-actin is enriched along the lateral membrane in both mesodermal and ectodermal cells (yellow and magenta arrows, respectively). During apical constriction, F-actin also accumulates at the apical domain of the constricting cells (cyan arrows). T=0 (min) is the onset of ventral furrow formation. +0 μm indicates the apex of the ventral most cells (red arrowheads) where the strongest accumulation of apical F-actin is observed. (b) Upon Opto-Rho1DN stimulation (immediately before 6.2 min), apical F-actin disappears within 1.2 min (cyan arrows). Lateral F-actin in the constricting cells was not immediately affected and only started to diminish 4 min after stimulation (yellow arrows). Lateral F-actin in the ectodermal cells was not significantly affected (magenta arrows). Note that the initial apical indentation (red arrowheads) quickly disappeared after stimulation upon relaxation of the cell apex. N=3 for unstimulated embryos and N=10 for stimulated embryos. Scale bars: cross-sections, 20 μm; en face views, 10 μm.

Consistent with the actomyosin phenotype, Opto-Rho1DN stimulation before or during apical constriction resulted in prevention or reversal of apical constriction, respectively (Figure 1—figure supplement 1). In addition, Opto-Rho1DN stimulation resulted in an immediate loss of cortical tension at the ventral surface of the embryo during apical constriction (Figure 3), in accordance with the previous finding that the increase in tissue tension in the ventral mesodermal region during apical constriction is due to activation of apical myosin contractility and integration of contractile forces across the constriction domain (Martin et al., 2010). Finally, by stimulating the embryo within a specific region of interest (ROI), we showed that membrane recruitment of CRY2-Rho1DN and inhibition of apical constriction are restricted to cells within the stimulated region (Figure 1g, Figure 1—figure supplement 1c, d).

Opto-Rho1DN stimulation during apical constriction results in an immediate loss of cortical tension at ventral surface of the embryo.

(a) Cartoon depicting the experimental setup for laser ablation to detect cortical tension. Yellow shaded regions indicate the ablated regions. For stimulated embryos, light-activation of Opto-Rho1DN was performed 3 min before the laser ablation. Due to apical relaxation after stimulation, multiple z-planes were ablated (yellow shaded region) in order to ensure the ablation of the very apical surface of the ventral cells. (b) Width changes of the ablated region along the A-P axis during the first 20 s after laser ablation. A clear tissue recoil was observed after laser cutting in the unstimulated control embryos. In contrast, little to no tissue recoil was observed in the stimulated embryos, indicating lack of apical tension after Rho1 inhibition. p value was calculated using two-sided Wilcoxon rank-sum test. (c–d) Kymographs showing the comparison between unstimulated and stimulated embryos. No obvious tissue recoil was observed in the stimulated embryos (N=6 for unstimulated embryos and N=5 for stimulated embryos). Dotted line indicates ablation site.

Acute inhibition of myosin contractility reveals mechanical bistability of the mesoderm during gastrulation

Using Opto-Rho1DN, we determined the role of actomyosin contractility at different stages of ventral furrow formation. We first obtained non-stimulated control movies by imaging embryos bearing Opto-Rho1DN on a multiphoton microscope using a 1040-nm pulsed laser, which excites mCherry but does not stimulate CRY2 (Guglielmi et al., 2015). Without stimulation, the embryos underwent normal ventral furrow formation (Figure 4a). The transition from the lengthening phase to the shortening phase (TL-S trans) occurred around 10 min after the onset of gastrulation (9.3±1.6 min, mean±s.d., n=10 embryos), which was similar to the wild-type embryos (8.9±0.8 min, mean±s.d., n=6 embryos) (Figure 4—figure supplement 1a-c). This transition was characterized by a rapid inward movement of the apically constricted cells and could be readily appreciated by the time evolution of distance between the cell apices and the vitelline membrane of the eggshell (invagination depth ‘D,’ Figure 4—figure supplement 1d).

Figure 4 with 4 supplements see all
Acute inhibition of actomyosin contractility results in stage-dependent response during ventral furrow formation.

(a) Still images from multiphoton movies showing different tissue responses to acute loss of myosin contractility during ventral furrow formation. Early Group, Mid Group, and Late Group embryos (N=8, 4, and 6 embryos, respectively) are defined based on their immediate response to myosin inhibition. For stimulated embryos, the first frame corresponds to the time point immediately after stimulation. The inset depicts the stimulation and imaging protocol. Arrowheads indicate the apex of the ventral most cells. (b) Time evolution of the invagination depth ‘D’ for the stimulated embryos and a representative unstimulated control embryo. For stimulated embryos, all movies were aligned in time to the representative control embryo based on furrow morphology at the time of stimulation. (c) Relationship between the transition phase for sensitivity to myosin inhibition (Ttrans) and lengthening-shortening transition (TL-S trans). (d) Scatter plot showing the relation between invagination depth at the time of stimulation (Ds) and the delay time (Tdelay) in furrow invagination compared to the representative control embryo. Tdelay is highly sensitive to Ds, with a switch-like change at Ds~6 μm (dashed line). Inset: Average Tdelay in Early (E, n=5 embryos that invaginated), Mid (M, n=4), and Late (L, n=6) Group embryos. Statistical comparisons were performed using two-sided Wilcoxon rank-sum test. (e) Cartoon depicting mechanical bistability of the mesoderm during gastrulation. Both the initial, pre-constriction state and the final, fully invaginated state are stable. During gastrulation, actomyosin contractility is critical for bringing the system from the initial state to an intermediate, transitional state, whereas the subsequent transition to the final state can occur independent of myosin contractility.

Figure 4—source data 1

Cell length measurements for determining lengthening-shortening transition time.

https://cdn.elifesciences.org/articles/69082/elife-69082-fig4-data1-v2.xlsx

Next, we examined the effect of Opto-Rho1DN stimulation on furrow invagination (Figure 4b; Figure 4—figure supplement 1e; see Materials and methods). Measuring the rate of furrow invagination (dD/dt) immediately after stimulation revealed three types of tissue responses to stimulation (Figure 4a and b; Figure 4—figure supplement 1f, g; Figure 4—video 1). For most embryos that were stimulated before T=7 min (‘Early Group’ embryos), the constricting cells relaxed immediately. The initial apical indentation disappeared (dD/dt<0). The flanking, non-constricting cells, which were previously apically stretched by neighboring constricting cells, shrunk back, indicating that the apical cell cortex was elastic (Figure 4—figure supplement 2a, b). In contrast, for most embryos that were stimulated after T=7 min (‘Late Group’ embryos), furrow invagination proceeded at a normal rate and dD/dt remained positive. Finally, some embryos stimulated around T=7 min (6.5±1.4 min, mean±s.d., n=4 embryos; ‘Mid Group’ embryos) displayed an intermediate phenotype, where ventral furrow invagination paused for around 5 min (4.7±0.5 min; dD/dt≈0) before invagination resumed. We confirmed that there was no residual apical myosin in Late Group embryos upon stimulation (Figure 4—figure supplement 3a). Furthermore, lateral myosin, which has been shown to facilitate furrow invagination (Gracia et al., 2019; John and Rauzi, 2021), also quickly disappeared after stimulation (Figure 4—figure supplement 3b, c). Overall, furrow invagination in Late Group embryos is not due to myosin activities at the apical or lateral cortices of the mesodermal cells. Taken together, these results indicate that actomyosin contractility is required in the early but not the late phase of furrow formation. The narrow time window when Mid Group embryos were stimulated defined the ‘transition phase’ for sensitivity to myosin inhibition (Ttrans), which is immediately before TL-S trans (Figure 4c).

While all Early Group embryos showed tissue relaxation after stimulation, five out of eight of them were able to invaginate after a prolonged delay (Figure 4b; Figure 4—figure supplement 2c-f). The invagination was associated with modest myosin reaccumulation at the cell apices that occurred approximately 10 min after the initial stimulation (Figure 4—figure supplement 2e-f). It is unclear why apical myosin sometimes reoccurred in the presence of persistent membrane localization of CRY2-Rho1DN. Nevertheless, the completion of furrow invagination when apical constriction is partially impaired is consistent with previous genetic studies and suggests that invagination is facilitated by additional mechanical input (Costa et al., 1994; Parks and Wieschaus, 1991).

Because the response of ventral furrow formation to myosin inhibition appears to be binary, we sought to identify morphological features of the progressing furrow that associate with Ttrans. Hence, we examined the relationship between D immediately after stimulation (Ds) and the extent by which furrow invagination is affected (assessed by the delay time in furrow invagination, Tdelay; see Materials and methods). We found that for stimulated embryos, Tdelay was ultrasensitive to Ds, with a steep transition at around 6 μm (Figure 4d). When Ds was above the threshold, Tdelay was close to 0. When Ds was below the threshold, Tdelay increased to 10 min (9.4±1.4 min, mean±s.d., n=5 Early Group embryos) or longer (fail to invaginate, n=3 Early Group embryos). Therefore, the response to acute myosin inhibition can be well predicted by the tissue morphology associated with the stage of furrow progression.

The binary response of the embryo to acute myosin inhibition led us to propose that the mesoderm during gastrulation is mechanically bistable. In this model, the mesoderm epithelium has two stable equilibrium states: the initial, pre-constricted conformation and the final, fully invaginated conformation (Figure 4e). Energy input from actomyosin-mediated apical constriction drives the transition from the initial state to an intermediate state (Ttrans), which has the propensity to transition into the final state without further requirement of actomyosin contractility. If myosin is inhibited before Ttrans, the furrow would relax back to the initial state. In contrast, if myosin is inhibited after Ttrans, the furrow would continue to invaginate without pause (Figure 4e).

Analysis of three-dimensional cell shape change after Opto-Rho1DN stimulation

To gain further insights into how cells respond to rapid myosin loss after Rho1 inhibition, we performed three-dimensional cell reconstruction using ilastik (Berg et al., 2019). For each embryo, we examined a single row of cells (cells 1–11) along the medial-lateral axis that include the ventral mesodermal cells (cells ~1–6, ‘constricting cells’), the lateral mesodermal cells (cells ~7–9, ‘non-constricting flanking cells’), and presumably some ectodermal cells (cells ~10 and 11) (Figure 5a and f). Our analysis covered time points with most notable cell shape changes after stimulation (0, 2.2, and 4.4 min for Early Group embryos, and 0, 1.1, and 2.2 min for Late Group embryos). Shorter time windows were used for Late group embryos because of the rapid furrow invagination despite the inhibition of actomyosin contractility.

Figure 5 with 1 supplement see all
Three-dimensional segmentation reveals details of cell response after Opto-Rho1DN stimulation.

(a–e) Early Group embryos and (f–j) Late Group embryos. (a, f) A row of cells along the medial-lateral axis on one side of the ventral midline was segmented in 3D (green box) over time after stimulation. Cells 1–6: constricting cells; Cells 7–9: flanking cells; Cells 10 and 11: ectodermal cells. Early Group: T=0, 2.2, and 4.4 min. Late Group: T=0, 1.1, and 2.2 min. 3D rendering of three representative cells, cell 1 (mid constricting cell), cell 6 (side constricting cell), and cell 9 (flanking cell), are shown in magenta, orange, and cyan boxes, respectively. These cells are also marked on the cross-section views (outlines) and the 3D tissue renderings (arrowheads) with the same color code. D: invagination depth. (b, g) Cartoon depicting apical surface area change (top) and characteristic cell shape changes (bottom) observed in Early Group (b) and Late Group (g) embryos. (c–e, h–j) Upper panels: quantifications showing the apical surface area change (c, h), cell length change (d, i), and cell curvature change (e, j) between 0 and 4.4 min (c–e) or between 0 and 2.2 min (h–j). Measurements from both Early and Late Groups are shown in (h-j) for comparison. Bottom panels show the measurements for each quantity immediately after stimulation (T=0). Error bars: ±s.e. N=3 embryos for each group.

As expected, in Early Group embryos, there was a prompt apical area relaxation in the constricting cells and an accompanying apical area reduction in the flanking cells following stimulation (see Figure 5b for a cartoon depiction). A number of features were observed in Early Group embryos. First, there was no substantial cell volume change (±5%, Figure 5—figure supplement 1), similar to the observation for normal ventral furrow formation (Gelbart et al., 2012; Polyakov et al., 2014). Second, the flanking cells that were most apically stretched before stimulation underwent most prominent apical area reduction, which is anticipated (comparing cells 7–9, Figure 5c). Surprisingly, the extent of apical relaxation in the constriction domain was not uniform. In general, cells located at the side of the constriction domain (‘side constricting cells,’ cells 4–6) relaxed more than those located closer to the ventral midline (‘mid constricting cells,’ cells 1–3) (Figure 5c). The lower degree of apical relaxation in the mid-constriction cells was accompanied with less prominent ‘reversal of lengthening’ (Figure 5d, see Figure 5b for a cartoon depiction). Finally, the alteration in apical cell area was associated with changes in cell curvature along the apical-basal axis. Before stimulation, the flanking cells and the side constricting cells were all tilted, with their apical side bending toward the ventral midline (e.g., cyan and orange cells in Figure 5a). After stimulation, the cells straightened up and partially restored their initial, columnar cell shape. This cell shape change is associated with a reduction in the apical-basal cell length and lateral cell area (Figure 5b, d and e; Figure 5—figure supplement 1). The ectodermal cells also showed some unbending, but their shape changes were in general moderate (Figure 5, cells 10 and 11).

Interestingly, despite the drastic tissue level difference between Early and Late groups of embryos, several trends we observed in Early group was also present in Late group, such as the conservation of cell volume (Figure 5—figure supplement 1), the straighten-up of some (but not all) flanking cells (e.g., cell nine in Figure 5j), and the uneven pattern of apical relaxation in the constriction domain (Figure 5h). Despite these similarities, Late Group embryos displayed several prominent features that were distinct from Early Group embryos (see Figure 5b and g for a cartoon depiction). First, the mid constricting cells barely showed any apical relaxation and underwent rapid cell shortening after stimulation (Figure 5h and i, cells 1–3), which were not observed in Early Group embryos but rather resembled the normal cell shortening process (Polyakov et al., 2014). Second, despite undergoing apical relaxation, the side constricting cells bent further toward the ventral midline instead of straightening up as their counterparts in Early Group embryos (Figure 5j, cells 4–6). The behavior of the middle and side constricting cells in Late Group embryos elucidates the cellular basis for the continued deepening of the furrow after Opto-Rho1DN stimulation and explains the tissue level difference between Early and Late Group embryos. Importantly, the observed cell shape change and tissue invagination in Late Group embryos occurred in the absence of actomyosin contractility, suggesting the presence of additional mechanical contributions.

A vertex model combining apical constriction in the mesoderm and apicobasal shortening in the ectoderm recapitulates the binary response to actomyosin inhibition

What mechanism may account for the apparent mechanical bistability during mesoderm folding? One possible mechanism is germband extension, a body axis elongation process in Drosophila that occurs soon after the onset of gastrulation (Kong et al., 2017). During germband extension, the ventral and lateral ectoderm undergo convergent extension and converge toward the ventral midline as they extend along the anterior-posterior (AP) axis. If germband extension-mediated ventral movement starts around Ttrans, it may account for the binary tissue response to actomyosin inhibition. However, previous literature indicates that germband extension does not start until ventral furrow formation is nearly completed (Lye et al., 2015). Furthermore, germband extension depends on Rho1-mediated activation of actomyosin contractility in the ectodermal cells (Kong et al., 2017). Since we stimulated Opto-Rho1DN in both the mesoderm and the ectoderm in our optogenetic experiment, we anticipate that germband extension would be inhibited. Consistent with this notion, we did not observe AP tissue movement in Opto-Rho1DN treated embryos (Figure 6—figure supplement 2), suggesting that other mechanisms are responsible for the invagination of ventral furrow in Late Group embryos.

Previous biophysical studies demonstrate that the embryonic epithelium in early Drosophila embryos can display an elastic response with a decay timescale of at least 4 min (Doubrovinski et al., 2017). The apparent bistable characteristic of the mesoderm is reminiscent of a compressed elastic beam undergoing snap-through buckling facilitated by a transverse ‘indentation force’ (Figure 6b; Qiu et al., 2004). Based on this analogy, we hypothesized that mesoderm invagination is mediated through tissue buckling enabled by in-plane compressive stresses. We tested this hypothesis by implementing compressive stresses from the ectoderm in a previously published two-dimensional vertex model for ventral furrow formation (Polyakov et al., 2014; Figure 6a; see Materials and methods). In this model, the sole energy input is apical constriction of the ventral cells, whereas the apical, lateral, and basal cortex of cells were represented by elastic springs that resist deformation (Polyakov et al., 2014; see Materials and methods). Note that the elasticity assumption in this model is a simplification of the actual viscoelastic properties of the embryonic tissue. However, this model successfully predicted the intermediate and final furrow morphologies with a minimal set of active and passive forces without prescribing individual cell shape changes (Polyakov et al., 2014; see Materials and methods). It is therefore advantageous to use this model to explore the main novel aspect of the folding mechanics underlying ventral furrow formation and to test the central concept of our hypothesis based on relatively small number of assumptions.

Figure 6 with 2 supplements see all
Mechanical bistability of the mesoderm during gastrulation can arise from ectoderm-derived compressive stresses.

(a) 2D vertex model testing the mechanisms of mechanical bistability of the mesoderm during gastrulation. In this model, the apical, lateral, and basal cortices of cells are modeled as elastic springs that resist deformations, the cell interiors are non-compressible, and the only two active energy inputs are apical constriction in the mesoderm (blue cells) and in-plane compression generated by the ectoderm (white cells). (b) Buckling of a compressed elastic beam triggered by a transverse indentation force. (c) The model recapitulates the bipartite response to acute loss of actomyosin contractility only when the in-plane compression is present. Column 1 shows normal ventral furrow formation. Columns 2–4 show tissue response when apical constriction is inhibited at different stages of ventral furrow formation with or without ectodermal compression. (d) Model predictions on the impact of reducing the width of apical constriction domain or the uniformity of apical constriction on final invagination depth. In the presence of compressive stress, final invagination depth becomes less sensitive to perturbations of apical constriction.

Figure 6—source data 1

Computer code for the energy minimization-based vertex model for ventral furrow formation.

https://cdn.elifesciences.org/articles/69082/elife-69082-fig6-data1-v2.zip

Using this model, we simulated the effect of acute myosin loss by removing the apical contractile forces at different intermediate states of the model (Materials and methods). Remarkably, in the presence of ectodermal compression, the model recapitulated the binary response to loss of apical constriction as observed experimentally (Figure 6c). In particular, the transitional state of the tissue revealed in the simulation is visually similar to that identified in our optogenetic experiments. As a control, in the absence of ectodermal compression, the model always returned to its initial configuration regardless of the stage when apical constriction was eliminated (Figure 6c). Therefore, our simulation indicates that buckling of an elastic cell sheet jointly mediated by local apical constriction and global in-plane compression can well reproduce the observed binary tissue response to acute actomyosin inhibition. Our model also predicts that the presence of ectodermal compression could compensate for partial impairment of apical constriction (Figure 6d), which explains why some Early Group embryos could still invaginate when myosin activity was greatly downregulated.

Interestingly, our simulation predicts that for embryos stimulated after Ttrans, the ventral furrow at the final invaginated state would be narrower with fewer cells being incorporated into the furrow compared to control embryos. Our observation in Late Group embryos well matched this prediction (Figure 6c; Figure 6—figure supplement 1a, b). Consistent with our 3D analysis (Figure 5h), we found that stimulation in Late Group embryos induced a mild but noticeable alteration in tissue flow that indicates the occurrence of apical relaxation in the constricted cells (Figure 6—figure supplement 1c, d). These results suggest that maintaining actomyosin contractility after Ttrans helps to prevent local relaxation of the apical membranes, thereby allowing more cells to be incorporated into the furrow. This ‘late’ function of actomyosin contractility is dispensable for furrow invagination per se but is important for the complete internalization of the prospective mesoderm.

Presence of myosin-independent cell shortening force in the constriction domain is important for the vertex model to recapitulate the observed binary response

In the vertex model described above, shortening of the constricting cells is mediated by passive elastic restoration forces generated in these cells as their lateral edges are stretched during cell lengthening (Polyakov et al., 2014). This passive elastic force is not affected by in silico myosin inactivation. However, our experiments show that Opto-Rho1DN stimulation results in rapid diminishing of lateral myosin in the constricting cells, suggesting that Rho1 inhibition may weaken the shortening forces in these cells (Gracia et al., 2019; John and Rauzi, 2021). To account for this effect, we implemented active lateral constriction forces in the ventral cells. In such a scenario, the active and passive lateral forces worked in combination to mediate ventral cell shortening, but only the active force was sensitive to myosin inactivation (Materials and methods). The active and passive lateral forces were given by KL_activel and  KL_passive(l l0), respectively, where l and l0 were the current and the resting length of the lateral edge, respectively. As expected, the addition of the active lateral force allows us to reduce the passive lateral force while still generate furrows with normal morphology (Figure 7a, comparing column KL_active=0 with column KL_active=2). The implementation of myosin-dependent active lateral forces allowed us to examine how the binary response would be influenced if myosin inactivation weakens the cell shortening forces in the mesoderm.

Modeling analysis of the impact of myosin-dependent and myosin-independent cell shortening forces in the constriction domain on binary tissue response.

(a) The myosin-dependent, active lateral shortening forces and myosin-independent, passive lateral shortening forces function additively to mediate cell shortening and furrow invagination in the 2D vertex model. The active and passive lateral forces are given by KL_activel and KL_passive(l- l0), respectively, where l and l0 are the current and the resting length of the lateral edge, respectively. Green shaded region represents the conditions where the model generates normal final furrow morphology. (b) When KL_passive=20, changing KL_active has little impact on the binary response of the model to stimulation. (c) When KL_active=2, changing KL_passive does not affect the binary tissue response per se but influences the final morphology after stimulation.

We performed the tests under the conditions where the model generates normal furrow morphology (green shaded region in Figure 7a, KL_passive=20 while KL_active ranges from 0 to 2, or KL_active=2 while KL_passive ranges from 0.002 to 20). When KL_passive=20, changing KL_active has little impact on the response of the model to stimulation (Figure 7b). When KL_active=2, changing KL_passive did not affect the binary tissue response per se but influenced the final morphology after stimulation (Figure 7c). When the passive lateral force was relatively high (KL_passive= 20), the model displayed normal binary response with realistic final furrow morphologies. Lowering the passive lateral force by a factor of ten (KL_passive= 2) did not change the tissue response aside from a mild reduction of the invagination depth ‘D’ (from 61 μm to 51 μm) in Late Group embryos. Strikingly, further lowering KL_passive to 0.2 resulted in a very shallow furrow and abnormal constricting cell morphology in Late Group embryos (D=30 μm). Taken together, these results demonstrate that the binary response of the model to myosin inhibition still exists when the cell shortening forces in the mesoderm become sensitive to myosin inhibition. However, a minimal level of myosin-independent lateral elastic force (KL_passive>0.2) is required for the model to generate realistic final furrow morphology in Late Group embryos. Interestingly, we found that in real embryos the lateral actin in the constricting cells persisted for several minutes after Opto-Rho1DN stimulation (Figure 2b, yellow arrows), which might provide the lateral elasticity described in our model.

The lateral ectoderm undergoes apical-basal shortening during gastrulation independently of ventral furrow formation

It remains unclear whether compressive stresses exist in the ectoderm during ventral furrow formation in real embryos. No cell division happens in the embryo during ventral furrow formation (Foe, 1989; Edgar et al., 1994), thereby excluding a role for cell proliferation in generating compressive stresses as seen in many other tissues (Nelson, 2016). Interestingly, it has been reported that ectodermal cells undergo apical-basal shortening during gastrulation (Brodland et al., 2010). In theory, this cell shape change could generate compressive stresses in the planar direction if the cell volume remains conserved. To further investigate the extent of ectodermal shortening during gastrulation, we performed 3D tissue volume measurement (Volec) in the lateral ectodermal region of wild-type embryos (ROI: 60°–120° away from the ventral midline and 75-μm long along the AP axis; Figure 8a). We focused our analysis on the lateral ectodermal region since a previous study shows that the dorsal ectoderm does not significantly contribute to ventral furrow formation (Rauzi et al., 2015). We reasoned that if the ectodermal cells undergo apical-basal shortening, there should be a net volume outflux from the ROI, causing a reduction in Volec (Figure 8a). Since cell shortening may not be uniform across the tissue, the change in Volec provides a less noisy measure of the change in average tissue thickness. To measure Volec, we performed 3D tissue segmentation without segmenting individual cells (Materials and methods).

Figure 8 with 3 supplements see all
The lateral ectoderm undergoes apical-basal shortening during gastrulation independently of ventral furrow formation.

(a) Top: cartoon showing a cross-section view of an embryo. Blue: mesoderm. Yellow: segmented ectoderm region (ROI). Green: the same group of ectoderm cells. The ROI (in 3D) covers the lateral ectodermal region that is 60°–120° away from the ventral midline and 75-μm long along the AP axis. The change in the volume of ROI (Volec) is used as a readout for change in average tissue thickness. Bottom: representative segmented 3D views showing the ectoderm at the onset of apical constriction (T0), reaches the thickest point (Tpeak) and at the last time frame of ventral furrow formation that is reliably segmented (Tend). (b) Definition for the direction of cell movement used in (c–f). (c, d) Change in Volec over time and cell movement along A-P and D-V axis in wild-type (c) and snail mutant embryos (d). Arrowheads indicate the start of ectoderm shortening. (e) Cell movement along A-P and D-V axis in wild-type and snail mutants are replotted together for better comparison. (f) Change in Volec and invagination depth D over time in wild-type and snail mutant embryos. Arrowheads indicate the start of ectoderm shortening. (g) Representative cross-section views showing the wild-type and snail mutant embryos at T0, Tpeak, and Tend, respectively. Arrowheads indicate the apex of the mid constricting cells. N=3 embryos for each genotype. Scale bars: 20 μm. (h) Comparison of the ectoderm volume reduction rate between the wild-type and snail mutant embryos. The descending part of the volume curve was fitted into a straight line to calculate the rate of volume reduction.

This analysis led to the following observations. First, Volec increased during the first 10 min of ventral furrow formation (Figure 8c). This observation is consistent with the previous measurement of ectodermal cell length during gastrulation (Brodland et al., 2010). Second, in all cases examined, Volec started to decrease approximately halfway through ventral furrow formation, but the exact time of this transition varied between embryos (11.1±2.3 min, mean±s.d., n=3 embryos; Figure 8c and f, arrowheads). Finally, the percent reduction in Volec from the peak volume to that at the end of ventral furrow formation ranged between 4% and 8%. These observations are consistent with an independent 2D measurement of the cross-section area (as a proxy for tissue thickness) of a lateral ectodermal region that was between 60° and 90° from the ventral midline (~6% reduction; Figure 8—figure supplement 1).

The observed ectodermal shortening could be driven by an active mechanism that reduces cell length or could be a passive response to the pulling from ventral furrow. To distinguish these possibilities, we repeated the Volec measurements in snail mutant embryos. As expected, ventral furrow failed to form in snail embryos (Figure 8g). Strikingly, the reduction of Volec occurred at about the same time as in the wild-type. In addition, the rate of Volec reduction was similar between the wild-type and snail embryos, although the snail embryos on average had a smaller Volec to start with (Figure 8d, f and h). This result indicates that pulling from ventral furrow cannot fully account for the apicobasal shrinkage of lateral ectoderm in the wild-type embryos and suggests the presence of ventral furrow-independent mechanism for ectodermal shortening.

In our vertex model, ectodermal compression was generated by letting all ectodermal cells shorten by 20% (ΔL=20%; Materials and methods). Further analysis demonstrates that a minimum of 5%–10% shortening in all 60 ectodermal cells is critical for generating the binary response, which was greater than the range measured in the actual embryos (4%–8% shortening, measured in ~45% ectodermal cells; Figure 8; Figure 8—figure supplement 2). A number of factors may account for the difference between the model and the actual embryo. For example, we observed residual cellularization in the ectoderm during the first 10 min of gastrulation, but there was no cell growth in our model (Figure 8c). While an apical-basal shrinking of the ectodermal cells can generate in-plane compression, the onset and the effect of this shrinking may be underestimated by volume or length measurement if the cells are growing at the same time. In addition, other morphogenetic processes occurring during gastrulation might also contribute to the generation of tissue compression. For example, Rauzi et al., 2015 found that both the onset of rapid ventral furrow invagination and the onset of ventrally directed movement of the lateral ectoderm are delayed in the mutant that disrupts cephalic furrow formation and posterior midgut invagination (Rauzi et al., 2015). Finally, the difference between an elastic model and the actual viscoelastic embryonic tissue may also contribute to the discrepancy between the measurements and model predictions. While our model recapitulated the major cell morphological features during ventral furrow formation and correctly predicted the binary response to acute actomyosin inhibition, future research is needed to elucidate the mechanism and function of ectodermal shortening in gastrulation. An interesting prediction from the Late Group embryo observation based on our proposed model is that ectodermal shortening does not depend on Rho1 activity, since ventral furrow invagination continued in Late Group embryos in which both the mesoderm and the ectoderm were stimulated (Materials and methods). Consistent with this prediction, we found that Opto-Rho1DN stimulation only had a mild and transient effect on ectoderm thickness change, suggesting that Rho1 inhibition did not directly impact ectodermal shortening (Figure 8—figure supplement 3).

Ventrally directed movement of the lateral ectoderm still occurs when apical constriction in the mesoderm is inhibited

In theory, compression generated by ectodermal shortening should result in displacement of the ectodermal cells toward the ventral region regardless of whether mesoderm invagination occurs. Previous studies have shown that ventral movement of the lateral ectoderm still occurs in mesoderm specification mutants that do not form ventral furrow (Rauzi et al., 2015). Our own observation in the snail mutant is also consistent with this result (Figure 8e). Interestingly, we found that in snail mutant embryos, cells moved toward both ventral and posterior sides as soon as ectodermal shortening began (Figure 8d). This is in sharp contrast to the wild-type embryos, where the onset of the posteriorly directed movement commenced 20–25 min later than the onset of Volec reduction and ventrally directed movement (Figure 8c). The altered pattern of cell movement in snail embryos could be explained by a combined effect of ectodermal compression and lack of ventral furrow invagination (Figure 8f). In this hypothetical scenario, the compression promotes ventral furrow invagination in wild-type embryos, which in turn functions as a ‘sink’ to facilitate (and thereby bias) the movement of the ectoderm in the ventral direction. However, since the snail mutant embryos do not form ventral furrow, an increase in ectodermal compression induces tissue movement in both AP and DV axis simultaneously, albeit slower.

So far, data from ours and others have shown that ventrally directed ectoderm movement still occurs without ventral furrow formation. It remains unclear, however, whether this decoupling of tissue movement can be observed without altering cell fate specification. Our optogenetic approach provided an opportunity for us to test this. First, we examined the ectodermal movement in the Early Group embryos immediately after stimulation. Our measurements confirmed the “reverse” movement of the ventral cells during tissue relaxation and demonstrated that during the same time period the lateral ectoderm moved in the opposite direction, toward the ventral midline (Figure 9a–e). Second, we inhibited Rho1 at the onset of apical constriction to mimic the ventral furrow phenotype of the snail mutants, but without changing mesodermal cell fate (Figure 9f–i). Despite the lack of apical constriction, we observed ventrally directed ectodermal movement similar to that in the snail mutants (Figure 9f–i). These results indicate that the ventrally directed lateral ectoderm movement can occur independent of ventral furrow formation when cell fate specification is expected to be normal. Taken together, our observations suggest that ectodermal shortening may serve as an independent mechanical input that could function together with other mechanical factors to facilitate mesoderm invagination. We propose that apicobasal cell shortening of the ectodermal cells (along with other unknown mechanisms) generates in-plane compression during gastrulation, which functions together with active cell shape changes in the mesoderm to facilitate mesoderm invagination through epithelial buckling (Figure 9j).

Lateral ectodermal cells still move ventrally in Early Group embryos when the ventral cells undergo apical relaxation upon Opto-Rho1DN stimulation.

(a) Montage showing the surface view of an Early Group embryo during the first 6 min after stimulation (T=0). Yellow dashed line depicts ventral midline. Despite the tissue relaxation in the ventral region of the embryo (green and cyan dots), the ectodermal cells continue to move toward the ventral midline. (b) Kymograph generated from the surface view of the same movie as shown in (a). Magenta box indicates the time window where the average velocity of tissue movement is measured. (c) Cross-section view of the same embryo in (a) showing the relaxation of the apical indentation at the ventral side of the embryo. Yellow line: DV axis. (d) Velocity of tissue movement as a function of the initial distance from the ventral midline. N=7 Early Group embryos. (e) Box plot showing the velocity of tissue movement at ventral mesodermal ((ii): from –10 μm to –40 μm, and (iii): from 10 μm to 40 μm) or lateral ectodermal ((i): from –60 μm to –120 μm, and (iv): from 60 μm to 120 μm) regions of the embryo. Statistical analysis testing whether the measured velocities are significantly larger (i, iii) or smaller (ii, iv) than 0 is performed using one-tailed one-sample t-test against 0. p<0.001 for all tests. (f, g) Surface kymograph (f) and cross-section views (g) of embryos expressing Opto-Rho1DN and stimulated at the onset of apical constriction. Yellow line: ventral midline. Ectoderm cells (magenta dots) still move toward ventral midline even though no ventral furrow was formed (g). The left and right panels in (g) show the mCherry and GFP signals from the same embryo. N=3 embryos. T=0 is the onset of apical constriction. All scale bars: 20 μm. (h) Velocity of tissue movement as a function of the initial distance from the ventral midline. N = 3 snail mutant embryos. (i) Box plot showing the velocity of tissue movement at lateral ectodermal regions of the embryo ((i): from –40 μm to –110 μm, and (ii): from 40 μm to 110 μm). Statistical analysis testing whether the measured velocities are significantly larger (i) or smaller (ii) than 0 is performed using one-tailed one-sample t-test against 0. p<0.001 for both tests. (j) Cartoon depicting the proposed model for mechanical bistability of the mesoderm epithelium during gastrulation. Actomyosin-mediated apical constriction is important for the system to transition from the initial state to an intermediate state, whereas the subsequent transition into the final, fully invaginated state is facilitated by ectodermal compression and is insensitive to loss of actomyosin contractility. The presence and potential function of ectodermal compression remain to be tested experimentally.

Figure 9—source data 1

Measurements of velocity of tissue movement as a function of the initial distance from the ventral midline in Early Group embryos.

https://cdn.elifesciences.org/articles/69082/elife-69082-fig9-data1-v2.xlsx
Figure 9—source data 2

Measurements of velocity of tissue movement as a function of the initial distance from the ventral midline in Opto-Rho1DN embryos stimulated before gastrulation.

https://cdn.elifesciences.org/articles/69082/elife-69082-fig9-data2-v2.xlsx

Discussion

In this work, we developed an optogenetic tool, Opto-Rho1DN, to acutely inhibit actomyosin contractility in developing embryos. Using this tool, we discovered that while actomyosin-mediated active cell deformation is critical for mesoderm invagination, it is dispensable during the subsequent rapid invagination step after a transitional stage (Ttrans), when the initial apical constriction mediated ‘tissue priming’ is completed. These observations suggest that the mesoderm epithelium is mechanically bistable during gastrulation—eliminating the active forces that deform the tissue at different stages of ventral furrow formation causes the tissue to either fall back to the initial stable state or progress to the final, fully invaginated state. Using computer modeling, we tested a possible mechanism based on the analogy of buckling of a compressed elastic beam induced by an indentation force. We demonstrate that the binary response of the tissue can be recapitulated in a simulated folding process jointly mediated by apical constriction in the mesoderm and apicobasal shrinkage of the ectoderm. In this simulation, apicobasal shrinking of the ectoderm is expected to generate in-plane compression and increase the propensity of the epithelium to buckle. Finally, we show that apicobasal shrinkage of lateral ectoderm occurs during gastrulation in both wild-type and snail mutant embryos and may thereby offer an independent mechanical input that functions together with other mechanical forces (such as apical constriction) to facilitate mesoderm invagination. We propose that ventral furrow formation is mediated through a joint action of multiple mechanical inputs. Apical constriction drives initial indentation of ventral furrow, which primes the tissue for folding, whereas the subsequent rapid folding of the furrow is promoted by the bistable characteristics of the mesoderm and by lateral myosin contractions in the constricting cells (Figure 9j). We further hypothesize that ectodermal compression induced by ectodermal shortening induces mechanical bistability in the mesoderm, which in turn facilitates mesoderm invagination through a buckling-like mechanism (Figure 9j). Future experiments investigating the presence and potential function of ectodermal compression would be key to testing these hypotheses.

Our model proposes that compressive forces from the surrounding ectoderm contribute to mesoderm invagination and result in the binary tissue response upon acute actomyosin inhibition. The processes that may potentially generate ectodermal compression during mesoderm invagination remain to be determined. One possible candidate is germband extension. During germband extension, the convergence of tissue in the DV axis may provide a pushing force on the ventral mesodermal cells and lead to the binary tissue response to acute Rho1 inhibition. However, we do not favor this model for the following reasons. Since germ band extension is a process of convergent extension, the cellular flow along the DV axis (the ‘convergence’) is expected to be associated with an accompanying cellular flow along the AP axis (the ‘extension’). While the DV movement of the ectoderm occurs at about the same time as the onset of rapid ventral furrow invagination, the AP movement of the ectoderm did not start until ~10 min later, when the ventral furrow has nearly fully invaginated (Figure 8c). These results are consistent with previous reports (Lye et al., 2015) and suggest that the initial DV movement is caused by the ventral furrow pull, rather than the convergence of the germband. Along the same vein, the apicobasal shortening of the lateral ectoderm also happens earlier than the onset of AP movement (Figure 8c), suggesting that germband extension does not account for ectodermal cell shortening during ventral furrow formation. Furthermore, in Late Group embryos, the cellular flow in the lateral ectodermal region displays little AP tissue movement during the course of ventral furrow invagination (Figure 6—figure supplement 2). Taken together, these observations suggest that germband extension is unlikely the cause of the binary tissue response observed in our optogenetic experiment. That being said, our results do not rule out the possibility that germband extension may contribute to the closing of ventral furrow at the final stage of furrow invagination. In addition, while we consider mechanical bistability as the most parsimonious explanation of the observed binary tissue response, we could not formally rule out other possible mechanisms.

In principle, bending of a cell sheet can be achieved through two general mechanisms. In the first mechanism, an active cell deformation (e.g., apical constriction) that results in cell wedging can cause the tissue to acquire a folded configuration. In this case, the mechanism is tissue autonomous and does not necessarily require active mechanical contributions from neighboring tissues. In the second mechanism, bending of a cell sheet is prompted by an increase in the compressive stresses within the tissue (Mao and Baum, 2015). This mechanism is tissue non-autonomous and depends on the tissues surrounding the region where folding happens. The canonical model for apical constriction-mediated cell sheet folding emphasizes the role of active cell deformation in the constriction domain (Wolpert et al., 2015). A number of recent studies, however, have implicated additional mechanical contributions from outside of the constriction domain that can influence apical constriction-mediated tissue folding (Rauzi et al., 2015; Perez-Mockus et al., 2017). Our results further support this notion and suggest that ventral furrow formation involves both tissue autonomous and non-autonomous mechanisms. In our proposed model, the generation of ectodermal compression increases the buckling propensity of the epithelium, whereas the active cell shape change induced by apical constriction triggers buckling at a defined position. Without apical constriction, ectodermal compression alone is not sufficient in triggering epithelial buckling due to the high energy barrier associated with the transition from the initial state to the final, fully invaginated state (e.g., in snail mutant or Opto-Rho1DN embryos stimulated prior to Ttrans). The active cell shape change induced by apical constriction allows the tissue to overcome this energy barrier. The proposed requirement for this tissue level cooperation may explain the recent data showing that ectopic activation of Rho1 activity at different regions of cellular blastoderm in Drosophila results in different final furrow morphologies (Rich et al., 2020).

Drosophila ventral furrow formation is a robust process in that it can still happen in various scenarios when apical constriction is weakened (Leptin and Grunewald, 1990; Parks and Wieschaus, 1991; Kölsch et al., 2007; Yevick et al., 2019). Recent studies have revealed that the supracellular actomyosin network is intrinsically robust, with build-in redundancies to secure global connectivity of the network (Yevick et al., 2019). We propose that mechanical bistability of the mesoderm enabled by ectodermal shortening provides an alternative, tissue-extrinsic mechanism that allows proper invagination of the mesoderm when actomyosin contractility in the mesodermal cells is partially impaired due to genetic and environmental variations.

The use of Opto-Rho1DN in our work has revealed an unexpected, tissue-specific involvement of Rho1 in regulating cortical F-actin. We show that lateral F-actin in the ectoderm is resistant to acute Rho1 inhibition, whereas both medioapical and lateral F-actin in the mesoderm are more sensitive. The reason for the tissue-specific response of F-actin to Rho1 inhibition is not entirely clear. Previous studies suggest that the Rho1 effector Diaphanous (Dia), a formin protein, is responsible for the assembly of medioapical F-actin in the mesoderm during apical constriction (Homem and Peifer, 2008; Mason et al., 2013). This notion is consistent with our observation that medioapical actin rapidly disappeared after Rho1 inhibition. In the ectoderm, F-actin appears to be mainly distributed along the lateral membrane. This is in contrast to active Rho1, which has been found to be enriched at the junctional regions but not along the lateral membrane in ectodermal cells (Garcia De Las Bayonas et al., 2019). This observation is consistent with our data that acute Rho1 inhibition has a negligible impact on the lateral F-actin in the ectoderm and suggests the involvement of actin regulators other than the Rho1-Dia pathway (Grevengoed et al., 2003). Interestingly, it has recently been shown that the mesodermal cell fate determinant Twist and Snail function together to specify the spatial pattern of F-actin distribution in the mesoderm (Denk-Lobnig et al., 2021). The difference in the sensitivity of F-actin to acute Rho1 inhibition likely reflects cell type and subcellular location specific regulation of F-actin.

Actomyosin contractility provides a common force generation mechanism in development. To date, the studies of tissue mechanics have been largely focused on the mechanisms that generate contractile forces that actively deform the tissues (Gilmour et al., 2017; Heisenberg and Bellaïche, 2013; Chanet and Martin, 2014). It remains less well understood how the ‘passive’ mechanical status of the tissue influences the outcome of tissue deformation. The Opto-Rho1DN optogenetic tool developed in this work provides an effective approach to acutely disrupt the myosin-dependent force generation machinery, and therefore can help us begin to discern the role of ‘active’ forces and ‘passive’ mechanical properties in morphogenesis. Furthermore, our work highlights the importance of cooperation between ‘local,’ tissue-autonomous active force production and ‘global,’ tissue-nonautonomous mechanical contribution in tissue morphogenesis. It will be interesting to explore whether such cooperation plays a role in other cell sheet folding processes and whether mechanisms other than apical constriction can be employed to trigger folding in a compressed cell sheet.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Genetic reagent (Drosophila melanogaster)Sqh-GFPRoyou et al., 2002
Genetic reagent (D. melanogaster)Sqh-mCherryMartin et al., 2009
Genetic reagent (D. melanogaster)E-cadherin-GFPOda and Tsukita, 2001
Genetic reagent (D. melanogaster)UASp-P4M-mCherryChen and He, 2021
Genetic reagent (D. melanogaster)Utrophin-VenusFigard and Sokac, 2011
Genetic reagent (D. melanogaster)halo snail/CyO, Sqh-GFPMartin et al., 2009
Genetic reagent (D. melanogaster)UASp-CIBN-pm-GFP (II)Guglielmi et al., 2015
Genetic reagent (D. melanogaster)UASp-CIBN-pm (I)Guglielmi et al., 2015
Genetic reagent (D. melanogaster)UASp-CRY2-Rho1DN -mCherry (III)This studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)UASp-CIBN-pm-GFP; CRY2-Rho1DNThis studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)UASp-CIBN-pm; CRY2-Rho1DNThis studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)Maternal-Tubulin-Gal4 (67)Hunter and Wieschaus, 2000Maternal-Tubulin-Gal4 on Chromosome II
Genetic reagent (D. melanogaster)Maternal-Tubulin-Gal4 (15)Hunter and Wieschaus, 2000Maternal-Tubulin-Gal4 on Chromosome III
Genetic reagent (D. melanogaster)67; 15Hunter and Wieschaus, 2000Maternal-Tubulin-Gal4 on II and III
Genetic reagent (D. melanogaster)67 Sqh-mCherry; 15 Sqh-GFPThis studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)67 Sqh-mCherry; 15 E-cadherin-GFPThis studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)67 Sqh-GFP; 15This studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)67 Sqh-GFP; 15 Sqh-GFPThis studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)67 Sqh-mCherry; 15 Utrophin-VenusThis studySee ‘Fly stocks and genetics’ for details
Genetic reagent (D. melanogaster)67 Sqh-GFP; 15 P4M-mCherryThis studySee ‘Fly stocks and genetics’ for details
Recombinant DNA reagentpTigerS. Ferguson, State University of New York at FredoniaTransformation vector containing the attB site and 14 upstream UAS sites
Recombinant DNA reagentpTiger-CRY2-mCherry-Rho1DNThis studyTransformation construct containing CRY2-mCherry-Rho1DN
Software, algorithmMATLABMathWorkshttps://www.mathworks.com/?s_tid=gn_logoRRID:SCR_001622
Software, algorithmFijiImageJhttp://fiji.scRRID:SCR_002285
Software, algorithmEmbryo Development Geometry Explorer (EDGE)Gelbart et al., 2012https://github.com/mgelbart/embryo-development-geometry-explorer, Gelbart, 2018
Software, algorithmilastikBerg et al., 2019https://www.ilastik.org/RRID:SCR_015246

Generation of CRY2-Rho1DN construct

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To generate the CRY2-mCherry-Rho1DN fusion gene, the Rho1 CDS bearing an ACT to AAT mutation and a TGC to TAC mutation (T19N and C189Y, respectively) was synthesized and inserted to the C-terminal end of pCRY2PHR-mCherryN1 (Addgene, deposited by Chandra Tucker) through Gibson assembly. A 12-aa flexible linker sequence (GGGGSGGGGSGG) was included between the mCherry and Rho1DN sequences. The resulting CRY2PHR-mCherry-Rho1N18Y189 fusion gene was subsequently inserted into pTiger, a transformation vector containing the attB site and 14 upstream UAS GAL4-binding sites (courtesy of S. Ferguson, State University of New York at Fredonia, Fredonia, NY). The resulting pTiger-CRY2-mCherry-Rho1DN construct was sent to Genetic Services for integration into the attP2 site using the phiC31 integrase system (Groth et al., 2004).

Fly stocks and genetics

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Fly lines containing the following fluorescent markers were used: Sqh-GFP (Royou et al., 2002), Sqh-mCherry (Martin et al., 2009), E-cadherin-GFP (Oda and Tsukita, 2001), P4M-mCherry (Chen and He, 2021), and Utrophin-Venus (Figard and Sokac, 2011). To generate snail mutant embryos, we used the halo snail/CyO, Sqh-GFP line (Martin et al., 2009).

Fly lines containing CIBN-pm-GFP (II), CIBN-pm (I) (Guglielmi et al., 2015), or CRY2-Rho1DN (III) (this study, note that CRY2-Rho1DN also contains the mCherry tag) were used to generate the following stocks: CIBN-pm-GFP; CRY2-Rho1DN and CIBN-pm; CRY2-Rho1DN. To generate embryos containing maternally deposited CIBN-pm-GFP and CRY2-Rho1DN proteins, CIBN-pm-GFP; CRY2-Rho1DN females were crossed to males from the Maternal-Tubulin-Gal4 line 67.15 to generate CIBN-pm-GFP/67; CRY2-Rho1DN/15 females (‘67’ and ‘15’ refer to Maternal-Tubulin-Gal4 on chromosomes II and III, respectively [Hunter and Wieschaus, 2000]). To generate embryos containing maternally deposited CIBN-pm (without GFP) and CRY2-Rho1DN proteins, CIBN-pm; CRY2-Rho1DN females were crossed to 67.15 males to generate CIBN-pm/+; +/67; CRY2-Rho1DN/15 females. In both cases, embryos derived from the F1 females were used in the optogenetic experiments. The use of CIBN-pm without a GFP tag was to allow better visualization of GFP-tagged myosin regulatory light chain Sqh. The following Maternal-Tubulin-Gal4 lines were used: (1) 67; 15, (2) 67 Sqh-mCherry; 15 Sqh-GFP, (3) 67 Sqh-mCherry; 15 E-cadherin-GFP, (4) 67 Sqh-GFP; 15, (5) 67 Sqh-GFP; 15 Sqh-GFP, (6) 67 Sqh-mCherry; 15 Utrophin-Venus, and (7) 67 Sqh-GFP; 15 P4M-mCherry. Specifically:

Viability test for Opto-Rho1DN embryos

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Embryo viability test was performed by hand selecting cellularizing embryos derived from 67/CIBNpm-GFP; 15/CRY2-Rho1DN female flies in a dark room where the only illuminating light is red light. The selected embryos were randomly divided into two groups and placed on two fresh apple juice plates. One plate was kept in the dark and the other was placed under a beam of white light with moderate intensity. Plates were kept at 18°C for more than 48 hr, after which the hatched and unhatched embryos were counted using a Nikon stereo microscope. Three independent trials were conducted. In total, 60 out of 70 stimulated embryos did not hatch whereas all 63 unstimulated embryos hatched.

Live imaging and optogenetic stimulation

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To prepare embryos for live imaging, embryos were manually staged and collected from apple juice plates, and dechorionated with ~40% bleach (i.e., ~3% sodium hypochlorite) in a dark room using an upright Nikon stereo microscope. In optogenetic experiments, embryos were protected from unwanted stimulation using an orange-red light filter placed on top of white light coming from the stereo microscope. After dechorionation, embryos were rinsed thoroughly with water, and transferred on a 35-mm glass-bottom dish (MatTek Corporation). Distilled water was then added to the dish well to completely cover the embryos. Live imaging was performed in water at room temperature with one of the following approaches.

To examine the plasma membrane recruitment of CRY2-Rho1DN and its impact on cortical association of myosin (Figure 1c–f), embryos were imaged using a Nikon inverted spinning disk confocal microscope equipped with the perfect focus system and Andor W1 dual camera, dual spinning disk module. A CFI Plan Apo Lambda 60×/1.40 WD 0.13-mm oil objective lens was used for imaging. For membrane recruitment of CRY2-Rho1DN, time-lapse movies were taken for post-cellularization embryos at a single z-plane close to the apical side of the tissue at a frame rate of 0.46 s per frame. For cortical association of myosin, a small Z-stack (five slices, 0.5 μm step size) was taken at a rate of 3.46 s per stack. The image size is 858 by 1536 pixels with a lateral pixel size of 0.11 μm, which corresponds to a 94 μm by 169 μm region. For each embryo, pre-stimulation images were acquired using a 561-nm laser, which did not activate Opto-Rho1DN. Stimulation and post-stimulation imaging were performed simultaneously as we subsequently imaged the embryo with both 488 nm and 561 nm lasers in an alternating pattern using triggered excitation.

To examine the spatial confinement of stimulation (Figure 1g) and the effect of activation of Opto-Rho1DN on apical constriction during ventral furrow formation (Figure 1—figure supplement 1), embryos were imaged on a Leica SP5 confocal microscope with a 63×/1.3 NA glycerin-immersion objective lens. A 2× zoom was used. Twenty confocal z-sections with a step size of 1 μm were acquired every 13 s. The image size is 1024×512 pixels with a lateral pixel size of 240 nm. The total imaged volume is approximately 246×123×19 μm3. First, a single-time-point pre-stimulated image stack was acquired using a 561-nm laser. Next, stimulation was performed by taking a single-time-point image stack using both 488 nm and 561 nm lasers, which took 13 s. Stimulation was followed by post-stimulation acquisition with 561 nm laser for 20 time points (260 s). The stimulation and post-stimulation acquisition cycle were repeated until the end of the movie (i.e., stimulated for 13 s every 273 s).

To examine the rate of tissue recoil after laser ablation (Figure 3), embryos were imaged on an Olympus FVMPE-RS multiphoton system with a 25×/1.05 numerical aperture water immersion objective lens. For unstimulated embryos, a pre-ablation Z-stack was obtained using a 1040-nm laser to image a 512×100-pixel region (171×33 μm2, 3× zoom) with a step size of 2 μm, which took approximately 16 s. Laser intensity increased linearly (4%–7%) from the surface of the embryo to 100-μm deep. This pre-ablation Z-stack was used to determine the stage of the embryo. Next, a 10-frame single Z-plane pre-ablation movie was obtained with a 1040-nm laser. A 512×512-pixel region (171×171 μm3, 3× zoom) was imaged, which took approximately 1 s per frame. Next, a 920-nm laser with 30% laser intensity was used to ablate a 3D region from immediately below the vitelline membrane to ~20-μm deep. This was achieved by taking a Z-stack with a step size of 1.5 μm. The ROI of the ablated region is ~3 μm along the AP axis. The purpose of targeting multiple Z-planes for ablation was to ensure that the very apical surface of the ventral cells was ablated. This was particularly important for the stimulated embryos since the ventral cells undergo rapid apical relaxation after Rho1 inhibition. Immediately after laser ablation, a 100-frame single Z-plane post-ablation movie was acquired using both 1040 nm and 920 nm lasers. The same ROI was imaged as the pre-ablation single Z-plane movie with identical image acquisition speed. For stimulated embryos, a stimulation procedure was included before laser ablation. First, a Z-stack was acquired to determine the stage of the embryo, as described above. Then, a 458-nm diode laser with 0.3% laser intensity was used to illuminate the whole embryo under 1× digital zoom for 12 s to activate the optogenetic module. A 3-min wait time was applied after stimulation to ensure the complete inactivation of myosin and disassembly of apical F-actin before laser ablation. Following this wait time, the same laser ablation procedure as used for the unstimulated embryos was applied (pre-ablation single Z-movie, laser ablation, and post-ablation single Z-movie), except that both 1040 nm and 920 nm lasers were used for acquiring the pre-ablation movie.

The analyses of F-actin (Utrophin-Venus) after Opto-Rho1DN activation (Figure 2) and the stage-specific effect of Opto-Rho1DN activation on ventral furrow invagination (Figure 4) were conducted using the following imaging protocol. Embryos were imaged on the Olympus FVMPE-RS multiphoton system with a 25×/1.05 numerical aperture water immersion objective lens. Pre-stimulation images were obtained by using a 1040-nm laser to excite a 512×100-pixel region (171×33 μm2, 3× zoom). Continuous imaging was conducted for pre-stimulation imaging. For each time point, a 100-μm Z-stack with step size of 1 μm was obtained with linearly increased laser intensity (4%–7.5%) over Z, which took 34 s. Stimulation was performed at different stages of ventral furrow formation using a 458 nm single-photon diode laser by illuminating the whole embryo under 1× zoom for 12 s with 0.3% laser intensity. A wait period of ~20 s was imposed after stimulation for examining the stage-specific effect. Post-stimulation images were obtained using both 1040 nm and 920 nm lasers. Same setting as pre-stimulation imaging was applied, with an addition of a linear increase in 920-nm laser intensity (0.1%–0.5%) over Z. Stimulation was conducted manually every five stacks during post-stimulation imaging (i.e., stimulation for 12 s every ~200 s) to ensure the sustained membrane recruitment of CRY2-Rho1DN.

To image wild-type and snail mutant embryos for the volume measurement in the lateral ectoderm (Volec, Figure 8), embryos were imaged on the Olympus FVMPE-RS multiphoton system with a 25×/1.05 numerical aperture water immersion objective lens. Z stacks of 120 μm with a step size of 1 μm was acquired continuously over a 512×150-pixel region (256×75 μm2, 2× zoom). A 920-nm laser with a linear increase of laser intensity over Z (0.3%–2.5%) was used for imaging. The temporal resolution of the movie was ~54 s/stack. halo snail homozygous mutant embryos were derived from the halo snail/CyO, Sqh-GFP stock and were recognized based on the ‘halo’ phenotype during cellularization.

To examine ectodermal cell movement after Opto-Rho1DN activation (Figure 9f–i), embryos were imaged on the Olympus FVMPE-RS multiphoton system with a 25×/1.05 numerical aperture water immersion objective lens. A pre-stimulation Z stack was acquired using 1040-nm laser to illuminate an ROI (512×150 pixels, 171×50 μm2, 3× zoom) with a step size of 1 μm for a total depth of 100 μm. The laser intensity increased linearly over Z (3%–7%). Next, whole embryo stimulation was achieved by illuminating the embryo with a 458-nm laser at 0.3% laser intensity for ~12 s under 1× zoom. Post-stimulation Z stacks were acquired using similar protocol as the pre-stimulation Z stack, except that both 920 nm and 1040 nm lasers were used for imaging. The intensity of the 920-nm laser also increased linearly over Z (0.5%–2.5%). The temporal resolution of the post-stimulation movie was ~41 s/stack. At the end of each experiment, a single frame image of the embryo was acquired at 1× zoom to record the AP orientation of the embryo.

Image analysis and quantification

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All image processing and analysis were processed using MATLAB (The MathWorks) and ImageJ (NIH).

To quantify the percent membrane recruitment of CRY2-Rho1DN after stimulation (Figure 1d), cell membrane was segmented based on the CIBN-pm-GFP signal using a custom MATLAB code. The resulting membrane mask was used to determine the signal intensity of CRY2-Rho1DN (mCherry tagged) along the cell membrane over time. The intensity of the CRY2-Rho1DN signal in the cytoplasm, which is determined as the average intensity in the cells before stimulation, was subtracted from the measured intensity. The resulting net membrane signal intensity was further normalized by scaling between 0 and 1.

To quantify the rate of tissue recoil after laser ablation (Figure 3), the width change of the ablated region along the AP axis was measured over time from the kymograph using ImageJ. The width change during the first 20 s after laser ablation (when the change over time was relatively linear) was reported to indicate the rate of tissue recoil.

The invagination depth D (i.e., the distance between the vitelline membrane and the apex of the ventral-most cell, e.g., Figure 4b) and the apical-basal length of the ventral cells over the course of ventral furrow formation (Figure 4—figure supplement 1a,b) were manually measured from the cross-section view of the embryos using ImageJ.

To categorize embryo responses to stage-specific Opto-Rho1DN stimulation, first, each post-stimulation movie was aligned to a representative control movie based on the intermediate furrow morphology at the time of stimulation. The purpose of this alignment was to minimize the impact of embryo-to-embryo variation in TL-S trans (Figure 4—figure supplement 1d). The resulting aligned time was used in the analysis because it provided a better measure of the stage of ventral furrow formation than the ‘absolute’ time defined by the onset of gastrulation in each embryo. Next, dD/dt immediately after stimulation is determined by a linear fitting of D over time in the first 4 min after stimulation (Figure 4—figure supplement 1f). Embryos with dD/dt<–0.3 μm/min are defined as Early Group. These embryos undergo tissue relaxation and result in decrease in D over time. Embryos with dD/dt between –0.3 μm/min and 0.3 μm/min are defined as Mid Group. These embryos do not undergo obvious tissue relaxation but display a temporary pause before they continue to invaginate. Embryos with dD/dt>0.3 μm/min are defined as Late Group. These embryos continue to invaginate despite the rapid inactivation of myosin. To quantify the impact of myosin inhibition on furrow invagination (Figure 4d), the delay time in furrow invagination (Tdelay) was measured. Tdelay is defined as the difference in time for a given stimulated embryo to reach a D of 20 μm compared to a representative control embryo.

To examine the change of apical cell area of the flanking, non-constricting cells in stimulated Opto-Rho1DN embryos (Figure 4—figure supplement 2a, b), a custom MATLAB script was used to generate a flattened surface view of the embryo which accounts for the curvature of the embryo. The apical cell area was measured by manually tracking and outlining flanking cells followed by area measurement using ImageJ.

To segment the mesoderm cells (Figure 5), Carving procedure from ilastik (Berg et al., 2019) was used in combination with manual correction using ImageJ and MATLAB to segment and reconstitute the 3D shape of individual cells. Cell length was calculated by summing up the distance between neighboring centroids at consecutive z planes to account for the curvature of the cell. Apical surface area was calculated based on the intersection area between the curved apical surface of the tissue and the 3D object of the cell. This intersection area was determined using a custom MATLAB code. The lateral surface area was calculated by a subtraction of the apical and basal surface area from the overall cell surface area, which was calculated from the 3D object of the cell. The volume is calculated directly from the 3D object of the cell.

To evaluate the volume flux in the lateral ectoderm region (Figure 8), pixel segmentation procedure from ilastik (Berg et al., 2019) was used to segment a fixed lateral ectoderm region (ROI: from 60° to 120° away from the ventral midline and 75 μm long along the AP axis) in both WT and snail mutant embryos. Segmentation was performed based on Sqh-GFP signal which clearly defines the apical and basal boundaries of the tissue. The tissue was segmented as a continuum without segmentation of individual cells. The volume of the selected ROI (Volex) at each time point was determined based on the segmented 3D tissue object. To measure the movement of lateral ectoderm during gastrulation, the position of a cell initially located 50° away from the ventral midline at the onset of gastrulation was manually tracked over time using ImageJ.

To measure ectodermal shortening from 2D cross-section view of the embryos (Figure 8—figure supplements 1 and 3), the thickness of the lateral ectoderm is reported as the cross-section area of the ectodermal cell layer between 60° and 90° from the DV axis, which provides a good readout of tissue volume redistribution in 3D. A custom MATLAB script was used to facilitate the measurement. Specifically, the apical surface of the tissue was determined by automatic segmentation of the vitelline membrane of the embryo. The resulting curve was subsequently fit into a circle (the outer membrane circle). The basal surface of the tissue was determined by automatic segmentation and tracking of the basal myosin signal, with necessary manual corrections when the basal signal became too dim to be segmented accurately. Tissue thickness was measured as the cross-section area of a section of the ectodermal tissue bounded by the outer membrane, the basal surface, and two radii of the outer membrane circle that are 60° and 90° away from the ventral midline, respectively.

To detect ectodermal cell movement in Early Group embryos after stimulation (Figure 9a–e) and in embryos stimulated at the onset of gastrulation (Figure 9f–i), kymographs were generated from the surface views with the CIBN-pmGFP signal or P4M-mcherry that marks the cell membrane. Individual membrane traces within the first 6 min after stimulation were manually tracked from the kymograph using the line selection tool in ImageJ. The slope of the lines was subsequently measured using ImageJ to calculate the rate of membrane movement, which was then plotted against the position of corresponding membrane relative to the ventral midline at the time of stimulation. The average velocity of tissue movement at ventral mesodermal (from –10 μm to –40 μm and from 10 μm to 40 μm) or lateral ectodermal (from –60 μm to –120 μm and from 60 to 120 μm for Early Group embryos or <–40 μm and >40 μm for embryos stimulated at the onset of gastrulation) regions of the embryos was calculated and tested using one-tailed one-sample t-test against 0.

Energy minimization-based vertex model for ventral furrow formation (Polyakov et al. Model)

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The 2D vertex model for ventral furrow formation, which considers the cross-section view of the embryo, was constructed as previously described (Polyakov et al., 2014). In brief, the model contains a ring of 80 cells that represent the cross-section of an actual embryo. The initial geometry of the cells in the model resembles that in the real embryo. The apical, basal, and lateral membranes of the cells are modeled as elastic springs that resist deformation, and the cells have a propensity to remain constant cell volume (area in 2D). In the model, apical constriction provides the sole driving force for tissue deformation, which is achieved by allowing the apex of the cells in the ventral region of the embryo to shrink. The resulting morphological changes of the tissue are determined by the stiffness of the apical, lateral, and basal springs, cell volume conservation, and a stepwise reduction of basal spring stiffness (see below).

The energy equation that approximates the mechanical properties of the system is described by the following expression:

E= iφiμiai2+i[Kl(lil0)2+Kb(bib0)2+Ka(aia0)2]+iCVOL(Vi)+CYOLK(Vyolk)

The first term in the equation stands for myosin-mediated apical constriction. The term φiμi defines the spatial distribution of myosin contractility. φi equals to one for cells within the mesoderm domain (9 cells on each side of the ventral midline, a total of 18 cells) and 0 for the rest of cells, which ensures that only the mesodermal cells will constrict apically. μi is a Gaussian function that peaks at the ventral midline. Specifically,

μi= μ0e-(i-imid)22σ2

Here, imid stands for the cell ID at the ventral midline, μ0 defines the strength of apical constriction, and σ defines the width of the force distribution. Note that due to the Gaussian-shaped force distribution, only 12 cells at the ventral most region of the embryo will undergo apical constriction, similar to what occurs in the real embryo.

The second term in the equation stands for the elastic resistance of the membranes. ai , bi , and li stand for the area (length in 2D) of the apical, basal, and lateral membrane of cell i, respectively. a0 , b0 , and l0 are the corresponding resting length, which are set to be their initial length. ka , kb, and kl are the spring constant of the apical, basal, and lateral springs, respectively.

The third and fourth terms, CVOL and CYOLK , are constraint functions that describe volume conservation of the cells and the yolk, respectively. These two terms impose penalties when the volume deviates from the resting value. Specifically,

CVOLVi= Kv(Vi-V0)2
CYOLKVyolk= KY(Vyolk-V0, yolk)2

Here, Vi stands for the volume of cell i. V0 stands for the initial equilibrium volume of cell i. Similarly, Vyolk stands for the volume of the yolk. V0,yolk stands for the initial equilibrium volume of the yolk. Kv and Kyolk control the deviation of the cell volume and the yolk volume from the initial equilibrium values, respectively.

In the simulation, the basal rigidity of the epithelium (Kb) decreases adiabatically, allowing the model to transition through a series of intermediate equilibrium states defined by the specific Kb values. These intermediate states recapitulate the lengthening-shortening dynamics of the constricting cells observed in the real embryos (Polyakov et al., 2014).

To produce an in-plane compressive stress in the ectodermal tissue, we set the resting length of the lateral springs (l0) 20% shorter than their original length. Because of the cell volume conservation constraint, shortening of the ectodermal cells in the apical-basal direction will result in an expansion of cells in the orthogonal direction, which is the planar direction of the epithelium. This expansion provides a mechanism to produce in-plane compressive stress in the epithelial sheet. As shown in Figure 8—figure supplement 2, when the percent reduction of the ectodermal cell length is lowered from 20% to 10%, the binary response to acute actomyosin inhibition is still present, although the final depth of the furrow in the simulated Late Group embryo is reduced. Further lowering the percent reduction to 5% abolishes the binary response.

To simulate the optogenetic inhibition of myosin contractility in the model, we set the program such that the apical contractility term will be reduced to 0 at defined intermediate equilibrium state. This approach allows us to inhibit apical contractility at any intermediate furrow configuration specified by Kb .

In order to simulate the impact of impairing apical constriction on invagination, we modified our model in two different ways. First, to reduce the number of apically constricting cells, we decreased the width of the Gaussian function that defines the spatial distribution of apical myosin contractility. Second, to recapitulate the weakened and uncoordinated apical constriction observed in certain apical constriction mutants, we inhibited apical constriction from every other cell within the ventral 18 cell region.

In order to account for the impact of optogenetic stimulation of Opto-Rho1DN on lateral myosin in the constricting cells, we implemented an active lateral constriction force along the lateral edges of the constricting cells, on top of the passive lateral restoration force described above. In such a scenario, the active and passive lateral forces worked in combination to mediate ventral cell shortening, but only the active force was sensitive to myosin inactivation. For each lateral cortex within the constriction domain, the active lateral constriction force is generated by a spring with a resting length of 0. The lateral shortening force in the constricting cells is determined by:

Fshortening=KL_activel+KL_passive(l-l0)

where KL_active and KL_passive are the spring constant of the active and passive lateral springs, respectively, and l and l0 are the current and the resting length of the lateral edges, respectively. As shown in Figure 7, a spring constant of KL_active= 2 generated the desired furrow morphology under a broad range of passive spring constant (KL_passive=[0.002, 20]). A further increase of KL_active result in deeper furrows as more cells are incorporated into the neck region of the furrow. To account for the observation that lateral myosin in the constricting cells rapidly diminishes after Opto-Rho1DN activation, KL_active is set to 0 after in silico myosin inhibition.

List of parameters used in Figure 6c:

ParameterValue
Ka30
Kl20
Kb*210 to 20
μ0**5000
σ3
Kv5000
KY1

*: In the simulation, Kb decreases adiabatically from 210 to 20, with a twofold reduction at each step. An energy equilibrium state is reached for each value of Kb . These energy equilibrium states define the intermediate and final furrow morphologies.

**: The model recapitulates the binary response to acute loss of actomyosin contractility under a wide range of μ0 tested (500 – 50,000).

Statistics

Sample sizes for the presented data and methods for statistical comparisons can be found in figure legends. p values were calculated using MATLAB ttest2 or rank-sum function.

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files. Source data files have been provided for the codes for the computer models described in this work and the numerical data for Figure 4 - figure supplement 1 and Figure 9.

References

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    2. Martin AC
    (2014) Mechanical force sensing in tissues
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    https://doi.org/10.1016/B978-0-12-394624-9.00013-0
    1. Foe VE
    (1989)
    Mitotic domains reveal early commitment of cells in Drosophila embryos
    Development (Cambridge, England) 107:1–22.
    1. Nelson CM
    (2016) On Buckling Morphogenesis
    Journal of Biomechanical Engineering 138:021005.
    https://doi.org/10.1115/1.4032128
  1. Book
    1. Wolpert L
    2. Tickle C
    3. Arias AM
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    Principles of Development
    Oxford University Press.

Decision letter

  1. Michel Bagnat
    Reviewing Editor; Duke University, United States
  2. Utpal Banerjee
    Senior Editor; University of California, Los Angeles, United States
  3. Sebastian J Streichan
    Reviewer; University of California, Santa Barbara, United States
  4. Magali Suzanne
    Reviewer; Centre de BIologie Integrative (CBI), France

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Mechanical bistability enabled by ectodermal compression facilitates Drosophila mesoderm invagination" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Utpal Banerjee as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Sebastian J Streichan (Reviewer #1); Magali Suzanne (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

The authors address how contractile forces near the apical surface of a cell sheet drive out-of-plane bending of the sheet. To determine whether actomyosin contractility is required throughout the folding process and to identify potential actomyosin independent contributions for invagination, they develop an optogenetic-mediated inhibition of myosin and show that myosin contractility is critical to prevent tissue relaxation during the early stage of folding but is dispensable for the deepening of the invagination. The results shown in the first two figures support the idea that the mesoderm is mechanically bistable during gastrulation.

In the second part of this study, the authors test the role of the coupling between mesoderm and ectoderm by using 2D computational modelling and infrared pulsed laser dissection. They propose that the ectoderm can generate compressive forces on the mesoderm facilitating mesoderm internalization (2nd phase).

They then propose that this mechanical bistability arises from an in-plane compression from the surrounding ectoderm and that mesoderm invagination is achieved through the combination of apical constriction and tissue compression.

While the optogenetic experiments require additional controls, the overall results are compelling and deemed both interesting and significant for the field. By contrast, figure 4,5,6 appear highly speculative, and have substantial issues (e.g. reporting effects orders of magnitude below diffraction limit).

The manuscript presents two different models for data interpretation. The first one is a modified version of an earlier model that provides some predictions that can be tested with relatively simple experiments. On the other hand, the second model is rather complex and should be further analyzed with great care, before considering it for publication. It appears highly overparameterized, oftentimes using ad-hoc modifications for generating a desired effect. Moreover, it is a well-known fact from thin sheet elasticity that contributions of bending to total elastic energy are weighted by thickness cubed. The cell thickness shown are considerably thinner than the equivalent of cells in the embryo. At a thickness comparable to embryonic cells, bending will become orders of magnitude more costly. It further remains unclear how a dynamic variable is obtained. Thus, it is also unclear how the simulations ensure a robust trajectory in a high dimensional phase space with likely multiple minima.

After vigorous discussion with all reviewers, there emerged the possibility to focus the present manuscript on the original optogenetics findings, described in figure 1 and 2, and then quantitative analysis of predictions made by model shown in figure 3. These tests should include lateral edge lengths, across all cells in the ectoderm. Here it will be important to distinguish passive effects in the ectoderm due to pulling from the ventral furrow: If the furrow pulls, cells might actually also shorten laterally. This can be tested using wide-spread twist mutants. Finally, the authors need down their claim of compression. In the discussion, it may be mentioned the possibility of compression. However, the existing data does not support for such a mechanism at all.

Essential revisions:

1. Provide quantitative data of cell shape changes near the ventral furrow. Analysis should include both apical as well as lateral cell surface areas.

2. The authors analyze the effects of RhoDN on MyoII but never on the F-actin network. Rho1 is known to control F-actin organization so this should also be tackled thoroughly.

3. Test actomyosin contractility by measuring network recoil after laser dissection in control (RhoDN non activated) and RhoDN activated embryo.

4. Test the modification of the Polyakov model using available data. Since the lateral rest lengths are modified such that cells shorten over time (by 20 % – in real cells this would be about 8µm), if all 60 ectoderm cells shrink that much, this can result in a considerable in-plane expansion, assuming volume conservation. This could be tested by measuring the time course of average lateral length change of cells in the ectoderm (on the dorsal pole, and in the lateral regions), and explain how this compares to model assumptions.

5. Some degree of lateral cell shrinking is expected from ventral furrow pull. To distinguish possible contributions from ventral furrow vs active processes shortening the cell edges as proposed in the model, the authors should repeat the lateral cell surface analysis from (5) in twist or snail mutants.

6. To address to some extent the role of the mesoderm the authors could perform an early optogenetic RhoDN of the ventral side. If the ectoderm is pushing, then one could predict that the ventral cells should reduce their size along DV but not along AP because of the DV pushing from left and right ectoderms.

These experiments and the contrast of data with the modified Polyakov model may allow the authors to arrive at a soft conclusion implicating other forces, e.g. ectoderm compression, in the discussion.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Mechanical bistability of the mesoderm epithelium facilitates mesoderm invagination during Drosophila gastrulation" for further consideration by eLife. Your revised article has been reviewed by 3 peer reviewers and the evaluation has been overseen by Utpal Banerjee as the Senior Editor, and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

Essential revisions:

The reviewers have found the manuscript much impproved and have praised the extensive work performed to address their criticism. This has resulted in a greatly improved manuscript that needs no further experimental work and is almost ready for publication pending some editorial changes. Specifically, they have pointed out one analysis that needs attention and several discussion points that should be addressed with text changes as follows:

1. For Figure 9f the authors have opted to track an ectoderm cell over time to demonstrate ectoderm displacement. This is fine, nevertheless the author should be consistent and perform again the analysis for Figure 9f by using the same analysis procedure they implemented for Figure 9a. More precisely the authors should follow a cell that is located 20 cells away from the midline (not just 10 cell away).

Discussion points:

2. The model should be described as a theory. In its current form it is hard to distinguish from the descriptions of experiments. It should be clearly labeled that it is purely elastic, and that it neglects the well-known viscous properties of tissues that dominate on the scale of at least 4 minutes and beyond.

3. On page 19 and eventually in the discussion the discrepancy between the model and the in vivo measurements should be discussed. More precisely, along a cross-section in the model the minimum necessary ectoderm cell shortening is 5-10% for 60 cells while in the real embryo the cell-shortening measured is 4-8% for a much smaller number of ectoderm cells closer to the mesoderm (20 cells?). Please consider these potential discrepancies and if they are indeed present discuss its possible origin and/or speculate in the discussion what might account for them.

4. In the new version of the paper, it is pointed out that F-actin in ectoderm cells is not affected by RhoDN optogenetic activation. This is quite puzzling and therefore merits at least further discussion.

5. In Figure 4A and in many other experiments, RhoDN is activated both in the mesoderm and in the ectoderm. Therefore, by following the logic of the model, ectoderm pushing is not dependent on Rho signaling. In other words, while mesoderm cell shortening depends on Rho, ectoderm cell shortening is Rho independent. This also is quite surprising thus merits further discussion.

6. The manuscript has valuable data on cell behaviors in the lateral ectoderm. But, the presentation is entirely focused around the idea of 'compression', and no alternatives discussed. One such alternative, could be the impact of germband extension on deepening the already formed furrow. The cellular flow of germband has a component directed towards the ventral pole, possibly allowing cells to flow into furrow, that has already formed. This extension will lead to the observed apicobasal shortening of lateral ectoderm cells, and deepening of the fold, but requires no mechanical bistablity. It would further be consistent with lack of fold and the described cell shapes at the apical surface in Snail and opto RhoDN experiment.

7. Stress is needed to build up for the proposed buckling by compression. However, the Snail experiments clearly demonstrate that no buckling of mesoderm occurs when cell behaviors in the mesoderm are perturbed. Stating mesoderm buckles due to compression from the ectoderm is, therefore, misleading and has not been demonstrated with an experiment. Any mention of this interpretation should be confined to the discussion.

8. In the discussion, the statement "Using computer modeling, we further demonstrated that mechanical bistablity in the mesoderm can arise from an apicobasal shrinkage of the ectoderm, which generates in-plane compression as the cell volume remains conserved" is misleading. This needs to be clarified, if the authors wish to raise the idea of bistability in the Discussion section.

9. While the authors consistently claim precise agreement between the model and data, it remains unclear to what extend this is the case. Visuals of simulations are provided. But there are no quantitative comparisons found that directly compare a model result with a corresponding measurement. Therefore, such claims (e.g. page 14 "In particular, the transitional state of the tissue revealed in the simulation is nearly identical to that identified in our ontogenetic experiments") have to be toned down (e.g. looks visually similar).

10. The quantitative analysis shown in figure 8 appears inconsistent with the descriptions. First, the authors refer to a rate of volume reduction, but show volume. Moreover, rate of volume reduction in WT appears consistently different from snail, yet is described as very similar. Such strong claims should either be toned down or backed up with a statistical significance test.

11. In the text page 20, the authors describe "In the wild type embryos, the compression promotes ventral furrow invagination, which in turn functions as a 'sink' to facilitate the movement of the ectoderm in the ventral direction". There is no experimental evidence provided for compression, and therefore this statement is speculation. Please rephrase.

Reviewer #1 (Recommendations for the authors):

The revised manuscript by Guo et al. has been revised, addressing some but not all of my concerns. In fact, the manuscript provides additional data, that argues strongly against the proposed mechanical bistability mechanism. The manuscript reads like two separate works.

The authors raise an interesting question in the abstract: Is myosin contractility at the apical surface required throughout folding? In the current version, characterization of the opto tool is much improved. It allows the authors to demonstrate apical myosin activity is not needed in a late phase of furrowing. The results presented in the first five figures are very interesting on their own, and provide a valuable contribution to the field. In my opinion, this would be an excellent point to stop the manuscript and enter the Discussion section. Such a work would be a great addition to eLife.

Instead, the authors enter a new direction, and propose mechanical bistablity of the mesoderm, to explain these ideas. This is somewhat unclear, as there are many possibly simpler explanations consistent with this very interesting observation that are not discussed (see more below). Instead, in figures six to nine, the manuscript hinges on speculation, and a purely elasticity-based model in combination with analysis of cell geometry along the apicobasal surface in support of their hypothesis. The conclusion, that invagination requires mechanical bistabilty of the mesoderm cannot be supported with the data presented. These claims should be toned down before publication, as already suggested in the first round of revisions. It seems this problem can be fixed by clarifications, and moving speculative data interpretation from the Results section to the Discussion sections.

(1) The manuscript offers no support for mechanical bistablity assumption. That purely elastic materials can buckle under compression is well established. But, the authors supply data that argues against mechanical bistablity in this system.

– Experimental data does not go beyond correlation, and yet the mechanism presented claims a causal role of ectoderm compression for ventral furrow folding. The original manuscript attempted to provide experimental data in support of the mechanical instability model. The current figures describe cell shapes in 3D, but there is no test of causality offered. This is somewhat puzzling, as the optogenetic tool should also function in the lateral ectoderm.

– The model should be described as a theory. In its current form it is hard to distinguish from the descriptions of experiments. It should be clearly labeled that it is purely elastic, and that it neglects the well-known viscous properties of tissues that dominate on the scale of at least 4 minutes and beyond.

– It is not clear to what extend the material in figure 8 supports the main argument. Instead, it seems to show the opposite. Lateral ectoderm shorting happens wether or not ventral furrow forms. This is a clear demonstration that the proposed mechanical bistability assumption is not able to drive tissue folding. Instead, these results suggest that folding needs to occur through an independent mechanism. Deepening of the fold could be generated by another mechanism (see below).

– Compression, as indicated by the authors, implies reduction in apical surface area of compressed cells, which is not shown in Snail or early opto RhoDN experiment. Early opto RhoDN experiments are described as heterogeneous cell morphology, but not further analyzed because of technical challenges. It is not clear how this is a technical problem, and not an issue of data interpretation. Heterogeneous apical cell area is consistent with cell shear, but not with external compression.

2) Discussion of model limitations and alternative scenarios.

– I congratulate the authors on their observation in figure 4b. It seems reasonable to further analyze this interesting phenomenon, and study the possible impact of tissue tissue interactions. In doing so, the manuscript would benefit from an open approach.

– The manuscript has valuable data on cell behaviors in the lateral ectoderm. But, the presentation is entirely focused around the idea of 'compression', and no alternatives discussed.

– One such alternative, could be the impact of germband extension on deepening the already formed furrow. The cellular flow of germband has a component directed towards the ventral pole, possibly allowing cells to flow into furrow, that has already formed. This extension will lead to the observed apicobasal shortening of lateral ectoderm cells, and deepening of the fold, but requires no mechanical bistablity. It would further be consistent with lack of fold and the described cell shapes at the apical surface in Snail and opto RhoDN experiment.

– The timescale of elasticity is very short compared to the 20 minutes of ventral furrow. As pointed out by the authors, the cited paper by Doubrovinsky provides an estimate for the transition to viscosity within 4 minutes. It is one of the longest currently published timescales for this process. As the authors clearly demonstrated turnover of the actomyosin cytoskeleton is very fast, further indicating the four minutes estimate is an upper bound of what is to be expected for these cells.

But, even if it is as long as four minutes, viscosity means, stresses will dissipate. Stress however is needed to build up for the proposed buckling by compression. Snail experiments clearly demonstrate that no buckling of mesoderm occurs when cell behaviors in the mesoderm are perturbed. Stating mesoderm buckles due to compression from the ectoderm is misleading, and has not been demonstrated with an experiment. Any mention of this interpretation should be confined to the discussion.

– In the discussion, the statement "Using computer modeling, we further demonstrated that mechanical bistablity in the mesoderm can arise from an apicobasal shrinkage of the ectoderm, which generates in-plane compression as the cell volume remains conserved" is misleading. The authors neither showed that ectoderm compresses mesoderm, nor is it a novel result that elastic systems can buckle under compression. This needs to be clarified, if the authors wish to raise the idea of bistability in the Discussion section.

– While the authors consistently claim precise agreement between the model and data, it remains unclear to what extend this is the case. Visuals of simulations are provided. But there are no quantitative comparisons found that directly compare a model result with a corresponding measurement. Therefore, such claims (e.g. page 14 "In particular, the transitional state of the tissue revealed in the simulation is nearly identical to that identified in our ontogenetic experiments") have to be toned down (e.g. looks visually similar).

Reviewer #2 (Recommendations for the authors):

Guo et al. have revised their paper by following the reviewer's suggestion.

The science presented is now more solid and merits publications after addressing the following 4 points:

1) In the new version of the paper, Guo and colleagues point out the fact that F-actin in ectoderm cells is not affected by RhoDN optogenetic activation. This is quite puzzling and therefore merits at least further discussion.

2) In Figure 4A and in many other experiments, RhoDN is activated both in the mesoderm and in the ectoderm. Therefore, by following the logic of the authors model, ectoderm pushing is not dependent on Rho signaling. In other terms, while mesoderm cell shortening depends on Rho, ectoderm cell shortening is Rho independent. This also is quite surprising thus merits further discussion.

3) For Figure 9f the authors have opted to track an ectoderm cell over time to demonstrate ectoderm displacement. This is fine, nevertheless the authors should be consistent and perform again the analysis for Figure 9f by using the same analysis protocol implemented for Figure 9a. More precisely the authors should follow a cell that is located 20 cells away from the midline (not just 10 cells away).

4) At page 19 and eventually in the discussion the authors should emphasize the discrepancy between the model and the in vivo measurements. More precisely they should make clear that along a cross-section in the model the minimum necessary ectoderm cell shortening is 5-10% for 60 cells while in the real embryo the cell-shortening measured is 4-8% for a much smaller number of ectoderm cells closer to the mesoderm (20 cells?).

Reviewer #3 (Recommendations for the authors):

The authors have done a great job responding to the concerns raised by the 3 referees with the quantitative analysis of cell shape after Rho1 inhibition, the analysis of the impact of Rho1 inhibition on F-actin, new laser ablation experiments to confirm the inactivation of myosin with their opto-Rho1DN construct, the analysis of the extend of endodermal shortening both on control and snail mutant embryos, the addition of active lateral contraction in the mesoderm in the model. They have added new data and analysis that strengthen the overall impact of the paper. They further discuss their findings in a more general context regarding previous works. I strongly support publication.

https://doi.org/10.7554/eLife.69082.sa1

Author response

[…] Essential revisions:

1. Provide quantitative data of cell shape changes near the ventral furrow. Analysis should include both apical as well as lateral cell surface areas.

We thank the reviewers for this suggestion and agree that a quantitative analysis of cell shape change after Rho1 inhibition will provide useful insights on the cellular basis of the different tissue behavior between the Early and Late Groups embryos. We performed three-dimensional reconstruction of a single row of cells along the medial-lateral axis from the ventral midline in three Early Group and three Late Group embryos. Because of the left-right symmetry of the embryo at this stage, we focused on the cells on one side of the ventral midline. The row of cells included the ventral mesodermal cells (~cell 1 – 6, “constricting cells”), the lateral mesodermal cells (~cell 7 – 9, “non-constricting flanking cells”), and presumably some ectodermal cells (~cell 10 – 11). The results of this analysis are now presented in Figure 5.

The major finding of this analysis is the differences in the cell behavior that explains the different tissue behavior in Early and Late Group embryos. For both group of embryos, the stimulation of Opto-Rho1DN resulted in an apical area relaxation in the constricting cells and an accompanying apical area reduction in the flanking cells (Figure 5). However, the apical area change results in very different change in cell curvature along the apical-basal axis. Before stimulation, cells located at the side of the constriction domain (cells 4 – 6, “side constricting cells”) and the flanking cells in both Early and Late Group embryos were bent over towards the ventral midline. After stimulation, all cells in Early Group embryos straightened up and partially restored their initial, columnar cell shape (Figure 5e, illustrated in Figure 5b). This cell shape change is associated with a reduction in the apical-basal cell length and lateral cell area (Figure 5d, i; Figure 5 – figure supplement 1; cells 4 – 9). In contrast, in Late Group embryos, only the flanking cell adjacent to the ectoderm (Figure 5j, cell 9) straightened up, whereas the side constricting cells and their neighboring flanking cells (cell 4 – 8) either remained their curvature or bent further towards the ventral midline (Figure 5j, illustrated in Figure 5g). A second major difference is that in Late Group embryos, the middle constricting cells (cell 1 – 3) underwent rapid cell shortening as the furrow continues to invaginate after stimulation (Figure 5i, illustrated in Figure 5g), which were not observed in Early Group embryos but rather resembles the normal cell shortening process during furrow invagination (Polyakov et al., 2014). Together, the continued bending of the side-constricting cells and the neighboring flanking cells and the continued shortening of the mid-constricting cells in Late Group embryos elucidates the cellular basis for the continued deepening of the furrow after Opto-Rho1DN stimulation and explains the tissue level difference between Early and Late Group embryos.

2. The authors analyze the effects of RhoDN on MyoII but never on the F-actin network. Rho1 is known to control F-actin organization so this should also be tackled thoroughly.

We thank the reviewers for raising this important question. During the revision, we examined the localization of an F-actin marker UtrophinABD-Venus before and after Opto-Rho1DN stimulation. The new data is now included in Figure 2.

In short, we found that stimulation of Opto-Rho1DN results in rapid diminishing of apical F-actin in the constricting cells. In unstimulated embryo, F-actin was enriched at the apical domain of the constricting cells (Figure 2a, cyan arrows). In addition, F-actin was also localized along the lateral membrane in both constricting and non-constricting cells (referred to as “lateral F-actin”, Figure 2a, yellow and magenta arrows). When we stimulated embryos during apical constriction, apical F-actin disappeared within 1.2 minutes after Rho1 inhibition (Figure 2b). The lateral F-actin in the constricting cells was not immediately affected and only appeared to diminish 4 minutes after stimulation (Figure 2b, T = 10.8 min, yellow arrows). In contrast, the lateral F-actin in the ectodermal cells was not significantly affected within the time frame of the experiment (Figure 2b, magenta arrows). Thus, stimulation of Opto-Rho1DN results in rapid inactivation of both apical myosin and diminishing of apical F-actin, both contributing to the rapid inactivation of apical actomyosin contractility.

3. Test actomyosin contractility by measuring network recoil after laser dissection in control (RhoDN non activated) and RhoDN activated embryo.

Following the reviewers’ suggestion, we measured tissue recoil along A-P axis after laser ablation in embryos at the lengthening phase of ventral furrow formation with or without stimulation. In Opto-Rho1DN stimulated embryos, we performed laser ablation 3 min after photo-activation of Opto-Rho1DN to ensure the complete inactivation of apical myosin and disassembly of apical F-actin. The new data are now included in Figure 3. One challenge associated with this experiment is that due to apical relaxation after stimulation, the cell apex moved rapidly to a more apical Z position. This change, together with the disappearance of apical myosin, made it difficult to identify the exact apical surface of the tissue. In order to ensure that we ablate the apical surface of the cells in both unstimulated and stimulated embryos, we ablated a 3D region of the tissue starting from immediately below the vitelline membrane to ~20 μm deep (illustrated in Figure 3a). Ablation was performed using a 920 nm femtosecond laser, with the laser intensity carefully tuned to avoid any non-specific heat-induced tissue damage. Quantifications showed a clear reduction of tissue recoil in the stimulated embryos compared to the unstimulated embryos, suggesting that Opto-Rho1DN stimulation during apical constriction results in a rapid loss of apical tension (Figure 3b, c and d).

4. Test the modification of the Polyakov model using available data. Since the lateral rest lengths are modified such that cells shorten over time (by 20 % – in real cells this would be about 8µm), if all 60 ectoderm cells shrink that much, this can result in a considerable in-plane expansion, assuming volume conservation. This could be tested by measuring the time course of average lateral length change of cells in the ectoderm (on the dorsal pole, and in the lateral regions), and explain how this compares to model assumptions.

We appreciate the reviewers’ suggestion. To further investigate the extent of ectodermal shortening during gastrulation, we performed 3D tissue volume measurement in the lateral ectodermal region of the wildtype embryo (ROI: 60-120 degrees away from the ventral furrow midline and 75 μm long along the AP axis; Figure 8a). We used the term “Volec” to refer to the volume of this ROI. We focused our analysis within the lateral ectodermal region since a previous study shows that the dorsal ectoderm does not significantly contribute to ventral furrow formation(Rauzi et al., 2015). We reasoned that if the ectodermal cells undergo apical-basal shortening, there should be a net volume outflux from the ROI, causing a reduction in Volec (Figure 8a). Since cell shortening may not be uniform across the tissue, the change in Volec provides a less noisy measure of the change in average tissue thickness. To measure Volec, we performed 3D tissue segmentation in the lateral ectoderm region without segmenting individual cells (Methods).

This analysis led to the following major observations, which are presented in Figure 8. First, Volec increased during the first ten minutes of ventral furrow formation (Figure 8c). This observation is consistent with the previous measurement of ectodermal cells during gastrulation (Brodland et al., 2010) and this increase in Volec likely reflects residual cellularization. Second, in all cases examined, Volec started to decrease approximately halfway through ventral furrow formation, but the exact time of this transition varied between embryos (11.1 ± 2.3 min, mean ± s.d., n = 3 embryos; Figure 8f, arrowheads). Finally, the percent reduction in Volec from the peak volume to that at the end of ventral furrow formation ranged between 4 – 8%. These observations are consistent with an independent 2D measurement of the cross-section area (as a proxy for tissue thickness) of a lateral ectodermal region (~ 6% reduction; Figure 8 – figure supplement 1).

In our vertex model, ectodermal compression was generated by letting all ectodermal cells shorten by 20% (ΔL = 20%; Methods), which was greater than the range measured in the actual embryos. We therefore tested the impact of reducing the extent of ectoderm shortening in our model We found that a minimum of 5 – 10% shortening in all 60 ectodermal cells is critical for generating the binary response (Figure 8 – figure supplement 2). When the percent reduction of the ectodermal cell length is lowered from 20% to 10%, the binary response to acute actomyosin inhibition is still present, although the final depth of the furrow in the simulated Late Group embryo is reduced. Further lowering the percent reduction to 5% abolishes the binary response – in this case, the intermediate furrow always relaxes back to the surface of the embryo after myosin inhibition regardless of the stage of stimulation.

A number of factors may account for the difference between the model and the actual embryo. For example, we observed residual cellularization in the ectoderm during the first few minutes of gastrulation, but there is no cell growth in the model. While an apical-basal shrinking of the ectodermal cells can generate in-plane compression, the onset and the effect of this shrinking may be underestimated by volume or length measurement if the cells are growing at the same time (Figure 8c, first 10 min). In addition, other morphogenetic processes happening during gastrulation, such as the formation of cephalic furrow and posterior midgut, might also contribute to the generation of tissue compression (Rauzi et al., 2015). Finally, while there is good evidence suggesting that the elasticity assumption is a reasonable approximation for tissue properties during ventral furrow formation (see our response to Reviewer #1’s comment #1 for further elaboration on this point), the embryonic tissue is viscoelastic instead of purely elastic.

However, despite these limitations, the model successfully recapitulated several main cell morphological features during ventral furrow formation, such as cell lengthening during apical constriction and shortening/wedging during invagination. The model also correctly predicted the ability of the mesoderm tissue to invaginate when apical constriction is partially impaired. Finally, the model recapitulated nearly all aspect of tissue response to acute actomyosin inhibition, including the binary tissue response, the morphology of the intermediate furrow at the transitional state, and the reduced final furrow width but not furrow depth in Late Group embryos. Using a minimalistic approach, the model was designed to identify the most critical active and passive mechanical inputs that can lead to the observed tissue behavior. The agreement between many major experimental observations and the modeling predictions suggests that the model captures the key features of the mechanism underlying the process. That being said, we acknowledge that future research is required to further understand the behavior and function of the ectodermal tissue in mesoderm invagination.

5. Some degree of lateral cell shrinking is expected from ventral furrow pull. To distinguish possible contributions from ventral furrow vs active processes shortening the cell edges as proposed in the model, the authors should repeat the lateral cell surface analysis from (5) in twist or snail mutants.

It is indeed important to distinguish the active (e.g. “apical-basal contraction” of the lateral ectoderm) versus passive (e.g. ventral furrow pull) mechanisms that may account for lateral cell shrinking during ventral furrow formation. Following this suggestion, we repeated the 3D lateral volume (Volec) measurements in snail mutant embryos. The new results are included in Figure 8. We observed a similar reduction in Volec in snail mutant embryos as in WT embryos. Specifically, the reduction of Volec occurred at about the same time as in the wildtype. In addition, the rate of Volec reduction was similar between the wildtype and snail mutant embryos, although the snail embryos on average had a smaller Volec to start with (Figure 8d and f). This result indicates that pulling from ventral furrow cannot fully account for the lateral cell shortening in the wildtype embryos and suggests the presence of ventral furrow-independent mechanism for ectodermal shortening.

Interestingly, we also observed a different pattern of lateral ectodermal movement in snail mutant embryos when compared to the wildtype embryos. In wildtype embryos, the onset of Volec reduction is in general aligned with the onset of ventrally directed cell movement, whereas the onset of the posteriorly directed movement happens 20 – 25 minutes earlier (Figure 8c). However, in snail mutant embryos, cells move towards both ventral and posterior sides as soon as ectodermal shortening begins (Figure 8d). This observation can be explained by the combined effect of ectodermal compression and lack of ventral furrow invagination (Figure 8f). In wildtype embryos, the compression promotes ventral furrow invagination, which in turn functions as a “sink” to facilitate (and thereby bias) the movement of the ectoderm in the ventral direction. However, since the snail mutant embryos do not form ventral furrow, an increase in ectodermal compression induces tissue movement in a both A-P and D-V axis simultaneously, albeit slower.

Taken together, our data show that both ectoderm shortening and the associated ventrally directed ectoderm movement still occur without ventral furrow formation. These observations suggest that ectoderm shortening may serve as an independent mechanical input that facilitates mesoderm invagination by promoting mechanical bistability in the mesoderm.

6. To address to some extent the role of the mesoderm the authors could perform an early optogenetic RhoDN of the ventral side. If the ectoderm is pushing, then one could predict that the ventral cells should reduce their size along DV but not along AP because of the DV pushing from left and right ectoderms.

Following the reviewers’ suggestion, we performed the early stimulation experiment and presented the new data in Figure 9. A minor deviation from the suggested experiment is that instead of measuring apical domain size/shape of the ventral cells, we directly measured the movement of the lateral ectodermal cells after stimulation. This is mainly due to technical reasons. After Rho1 inhibition and disappearance of apical myosin, the cell morphology became more heterogeneous over time, which complicated the evaluation of cell aspect ratio. We reasoned that if the ectoderm is pushing the mesoderm from both sides, we should be able to observe movement of the ectodermal cells towards the ventral region even if the ventral furrow is inhibited (like in the snail mutant). Our result is consistent with this prediction (Figure 9f-g). In addition, the result is also consistent with our observation in Group Early embryos (Figure 9d and 9f-g). As reported in our original manuscript, in Early Group embryos, despite the immediate relaxation of the constricting cells after stimulation, the lateral ectoderm continued to move towards the ventral midline during the same time period (Figure 9a-9e). Together, the results of these experiments indicate that the ventrally directed movement of lateral ectoderm can occur independent of ventral furrow formation. These observations suggest that ectodermal shortening may serve as an independent mechanical input that promotes mechanical bistability in the mesoderm and thereby facilitates mesoderm invagination.

These experiments and the contrast of data with the modified Polyakov model may allow the authors to arrive at a soft conclusion implicating other forces, e.g. ectoderm compression, in the discussion.

Additional modeling analysis to test the known mechanisms for mesoderm invagination:

In addition to the new experiments mentioned above, we also included additional modeling analysis to further test the known mechanisms that promote ventral furrow invagination (as suggested by Reviewer #3). In particular, we sought to further test the model in light of the recent findings that myosin accumulates along the lateral membrane of the constricting cells (“lateral myosin”) facilitates cell shortening and tissue invagination (Gracia et al., 2019; John and Rauzi, 2021). In our optogenetic experiments, we found that lateral myosin diminished rapidly after Opto-Rho1DN stimulation, but this phenomenon was not considered in our original vertex model.

In our vertex model presented in original Figure 3 (current Figure 6), shortening of the constricting cells is mediated by passive elastic restoration forces generated in these cells as their lateral edges are stretched during cell lengthening (Polyakov et al., 2014). This passive elastic force is not affected by in silico myosin inactivation. To account for our observations in the actual optogenetic experiments, we incorporated an active lateral constriction force in the constricting cells that is sensitive to in silico myosin inactivation. In such a scenario, the active and passive lateral forces worked in combination to mediate ventral cell shortening, but only the active force was sensitive to myosin inactivation (Methods). The active and passive lateral forces are given by KL_activel and KL_passive(l-l0), respectively, where l and l0 are the current and the resting length of the lateral edge, respectively. As expected, the addition of the active lateral force allows us to reduce the passive lateral force while still generate furrows with normal morphology (Figure 7a). This modification allowed us to examine how the binary response of our model would be influenced if myosin inactivation impairs both apical constriction and cell shortening in the constriction domain. The result of this new analysis is presented in Figure 7 and discussed in the main text.

In short, we tested the impact of myosin inhibition with the combinations of KL_active and KL_passive that give rise to normal furrow morphology (Figure 7a, green shaded region). We found that the binary response of the model to myosin inhibition still exists when the cell shortening forces in the mesoderm become sensitive to myosin inhibition. However, a minimal level of myosin-independent lateral elastic force (KL_passive > 0.2) is required for the model to generate realistic final furrow morphology in Late Group embryos (Figure 7b). These results predict that mesoderm invagination demands an adequate amount of cell shortening force in the constricting cells that is insensitive to Rho1 inhibition. Interestingly, we found that lateral F-actin in the constricting cells persisted for several minutes after Opto-Rho1DN stimulation (Figure 2b, yellow arrows), which might provide the lateral elasticity described in our model.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Essential revisions:

The reviewers have found the manuscript much improved and have praised the extensive work performed to address their criticism. This has resulted in a greatly improved manuscript that needs no further experimental work and is almost ready for publication pending some editorial changes. Specifically, they have pointed out one analysis that needs attention and several discussion points that should be addressed with text changes as follows:

1. For Figure 9f the authors have opted to track an ectoderm cell over time to demonstrate ectoderm displacement. This is fine, nevertheless the author should be consistent and perform again the analysis for Figure 9f by using the same analysis procedure they implemented for Figure 9a. More precisely the authors should follow a cell that is located 20 cells away from the midline (not just 10 cell away).

We thank the reviewers for this suggestion. Following the suggestion, we performed the same quantification as in Figure 9a for the phenotype described in Figure 9f. The new results are now incorporated into the updated Figure 9 (Figure 9h and 9i). Although the data are a little noisy, there is a quasi-linear relationship between the velocity of cell movement and the initial distance of the cell from the ventral midline (Figure 9h). We further examined the cell velocity in the -110 μm to -40 μm region and the 40 μm to 110 μm region, respectively. These regions correspond to ~ 7 cells to ~14 cells away from the ventral midline at the beginning of gastrulation. Our result shows that the velocities of both groups of cells are significantly different from zero (Figure 9i). These quantifications are consistent with our visual inspection and confirm that ectoderm cells move towards ventral midline even when apical constriction is inhibited.

Discussion points:

2. The model should be described as a theory. In its current form it is hard to distinguish from the descriptions of experiments. It should be clearly labeled that it is purely elastic, and that it neglects the well-known viscous properties of tissues that dominate on the scale of at least 4 minutes and beyond.

Following reviewer’s suggestion, in the revised manuscript, we carefully distinguished between descriptions of experimental observations and conclusions drawn from the modeling analysis. Since the computer model is chosen to test the ideas of compression and buckling-like behavior, we are not able to avoid mentioning these terms before the Discussion section; however, we make it explicit that these concepts are proposed model rather than experimental conclusions.

To emphasize the difference between the actual embryo mechanical properties and that of the model we used in our work, we added the following sentence to the revised manuscript to state the limitation of the model and the reason why we chose this model to test our hypothesis:

“Note that the elasticity assumption in this model is a simplification of the actual viscoelastic properties of the embryonic tissue. […] It is therefore advantageous to use this model to explore the main novel aspect of the folding mechanics underlying ventral furrow formation and to test the central concept of our hypothesis based on relatively small number of assumptions.” (Page 15).

3. On page 19 and eventually in the discussion the discrepancy between the model and the in vivo measurements should be discussed. More precisely, along a cross-section in the model the minimum necessary ectoderm cell shortening is 5-10% for 60 cells while in the real embryo the cell-shortening measured is 4-8% for a much smaller number of ectoderm cells closer to the mesoderm (20 cells?). Please consider these potential discrepancies and if they are indeed present discuss its possible origin and/or speculate in the discussion what might account for them.

There is indeed a difference between the model and the in vivo measurements regarding the extent of ectodermal shortening that is needed in order to generate the binary tissue response in the model versus the actual extent of ectodermal shortening observed in the embryo. In the previous version of the manuscript (page 19), we discussed these discrepancies as follows:

“In our vertex model, ectodermal compression was generated by letting all ectodermal cells shorten by 20% (ΔL = 20%; Methods), which was greater than the range measured in the actual embryos. […] While our model recapitulated the main cell morphological features during ventral furrow formation and correctly predicted the binary response to acute actomyosin inhibition, future research is needed to elucidate the mechanism and function of ectodermal shortening in gastrulation.”

In the revised manuscript, we further expanded the discussion on this matter, as shown below:

“In our vertex model, ectodermal compression was generated by letting all ectodermal cells shorten by 20% (ΔL = 20%; Methods). […] While our model recapitulated the major cell morphological features during ventral furrow formation and correctly predicted the binary response to acute actomyosin inhibition, future research is needed to elucidate the mechanism and function of ectodermal shortening in gastrulation.” (Page 20)

4. In the new version of the paper, it is pointed out that F-actin in ectoderm cells is not affected by RhoDN optogenetic activation. This is quite puzzling and therefore merits at least further discussion.

This is a very interesting question. Our Rho1 inhibition showed that F-actin in the ectoderm is resistant to acute Rho1 inhibition, whereas the F-actin in the mesoderm is more sensitive. Specifically, apical F-actin rapidly disappeared within 1.2 minutes after Rho1 inhibition, whereas the lateral F-actin appeared to diminish 4 minutes after stimulation in the mesoderm. The reason for the tissue-specific response of F-actin to Rho1 inhibition is not entirely clear. Previous studies suggest that the Rho1 effector Diaphanous (Dia), a formin protein, is responsible for the assembly of medioapical F-actin in the mesoderm during apical constriction (Homem and Peifer, 2008; Mason et al., 2013). This notion is consistent with our observation that medioapical actin rapidly disappeared after Rho1 inhibition. In the ectoderm, F-actin appears to be mainly distributed along the lateral membrane. While Rho1 protein is uniformly distributed along the apical-basal axis of the cell, active Rho1(as detected by GFP-tagged Rho1 binding domain of anillin) is enriched at the junctional regions (and to some extent medioapcial region), but not along the lateral membrane in ectodermal cells (Garcia De Las Bayonas et al., 2019). This observation is consistent with our data that acute Rho1 inhibition has a negligible impact on the lateral F-actin in the ectoderm and suggests the involvement of actin regulators other than the Rho1-Dia pathway (Grevengoed et al., 2003). Interestingly, it has recently been shown that the mesodermal cell fate determinant Twist and Snail function together to determine specify the spatial pattern of F-actin distribution in the mesoderm (Denk-Lobnig et al., 2021). The difference in the sensitivity of F-actin to acute Rho1 inhibition likely reflects cell type and subcellular location specific regulation of F-actin.

We have now included the above discussion in the revised Discussion (Page 27 – 28).

5. In Figure 4A and in many other experiments, RhoDN is activated both in the mesoderm and in the ectoderm. Therefore, by following the logic of the model, ectoderm pushing is not dependent on Rho signaling. In other words, while mesoderm cell shortening depends on Rho, ectoderm cell shortening is Rho independent. This also is quite surprising thus merits further discussion.

This is an excellent point. To gain further insight on this issue, we examined the potential impact of acute Rho1 inhibition on ectodermal cell length change. We measured the ectoderm cross-section area as a proxy of the ectodermal cell length for Late Group embryos (following the protocol described in Figure 8—figure supplement 1). The new data is presented in Page 21 – 22 and Figure 8—figure supplement 3 of the revised manuscript.

Interestingly, in three out of five Late Group embryos we examined, Opto-Rho1DN stimulation resulted in a slight increase in ectoderm thickness (Figure 8—figure supplement 3a, magenta boxes). This increase is both mild and transient, and the trend of change in thickness rapidly (in 2 – 3 min) resumed the pre-stimulation pattern. In the remaining two out of five Late Group embryos, Opto-Rho1DN stimulation did not have an obvious impact on ectoderm thickness (Figure 8—figure supplement 3a, blue boxes). To further analyze this observation, we compared the Late Group embryo to the non-stimulated control embryos for ectoderm area change within a 1.7 min duration, either immediately after stimulation (Late Group embryos) or after the maximal ectoderm area was reached (non-stimulated control embryos). In control embryos, the ectoderm area on average reduced by 1%, whereas in Late group embryos, the area on average increased 0.6% (Figure 8—figure supplement 3b).

Since the impact of Opto-Rho1DN stimulation on ectoderm thickness change was both mild and transient even though Rho1 inhibition is persistent (Methods), we do not favor the explanation that ectodermal shortening is directly influenced by Rho1 inhibition. Instead, we hypothesized that this impact was an indirect consequence of what happened near the constriction domain upon Rho1 inhibition. Interestingly, we have previously observed that Rho1 inhibition results in moderate but detectable relaxation of the flanking non-constricting cells in some Late Group embryos (cell 9 in original Figure 5f – j; original Figure 6—figure supplement 1). This mild and transient relaxation, although not sufficient to cause a pause in the invagination of the constricting cells (original Figure 5f), may result in a slight “ventral-to lateral” flow of cell volume, thereby causing a temporary increase in ectodermal cell length. To test this possibility, we examined the correlation between the extent of ectoderm cell length increase and the reverse movement of flanking cells after Rho1 inhibition and detected a positive correlation (R2 = 0.667, n = 5 embryos, plot not shown). Although we were not able to draw solid conclusion on this correlation due to the small sample size, this result is consistent with the idea that Rho1 inhibition does not directly impact ectoderm shortening but transiently and mildly influences ectoderm cell length due to relaxation of cells adjacent to the constricting domain.

6. The manuscript has valuable data on cell behaviors in the lateral ectoderm. But, the presentation is entirely focused around the idea of 'compression', and no alternatives discussed. One such alternative, could be the impact of germband extension on deepening the already formed furrow. The cellular flow of germband has a component directed towards the ventral pole, possibly allowing cells to flow into furrow, that has already formed. This extension will lead to the observed apicobasal shortening of lateral ectoderm cells, and deepening of the fold, but requires no mechanical bistablity. It would further be consistent with lack of fold and the described cell shapes at the apical surface in Snail and opto RhoDN experiment.

We thank the reviewers for raising this very interesting possibility. However, based on previous literature and our own observations, we do not favor the model that germband extension contribute to the transition from apical constriction to invagination. Although there is a prominent DV-directed cellular flow of the lateral ectoderm during ventral furrow, this cellular flow could be due to ventral furrow pull or germ band extension, or both. Since germ band extension is a process of convergent extension, the cellular flow along the DV axis (the “convergence”) is expected to be associated with an accompanying cellular flow along the AP axis (the “extension”). To ask whether the DV tissue flow is due to germband extension or due to ventral furrow pull, we compared the timing of ventral furrow invagination and the ectoderm cell movement in the wildtype embryos by examining the data from original Figure 8. While the DV movement of the ectoderm occurred at about the same time as VF invagination, the AP movement of the ectoderm did not start until ~ 20 min after the onset of apical constriction, when the ventral furrow has nearly fully invaginated (with an invagination depth D larger than 40 μm; Figure 8c and 8f). For comparison, the transition phase (Ttrans) of VF formation is at about ~6 min after the onset of apical constriction, and the invagination depth D at Ttrans is ~ 7 μm (Figure 4). These results are most consistent with a model that the DV movement of the ectoderm before T = 20 min is due to ventral furrow pull, instead of germ band extension. Along the same vein, a similar comparison between ectodermal shortening and ectodermal tissue flow indicates that ectodermal shortening occurs before the AP movement (Figure 8c), suggesting that germband extension does not account for ectodermal cell shortening before T = 20 min. These results are consistent with the notion in the literature that germband extension starts when ventral furrow formation is about to complete (Lye et al., 2015). Finally, we examined the cellular flow in Late Group embryos and observed little AP tissue movement during the course of VF invagination (Figure 6—figure supplement 2). Together, these results suggest that germband extension is unlikely to be the cause of the binary tissue response observed in our optogenetic experiment or the cause of ectodermal shortening before T = 20 min. That being said, our results do not rule out the possibility that germband extension may contribute to the closing of VF at the final stage of VF invagination, which will be an interesting topic for future research. In addition, while we consider mechanical bistability as the most parsimonious explanation of the observed binary tissue response, we could not formally rule out other possible mechanisms.

The above discussion about germband extension is included in the revised manuscript (Results: Page 15; Discussion: Page 25 – 26).

Regarding the comment on the snail mutant and early Opto-Rho1DN experiments:

The mechanical bistability model is not in odd with the lack of tissue folding in snail mutant and the early Opto-Rho1DN experiments. In our proposed model, ectodermal compression poises the mesoderm for invagination (i.e., transition from the initial stable equilibrium to the final, fully invaginated stable equilibrium); however, invagination does not happen instantaneously due to a high energy barrier between the two stable configurations. We propose that apical constriction provides the mechanism for the system to overcome this energy barrier and allows the tissue to invaginate. In this process, apical constriction is necessary to bring the system to a transitional state but is dispensable for the subsequent transition into the final fully invaginated state. In the snail mutant, the ventral epithelium is “stuck” at the initial configuration even in the presence of ectodermal compression due to the lack of mechanical input (“apical constriction”) to allow it to overcome the high energy barrier. Such a system is analogous to a bistable light switch. The switch lever has two stable positions (“on” and “off); however, the lever will not spontaneously switch between the two states unless there is an energy input from a push.

We have modified the text in the revised manuscript to make our points more explicit. In particular, we emphasize that in the model/mechanism we propose, apical constriction provides the essential trigger for mesoderm buckling in the presence of ectodermal compression.

7. Stress is needed to build up for the proposed buckling by compression. However, the Snail experiments clearly demonstrate that no buckling of mesoderm occurs when cell behaviors in the mesoderm are perturbed. Stating mesoderm buckles due to compression from the ectoderm is, therefore, misleading and has not been demonstrated with an experiment. Any mention of this interpretation should be confined to the discussion.

As elaborated in our response to Essential Revisions #6, we believe a model where apical constriction and ectodermal compression jointly mediate mesoderm buckling is compatible with the observation in the snail mutant embryos. In short, in our model, the level of compressive stress from the ectoderm is not sufficient to trigger mesoderm buckling on its own, but it can facilitate mesoderm buckling when the mesodermal cells undergo apical constriction. That being said, we agree with the reviewers that we have not tested this mechanism experimentally. In the revised manuscript, we emphasize that mesoderm buckling facilitated by ectodermal compression is a proposed mechanism that has only been tested by computer modeling.

8. In the discussion, the statement "Using computer modeling, we further demonstrated that mechanical bistablity in the mesoderm can arise from an apicobasal shrinkage of the ectoderm, which generates in-plane compression as the cell volume remains conserved" is misleading. This needs to be clarified, if the authors wish to raise the idea of bistability in the Discussion section.

Following this suggestion, we modified the sentence as following to clarify our point:

“Using computer modeling, we tested a possible mechanism based on the analogy of buckling of a compressed elastic beam induced by an indentation force. […] In this model, apicobasal shrinking of the ectoderm is expected to generate in-plane compression due to cell volume conservation.”

9. While the authors consistently claim precise agreement between the model and data, it remains unclear to what extend this is the case. Visuals of simulations are provided. But there are no quantitative comparisons found that directly compare a model result with a corresponding measurement. Therefore, such claims (e.g. page 14 "In particular, the transitional state of the tissue revealed in the simulation is nearly identical to that identified in our ontogenetic experiments") have to be toned down (e.g. looks visually similar).

Following the reviewers’ suggestion, we have modified the statement regarding the comparison between the modeling results and the actual observations in the embryos:

“In particular, the transitional state of the tissue revealed in the simulation is visually similar to that identified in our optogenetic experiments.”

10. The quantitative analysis shown in figure 8 appears inconsistent with the descriptions. First, the authors refer to a rate of volume reduction, but show volume. Moreover, rate of volume reduction in WT appears consistently different from snail, yet is described as very similar. Such strong claims should either be toned down or backed up with a statistical significance test.

We thank the reviewers for pointing this out. In the original text, we mentioned the “rate of volume reduction” one time in the following sentence: “In addition, the rate of Volec reduction was similar between the wildtype and snail embryos, although the snail embryos on average had a smaller Volec to start with (Figure 8d and f).”. In the actual plot, we displayed the volume over time, but did not display the rate of volume change directly. We apologize for the confusion.

To address this issue and also the reviewers’ comments regarding the comparison between the wildtype and snail mutants, we now provide quantification of the data to further support our statement. We fitted the descending part of the volume curve into straight lines and calculated the rate of volume reduction. The result is shown in Figure 8h in the revised manuscript. We found that there is no significant difference between the two groups (two-sample two tailed student t-test).

11. In the text page 20, the authors describe "In the wild type embryos, the compression promotes ventral furrow invagination, which in turn functions as a 'sink' to facilitate the movement of the ectoderm in the ventral direction". There is no experimental evidence provided for compression, and therefore this statement is speculation. Please rephrase.

Following the reviewers’ suggestion, we rephrased the description as follows:

“The altered pattern of cell movement in snail embryos could be explained by a combined effect of ectodermal compression and lack of ventral furrow invagination (Figure 8f). In this hypothetical scenario, the compression promotes ventral furrow invagination in wildtype embryos, which in turn functions as a “sink” to facilitate (and thereby bias) the movement of the ectoderm in the ventral direction.” (Page 22)

During the revision of this manuscript, we became aware of an earlier study that investigated the stage-specific role of Rok during ventral furrow formation by timed injection of Rok inhibitor (Krajcovic and Minden, 2012). In this work, the authors presented evidence that VF formation is less sensitive to injection of Rok inhibitor once the furrow started to form a small apical indentation. Due to technical limitations, the exact timing of Rok inhibition and its impact on the 3D morphology of the furrow were not provided in this study. However, their major observation is consistent with our results. We now cited this work in the Introduction of the revised manuscript (Page 4) to acknowledge the finding from this pioneer work.

Reviewer #1 (Recommendations for the authors):

[…] (1) The manuscript offers no support for mechanical bistablity assumption. That purely elastic materials can buckle under compression is well established. But, the authors supply data that argues against mechanical bistablity in this system.

– Experimental data does not go beyond correlation, and yet the mechanism presented claims a causal role of ectoderm compression for ventral furrow folding. The original manuscript attempted to provide experimental data in support of the mechanical instability model. The current figures describe cell shapes in 3D, but there is no test of causality offered. This is somewhat puzzling, as the optogenetic tool should also function in the lateral ectoderm.

As described in our response to Essential Revision #6, we agree with the reviewers that we could not entirely rule out other possible mechanisms that may result in the observed binary tissue response upon acute Rho1 inhibition. However, we consider mechanical bistability as the most parsimonious explanation of the observed binary tissue response. In addition, while it has been well established that elastic material can buckle under compression, the notion that tissue can undergo buckling through a joint action of global compression and local apical constriction and will display bistable characteristics during the folding process, to the best of our knowledge, have not been previously proposed. We therefore believe that such a model would provide new perspective to the understanding of tissue folding process.

To address the reviewer’s concern about the lack of causal role of ectodermal compression for ventral furrow folding, in the revised text, we clearly distinguish between the proposed model and the description of experimental observations.

– The model should be described as a theory. In its current form it is hard to distinguish from the descriptions of experiments. It should be clearly labeled that it is purely elastic, and that it neglects the well known viscous properties of tissues that dominate on the scale of at least 4 minutes and beyond.

Following reviewer’s suggestion, we carefully distinguished between descriptions of experimental observations and conclusions drawn from the modeling analysis. We have also emphasized the difference between the actual embryo mechanical properties and that of the model we used in our work, as elaborated in our response to Essential Revision #2.

– It is not clear to what extend the material in figure 8 supports the main argument. Instead, it seems to show the opposite. Lateral ectoderm shorting happens wether or not ventral furrow forms. This is a clear demonstration that the proposed mechanical bistability assumption is not able to drive tissue folding. Instead, these results suggest that folding needs to occur through an independent mechanism. Deepening of the fold could be generated by another mechanism (see below).

We thank the reviewer for raising this interesting and important point. As elaborated in our response to Essential Revision #7, we believe that compression alone is not sufficient to trigger tissue folding due to the high energy barrier between the two stable tissue configurations. The lack of tissue invagination in snail mutant is consistent with our notion that apical constriction is critical for letting the tissue to overcome this energy barrier and invaginate.

– Compression, as indicated by the authors, implies reduction in apical surface area of compressed cells, which is not shown in Snail or early opto RhoDN experiment. Early opto RhoDN experiments are described as heterogeneous cell morphology, but not further analyzed because of technical challenges. It is not clear how this is a technical problem, and not an issue of data interpretation. Heterogeneous apical cell area is consistent with cell shear, but not with external compression.

We agree with the reviewer that compression would be expected to result in a reduction in apical surface area of the compressed cells. The extent of this reduction would be jointly dependent on the magnitude of compression and the “resistance” – the rigidity of the cells. We did not directly measure apical surface area in snail mutant embryos due to lack of a membrane marker in the line we used. Based on our measurement of lateral cell movement, we anticipate that ventral cells will reduce their size mainly along the DV axis (since there is no sign of cell loss from the ventral surface). It would be interesting to test this in the future by direct measurements. For the early Opto-Rho1DN experiment, the apical cell area in the ventral region became heterogeneous ~6 min after the onset of gastrulation, which made it difficult to appreciate the cell aspect ratio change in individual cells. However, following the same reasoning for the snail mutant, we anticipate that the average cell aspect ratio (DV/AP) would reduce upon the observed ventral movement of the lateral cells. The cause of the heterogeneous apical cell area in the ventral region is unclear. Rho1 inhibition prevented activation of apical actomyosin contractility, which would be expected to cause disassembly of adherens junctions in ventral cells due to the expression of Snail (Weng and Wieschaus, 2016). This may in turn cause destabilization of the apical domain of the cells. We propose that what occurred in the ventral cells in early Opto-Rho1DN treated embryos was a combination of compression (as indicated by the ventral movement of the lateral cells, Figure 9f-i) and Snail-mediated loss of cell adhesion. It would be interesting to test these ideas in the future.

2) Discussion of model limitations and alternative scenarios.

– I congratulate the authors on their observation in figure 4b. It seems reasonable to further analyze this interesting phenomenon, and study the possible impact of tissue tissue interactions. In doing so, the manuscript would benefit from an open approach.

– The manuscript has valuable data on cell behaviors in the lateral ectoderm. But, the presentation is entirely focused around the idea of 'compression', and no alternatives discussed.

– One such alternative, could be the impact of germband extension on deepening the already formed furrow. The cellular flow of germband has a component directed towards the ventral pole, possibly allowing cells to flow into furrow, that has already formed. This extension will lead to the observed apicobasal shortening of lateral ectoderm cells, and deepening of the fold, but requires no mechanical bistablity. It would further be consistent with lack of fold and the described cell shapes at the apical surface in Snail and opto RhoDN experiment.

We thank the reviewer for the suggestion and for raising the very interesting hypothesis about germband extension. As elaborated in our response to Essential Revision #6, our current data and previous published observations do not seem to support a major role of germband extension in triggering VF invagination. As suggested by the reviewer, we now included these discussions in the revised manuscript to broaden the discussion on the possible mechanisms that may lead to the binary tissue response observed in Figure 4.

– The timescale of elasticity is very short compared to the 20 minutes of ventral furrow. As pointed out by the authors, the cited paper by Doubrovinsky provides an estimate for the transition to viscosity within 4 minutes. It is one of the longest currently published timescales for this process. As the authors clearly demonstrated turnover of the actomyosin cytoskeleton is very fast, further indicating the four minutes estimate is an upper bound of what is to be expected for these cells.

But, even if it is as long as four minutes, viscosity means, stresses will dissipate. Stress however is needed to build up for the proposed buckling by compression. Snail experiments clearly demonstrate that no buckling of mesoderm occurs when cell behaviors in the mesoderm are perturbed. Stating mesoderm buckles due to compression from the ectoderm is misleading, and has not been demonstrated with an experiment. Any mention of this interpretation should be confined to the discussion.

We thank the reviewer for raising the important question about the elasticity assumption. On the one hand, previous biophysical experiments suggest that the tissue display elastic response in the time scale of at least four minutes (Doubrovinski et al., 2017). On the other hand, rapid remodeling of cytoskeleton would be expected to rapidly dissipate elastic energy. Tissue buckling has been previously described in processes that have longer time scales than ventral furrow formation (Nelson, 2016). How observations at the molecular level link to the tissue level mechanical properties remains only partially understood and would be an important direction for future research. As elaborated in our response to the Essential Revision #7, we consider the proposed mechanism of tissue buckling by apical constriction and compression as the most parsimonious explanation for the binary tissue behavior we observed, but we also acknowledge that future experiments are needed to further support the proposed model. In the revised manuscript, we clearly indicate that buckling by compression is a proposed model that requires future experiments to further validate.

– In the discussion, the statement "Using computer modeling, we further demonstrated that mechanical bistablity in the mesoderm can arise from an apicobasal shrinkage of the ectoderm, which generates in-plane compression as the cell volume remains conserved" is misleading. The authors neither showed that ectoderm compresses mesoderm, nor is it a novel result that elastic systems can buckle under compression. This needs to be clarified, if the authors wish to raise the idea of bistability in the Discussion section.

Please see our response to the Essential Revision #7 for details.

– While the authors consistently claim precise agreement between the model and data, it remains unclear to what extend this is the case. Visuals of simulations are provided. But there are no quantitative comparisons found that directly compare a model result with a corresponding measurement. Therefore, such claims (e.g. page 14 "In particular, the transitional state of the tissue revealed in the simulation is nearly identical to that identified in our ontogenetic experiments") have to be toned down (e.g. looks visually similar).

Please see our response to Essential Revision #9 for details.

Reviewer #2 (Recommendations for the authors):

Guo et al. have revised their paper by following the reviewer's suggestion.

The science presented is now more solid and merits publications after addressing the following 4 points:

1) In the new version of the paper, Guo and colleagues point out the fact that F-actin in ectoderm cells is not affected by RhoDN optogenetic activation. This is quite puzzling and therefore merits at least further discussion.

This is indeed a very interesting point. Please see our response to Essential Revision #4 for details.

2) In Figure 4A and in many other experiments, RhoDN is activated both in the mesoderm and in the ectoderm. Therefore, by following the logic of the authors model, ectoderm pushing is not dependent on Rho signaling. In other terms, while mesoderm cell shortening depends on Rho, ectoderm cell shortening is Rho independent. This also is quite surprising thus merits further discussion.

We thank the reviewer for this important comment. Please see our response to Essential Revision #5 for details.

3) For Figure 9f the authors have opted to track an ectoderm cell over time to demonstrate ectoderm displacement. This is fine, nevertheless the authors should be consistent and perform again the analysis for Figure 9f by using the same analysis protocol implemented for Figure 9a. More precisely the authors should follow a cell that is located 20 cells away from the midline (not just 10 cell away).

Following the reviewer’s suggestion, we have now tracked the cell movement in snail mutant embryos. Please see our response to Essential Revision #1 for details.

4) At page 19 and eventually in the discussion the authors should emphasize the discrepancy between the model and the in vivo measurements. More precisely they should make clear that along a cross-section in the model the minimum necessary ectoderm cell shortening is 5-10% for 60 cells while in the real embryo the cell-shortening measured is 4-8% for a much smaller number of ectoderm cells closer to the mesoderm (20 cells?).

We thank the reviewer for this suggestion. Please refer to our response to Essential Revision #3 for details.

References:

Denk-Lobnig M, Totz JF, Heer NC, Dunkel J, Martin AC. Combinatorial patterns of graded RhoA activation and uniform F-actin depletion promote tissue curvature. Dev Camb Engl 2021;148:dev199232. https://doi.org/10.1242/dev.199232.

Doubrovinski K, Swan M, Polyakov O, Wieschaus EF. Measurement of cortical elasticity in Drosophila melanogaster embryos using ferrofluids. Proc Natl Acad Sci U A 2017;114:1051–6.

Garcia De Las Bayonas A, Philippe J-M, Lellouch AC, Lecuit T. Distinct RhoGEFs Activate Apical and Junctional Contractility under Control of G Proteins during Epithelial Morphogenesis. Curr Biol CB 2019;29:3370-3385.e7. https://doi.org/10.1016/j.cub.2019.08.017.

Grevengoed EE, Fox DT, Gates J, Peifer M. Balancing different types of actin polymerization at distinct sites: roles for Abelson kinase and Enabled. J Cell Biol 2003;163:1267–79.

Homem CCF, Peifer M. Diaphanous regulates myosin and adherens junctions to control cell contractility and protrusive behavior during morphogenesis. Development 2008;135:1005–18. https://doi.org/10.1242/dev.016337.

Krajcovic MM, Minden JS. Assessing the critical period for Rho kinase activity during Drosophila ventral furrow formation. Dev Dyn Off Publ Am Assoc Anat 2012;241:1729–43. https://doi.org/10.1002/dvdy.23859.

Lye CM, Blanchard GB, Naylor HW, Muresan L, Huisken J, Adams RJ, et al. Mechanical Coupling between Endoderm Invagination and Axis Extension in Drosophila. PLoS Biol 2015;13:e1002292. https://doi.org/10.1371/journal.pbio.1002292.

Mason FM, Tworoger M, Martin AC. Apical domain polarization localizes actin-myosin activity to drive ratchet-like apical constriction. Nat Cell Biol 2013;15:926–36.

Nelson CM. On Buckling Morphogenesis. J Biomech Eng 2016;138:021005. https://doi.org/10.1115/1.4032128.

Polyakov O, He B, Swan M, Shaevitz JW, Kaschube M, Wieschaus E. Passive mechanical forces control cell-shape change during Drosophila ventral furrow formation. Biophys J 2014;107:998–1010.

Weng M, Wieschaus E. Myosin-dependent remodeling of adherens junctions protects junctions from Snail-dependent disassembly. J Cell Biol 2016;212:219–29. https://doi.org/10.1083/jcb.201508056.

https://doi.org/10.7554/eLife.69082.sa2

Article and author information

Author details

  1. Hanqing Guo

    Department of Biological Sciences, Dartmouth College, Hanover, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Validation, Visualization, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8722-6253
  2. Michael Swan

    Department of Molecular Biology, Princeton University, Princeton, United States
    Contribution
    Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Bing He

    Department of Biological Sciences, Dartmouth College, Hanover, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Software, Supervision, Validation, Visualization, Writing - original draft, Writing - review and editing
    For correspondence
    Bing.He@Dartmouth.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8564-0933

Funding

National Institute of General Medical Sciences (ESI-MIRA R35GM128745)

  • Bing He

American Cancer Society (#IRG -82-003-33)

  • Bing He

Cystic Fibrosis Foundation (STANTO15R0)

  • Bing He

National Institute of Diabetes and Digestive and Kidney Diseases (P30-DK117469)

  • Bing He

Centers of Biomedical Research Excellence (P20-GM113132)

  • Bing He

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

The authors thank members of the He lab and the Griffin lab at Dartmouth College for sharing valuable thoughts during this work; James Moseley and Magdalena Bezanilla for providing constructive suggestions and comments on the manuscript; Ann Lavanway for imaging support. The authors thank the Wieschaus lab and the De Renzis lab for sharing reagents, Oleg Polyakov for sharing the code for the energy minimization-based vertex model, and the Bloomington Drosophila Stock Center for fly stocks. This study is supported by NIGMS ESI-MIRA R35GM128745 and American Cancer Society Institutional Research Grant #IRG-82-003-33 to BH. The study used core services supported by STANTO15R0 (CFF RDP), P30-DK117469 (NIDDK P30/DartCF), and P20-GM113132 (bioMT COBRE).

Senior Editor

  1. Utpal Banerjee, University of California, Los Angeles, United States

Reviewing Editor

  1. Michel Bagnat, Duke University, United States

Reviewers

  1. Sebastian J Streichan, University of California, Santa Barbara, United States
  2. Magali Suzanne, Centre de BIologie Integrative (CBI), France

Publication history

  1. Preprint posted: March 19, 2021 (view preprint)
  2. Received: April 3, 2021
  3. Accepted: February 22, 2022
  4. Accepted Manuscript published: February 23, 2022 (version 1)
  5. Version of Record published: March 4, 2022 (version 2)

Copyright

© 2022, Guo et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Hanqing Guo
  2. Michael Swan
  3. Bing He
(2022)
Optogenetic inhibition of actomyosin reveals mechanical bistability of the mesoderm epithelium during Drosophila mesoderm invagination
eLife 11:e69082.
https://doi.org/10.7554/eLife.69082

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