Mechanics of development

Abstract

Mechanical forces are integral to development, from the earliest stages of embryogenesis to the construction and differentiation of complex organs. Advances in imaging and biophysical tools have allowed us to delve into the developmental mechanobiology of increasingly complex organs and organisms. Here, we focus on recent work that highlights the diversity and importance of mechanical influences during morphogenesis. Developing tissues experience intrinsic mechanical signals from active forces and changes to tissue mechanical properties, as well as extrinsic mechanical signals, including constraint and compression, pressure, and shear forces. Finally, we suggest promising avenues for future work in this rapidly expanding field.

Introduction

Much of developmental biology has sought to uncover the roles of morphogens and the molecular programs that drive development (Briscoe and Small, 2015; Freeman and Gurdon, 2002; Kicheva and Briscoe, 2015). More recently, attention has turned to studying how mechanical forces influence developing tissues and to understanding the physical mechanisms of morphogenesis (Anlas and Nelson, 2018; Mammoto and Ingber, 2010; Stooke-Vaughan and Campas, 2018). These two schools of thought are not necessarily separate; indeed, local mechanical properties and their effects on tissue topology can shape morphogen fields (Manning et al., 2015; Shyer et al., 2015), and cell displacements can change how a tissue experiences a static molecular gradient (Nerurkar et al., 2019).

Mechanical forces influence all stages of development, from the early embryo, to gastrulation and establishment of the body plan, to organogenesis. Pioneering reviews laid the foundation for our current appreciation of the role of mechanical forces in development (Beloussov et al., 1994; Keller et al., 2003; Wolpert, 1971), and over the past 10 years, advances in animal models, live-imaging approaches, and biophysical measurements have shed light on the mechanics of morphogenesis (Ayad et al., 2019). In particular, these advances have allowed us to extend principles of developmental mechanobiology first established primarily in invertebrate embryos (Heisenberg and Bellaiche, 2013; Lecuit and Lenne, 2007; Mammoto and Ingber, 2010) to the morphogenesis of larger, more complex, and less accessible organs and organisms.

Here, we provide an overview of the different kinds of mechanical forces that drive morphogenesis and highlight recent progress in understanding how they regulate developmental processes from the earliest stages of embryogenesis to the finishing touches of organogenesis. We include examples of intrinsic mechanics driving morphogenesis, including the generation of active forces and tuning of tissue fluidity, as well as examples of extrinsic mechanical signals, such as physical constraints and microenvironment-mechanics, static and dynamic pressure, and shear forces from fluid flow. In this review, we focus on how mechanical forces directly influence tissue form. However, it is important to note that mechanical forces also influence another key aspect of development, cellular differentiation: physical forces activate intracellular signaling cascades or change nuclear envelope mechanics to regulate the activation or nuclear localization of mechanosensitive transcription factors, leading to downstream changes in gene expression, and altering cell behavior and fate (Hampoelz and Lecuit, 2011; Kumar et al., 2017; Vining and Mooney, 2017). These changes at the nuclear level could indirectly influence the material properties of the tissue and, thus, changes in tissue form.

Tension and active forces

Actomyosin networks coupled to cell-cell and cell-extracellular matrix (ECM) adhesions are responsible for generating the forces required for cells to pull on their neighbors or on their underlying substratum (Hoffman et al., 2011). Actomyosin contractions can locally constrict parts of the cell, as in apical constriction, in order to change cell shape, or allow it to exert traction in order to move over a surface, as in cell migration. These active forces at the level of single cells can be translated into dramatic changes in shape at the level of the tissue. Actomyosin networks often contract with pulsatile dynamics. Indeed, pulsed actomyosin contractions are an evolutionarily conserved mechanism for driving tissue morphogenesis. These contractions promote ventral-furrow ingression in Drosophila (Martin et al., 2009), compaction in the mouse blastocyst (Maitre et al., 2015), and have even been observed in collectives of unicellular organisms that form sheets that can fold in response to light (Brunet et al., 2019).

Active forces generated by actomyosin networks orchestrate cell-shape changes required for transforming the early mouse embryo from an agglomeration of roughly spherical cells into a compacted embryo, and then into a hollow blastocyst. As the embryo compacts, the angle between adjacent cells at the periphery of the embryo increases and the surface tension at the cell-medium interface increases (Fig. 1A, Table 1) (Maitre et al., 2015). Consistently, actomyosin activity is enriched at cell-medium compared to cell-cell interfaces, and the increase in tension at the cell-medium interface requires actin polymerization and myosin activity. Strikingly, this increase in tension is cell autonomous (occurs even in dissociated cells), and is accompanied by oscillatory cell deformations and actin accumulation; whether the oscillations themselves are required for compaction has yet to be shown. Cells of the 8-cell embryo also extend cadherin-containing filopodia over their neighbors that deform the apical part of cell to drive compaction (Fierro-Gonzalez et al., 2013).

Figure 1. Intrinsic mechanical forces driving morphogenesis.

(A) During compaction of the 8-cell embryo, tension at the cell-medium interface increases, changing the angles between cells. The increase in tension is accompanied by pulsatile actin dynamics at the cell cortex. (B) Apical actin rings in the 16-cell mouse embryo expand towards intercellular junctions to seal the blastocyst. (C) Hindgut morphogenesis in the chick embryo requires the gut endoderm to fold over onto itself to form the gut tube. This movement is generated by collective cell migration resulting from a gradient of cell contractility that interacts with a stable morphogen gradient to drive tissue flow. (D) Deep cells fluidize at the onset of zebrafish epiboly as they undergo divisions and lose integrity of their cell-cell contacts. As a result, the expanding EVL compresses the deep-cell layer to drive blastoderm thinning.

Table 1.

Physical concepts and equations related to the mechanics of development.

Physical concepts	Relevant quantities	Equations	Examples of related biological processes
Viscoelasticity	stress (σ) elastic modulus (E) viscosity (μ) displacement (ε$\epsilon$) time (t)	elastic element $\sigma =E\epsilon$ viscous element $\sigma =\mu \frac{d\epsilon }{dt}$	cortical tension and recoil after laser ablation (Samarage et al., 2015; Zenker et al., 2018)
Shape index and the jamming transition	shape index ($\overline{p}$) cell area (A) cell perimeter (P) critical shape index (p0)	$\overline{p}=\frac{P}{\sqrt{A}}$ if $\overline{p}>p_0$ , the tissue is fluid-like if $\overline{p}<p_0$, the tissue is solid-like	Drosophila germband extension (Wang et al., 2020), zebrafish tailbud elongation, avian gastrulation (Firmino et al., 2016; Saadaoui et al., 2020)
Differential stiffness driving buckling of a tube attached to a sheet	buckling wavelength (λ) elastic modulus of the tube (Et) elastic modulus of the sheet (Es) moment of inertia of the tube (It) height of the sheet (h)	$\lambda \propto {\left(\frac{E_t{I}_t}{E_{s}h}\right)}^{1/3}$	looping of the avian gut (Savin et al., 2011)
Differential growth driving buckling of a sheet attached to a viscoelastic foundation	buckling wavelength (λ) elastic modulus of the sheet (E) critical in-plane load (N, scales with growth rate)	$\lambda \propto \sqrt{\frac{E}{N}}$	branching of the mesenchyme-free murine airway epithelium (Varner et al., 2015)
Pressure in a spherical vessel	stress (σ) pressure (P) radius of the vessel (R) thickness of the walls (t)	$\sigma =\frac{PR}{2t}$	pressurization of the blastocyst (Chan et al., 2019; Dumortier et al., 2019; Leonavicius et al., 2018), zebrafish inner ear morphogenesis (Hoijman et al., 2015; Mosaliganti et al., 2019)
Young-Laplace equation as applied to micropipette aspiration	surface tension (γ) applied pressure (P) radius of curvature (Rc) micropipette radius (Rp)	$\gamma =\frac{P}{2\left(\frac{1}{R_p}-\frac{1}{R_c}\right)}$	surface tension at the cell-medium interface of early mouse embryo (Maitre et al., 2015)
Fluid shear stress	shear stress (τ) viscosity (µ) fluid velocity (u) height above the boundary (y)	$\tau =\mu \frac{du}{dy}$	Zebrafish heart valve morphogenesis (Boselli et al., 2017)

From the 8-cell to the 16-cell stage, a subset of cells end up in the center of the embryo. These inner cells give rise to the embryonic tissues and the primitive endoderm, whereas the outer cells (the trophectoderm) give rise to extraembryonic tissues that encapsulate the embryo. Inner cells are positioned by apical constriction (Maitre et al., 2016; Samarage et al., 2015). Myosin is enriched at constricting intercellular junctions, and laser ablation revealed more recoil (a proxy for cortical tension) along junctions between constricting cells and their neighbors than between non-constricting cells. Further, ablations oriented perpendicular to junctions between constricting cells caused more recoil of the cortical actin network than ablations parallel to junctions, demonstrating that the apical cortex of constricting cells is under greater tension. Specification and positioning of trophectoderm cells may also be mechanically regulated: keratin-rich intermediate filaments stabilize F-actin within the apical domain, and additionally serve as an asymmetrically inherited structural component that predicts trophectoderm fate (Lim et al., 2020).

During the next major phase of mouse embryogenesis (16- to 32-cell stages), the embryo transforms from a solid mass of cells (called the morula) to a fluid-filled blastocyst with an inner cell mass (ICM) and an outer layer of cells called the trophoblast. During this transition, the embryo undergoes further actomyosin-dependent remodelling in order to seal junctions between the outer cells and to facilitate the formation of a fluid-filled cavity within the center of the embryo called the blastocoel (Zenker et al., 2018). In the apical (outward-facing) domains of the cells of the 16-cell embryo, actin rings assemble and passively expand until they reach cell-cell junctions (Fig. 1B). Actin ring expansion appears to be driven indirectly by constriction of the basal (inward-facing) portion of the cells, which causes the apical portion of the cell to expand and push the actin ring outward. Active processes are then initiated: myosin becomes enriched at the contact between the actin ring and the membrane and helps to capture rings in neighboring cells and pull them to the same contact point. The two actin rings are then zippered together in a myosin- and tension-dependent process that helps to seal the cell-cell junction by recruiting and stabilizing adherens and tight junction components.

Active forces drive the formation of increasingly complex tissue architectures throughout embryogenesis. Actomyosin activity is required for several aspects of neural tube closure including apical constriction, apical-basal cell lengthening, and cell migration (Nikolopoulou et al., 2017), and for formation of the gut tube (Nerurkar et al., 2019). After gastrulation, the endoderm (which gives rise to the digestive tract and lungs) is transformed from a sheet of cells into a tube. Morphogenesis of the hindgut from the endoderm during embryonic development of the chick involves a gradient in contractile forces that drives coordinated migration of the gut endoderm sheet, allowing it to fold over on itself to generate the gut tube (Nerurkar et al., 2019) (Fig. 1C). Cells of the endoderm are exposed to a gradient of fibroblast growth factor 8 (FGF8) that is highest at the posterior end of the tissue. Posterior-most cells contract, pulling their trailing neighbors into the region of high FGF8 concentration, causing them to contract as well. This positive feedback generates a gradient of contractile forces that drives tissue elongation.

Actomyosin-driven apical constriction is responsible for generating tissue topology in many different organs. Pulsed actomyosin contractions promote apical constriction during invagination of the Drosophila salivary gland (Chung et al., 2017). Suprisingly, apical constrictions are not strictly required for invagination in this context (tissue-wide compressive forces can compensate to induce buckling – a concept discussed below), but are necessary to achieve the correct tissue morphology. In the avian lung, lateral branches form by apical constriction of epithelial cells in the primary bronchus (Kim et al., 2013). This active tissue folding requires growth factor signaling and myosin contractility, but how growth factor signaling elicits localized apical constriction remains unclear. During postnatal development of the murine intestine, crypts form through apical constriction of epithelial cells, and “hinge” cells basally constrict to generate a boundary between crypts and villi (Sumigray et al., 2018). The initial step requires myosin activity, whereas the subsequent step depends upon remodelling of cell-cell adhesion; both are necessary to achieve correct tissue architecture and spacing between villi.

Aside from apical constriction, actomyosin-based contractility can drive other cellular behaviors important for tissue morphogenesis. For example, contractility drives condensation of dermal progenitors in the avian skin, inducing a mechanosensitive response in the neighboring epidermal cells to initiate specification of feather follicles (Shyer et al., 2017). In the murine hair follicle, actomyosin-mediated rearrangements of epidermal follicle progenitors lead to rotational cell flows that relocate anterior cells to the periphery of the follicle and posterior cells to the center; these rearrangements are required to establish an asymmetric follicle morphology and the constituent cell fates (Cetera et al., 2018). In this case, active forces drive morphogenesis by promoting cell migration and, further, promote patterned differentiation of the murine hair follicle.

Cellular mechanosensing of physical signals like substratum stiffness and tension can inform fate decisions, even in cells that may not be actively contributing to morphogenetic movements. For example, airway epithelial cells cultured in tubes of developmentally relevant geometries (wider to mimic proximal airways, narrower to mimic distal airways) experience curvature-dependent tension that is interpreted to dictate the expression of either proximal or distal fate markers (Soleas et al., 2020). Future work will elucidate whether and how the active forces that generate complex tissue topologies are also directly sensed and interpreted by differentiating cells.

Fluid-to-solid transitions

Fluidity and solidity have recently emerged as useful descriptors of cell migration that can predict whether cells migrate independently or collectively (or somewhere in between). In fluid-like tissues, cells can rearrange and move past their neighbors, whereas in solid-like tissues, neighbors are maintained and rearrangements are minimal (Bi et al., 2015; Schotz et al., 2013). The transition between these two states is called the jamming transition and can be predicted in diverse biological systems by changes in cell geometry (Atia et al., 2018; Bi et al., 2015; Wang et al., 2020). This framework has been applied to studies of cancer (Haeger et al., 2014) and asthma (Park et al., 2015), and is being increasingly applied to developmental biology.

In simple two-dimensional cell sheets, the jamming transition is predicted to occur when the shape index, a simple parameter that depends on the cell perimeter and area, surpasses a specific value (Table 1). Tissues composed of hexagonal cells have a shape index below this threshold and exhibit solid-like behavior, whereas those with irregularly-shaped or elongated cells have a shape index above the threshold and exhibit fluid-like behavior (Bi et al., 2015). However, recent work in Drosophila germband extension has shown that the parameters regulating jamming may depend on whether the tissue is experiencing isotropic or anisotropic forces (Wang et al., 2020). Convergent extension is an anisotropic process, and cell shape is insufficient to predict the jamming transition that marks the onset of tissue flow in the germband. An additional parameter describing cell-shape alignment is required to predict these changes and altered cell behavior in mutants: snail mutants, which fail to undergo mesoderm invagination, have a lesser degree of cell-shape alignment and a delayed onset of tissue flow.

During epiboly, the first stage of zebrafish embryogenesis, the blastoderm (comprised of an epithelial enveloping layer (EVL) covering deep cells) spreads over the massive yolk cell by expanding the EVL and radially contracting via intercalation of deep cells (Bruce, 2016; Morita et al., 2017). The active forces that drive this process are well appreciated; additionally, recent work showed that tuning the mechanical properties of deep cells allows for their passive deformation at the onset of epiboly before radial intercalation has begun (Petridou et al., 2019). Deep cells lose cell-cell junctional integrity as they divide, leading to lower tissue viscosity and fluidization and allowing the expanding EVL to compress the deep cell layer and drive blastoderm thinning (Fig. 1D).

Axis elongation in zebrafish embryos involves extensive collective cell migration and depends upon tissue fluidity (Lawton et al., 2013; Mongera et al., 2018). Cells of the tailbud migrate posteriorly in a coherent fashion until they reach the tip, where their movement becomes less coherent, a hallmark of fluidization (Lawton et al., 2013). The observed patterns of cell migration point to a role for graded tissue mechanics in regulating axis elongation. Recently, new biophysical tools have been applied to uncover the mechanical properties of the tailbud and have revealed that axis elongation involves a jamming transition along the anterior-posterior axis (Mongera et al., 2018). A deformable, magnetic oil droplet injected into the tailbud was used to measure local stresses (from passive droplet deformations at rest) and mechanical properties (from active droplet deformation upon application of a magnetic field). These experiments revealed that the fluid-like cells at the tip of the tailbud exhibit lower mechanical integrity than the more solid-like anterior cells that comprise the presomitic mesoderm (PSM). Further, recent work has shown that these tissue mechanical properties are a critical component of signal transmission in the tailbud. The cells at the tip of the tailbud form the tailbud organizer and have long been appreciated as an important source of morphogens required for axis elongation. Recent work has also shown that mechanical information can be communicated between the cells of the tailbud, thus extending the influence of the tail organizer cells beyond the range of morphogen diffusion (Das et al., 2019). The transmission of this mechanical information is likely dependent on the fluidity of the tissue.

The zebrafish tailbud has proven useful for uncovering the physical parameters that regulate tissue fluidity in embryonic tissues. Similar to the deep cell layer during early epiboly (Petridou et al., 2019), there are greater spaces between cells at the tip of the tailbud that allow for fluid-like behavior (Mongera et al., 2018). Additionally, cell-cell contact lengths are more variable and fluctuate to a greater extent at the tip of the tailbud, and inhibiting myosin activity abrogates this behavior and solidifies the tissue. Finally, cell-cell adhesion supports tissue coherence and solidity in the tailbud, likely by reducing the space between neighboring cells (Lawton et al., 2013; Mongera et al., 2018).

Jamming transitions in embryogenesis are also regulated by cell division and changes in cell phenotype. During gastrulation in chick embryos, cells of the epiblast exhibit dramatic, counter-rotational flows called “Polonaise movements” to specify the embryonic midline and allow mesendoderm initiation. These rotational movements require cell divisions, without which the epithelial layer is too stable (or solid) for flows to occur (Firmino et al., 2016). Further, recent work combining quantitative live-imaging, mechanical modeling, and laser ablations to infer tissue strains in quail embryos showed that the epiblast is surrounded by a supracellular tensile ring that contracts to drive rotational tissue flows and gastrulation (Saadaoui et al., 2020). The onset of neural crest cell migration requires an epithelial-to-mesenchymal transition, which involves a reduction in cell-cell adhesion. In Xenopus embryos, internalization of N-cadherin allows neural crest cells to fluidize and thus migrate collectively through narrow spaces (Kuriyama et al., 2014).

Thus far, there are few examples of jamming transitions and roles for tissue solidity and fluidity in organogenesis. In the avian lung, fluidity of the pulmonary mesenchyme helps to distribute the large extracellular matrix protein tenascin around extending branches (Spurlin et al., 2019). During murine eyelid closure, collective migration of vast epithelial sheets is driven by intercalations of cells at the eyelid front, which may involve localized changes in tissue fluidity (Heller et al., 2014). Jamming transitions have been shown to occur in airway epithelial remodelling in asthma (Park et al., 2015), raising the possibility that airway epithelial fluidity may also be developmentally regulated as the lung branches and differentiates during embryogenesis.

Physical constraints and microenvironmental mechanics

Developing tissues are subject to a variety of physical constraints, including wrapping by ECM, attachment to a neighboring tissue, and sculpting by surrounding contractile tissues. Physical signals from the tissue microenvironment can elicit cellular behaviors via mechanotransduction (DuFort et al., 2011), and physical constraints can prevent or direct tissue growth, or lead to mechanical instabilities and subsequent buckling of tissues (Nelson, 2016).

Changes to basement membrane composition, stiffness, and mechanical integrity provide physical signals that guide tissue morphogenesis (Dzamba and DeSimone, 2018; Walma and Yamada, 2020). Basement membrane around the post-implantation mouse embryo is remodelled to shape the embryo prior to gastrulation and then locally weakened at the posterior pole of the embryo to allow for extension of the primitive streak (Kyprianou et al., 2020). Initially, perforations in the basement membrane around the epiblast allow growth of the embryo along its long axis. As the anterior visceral endoderm migrates from its position at the base of the embryo to the anterior pole, it locally inhibits matrix metalloproteinases, limiting the domain of perforations to the posterior pole. This local reduction in the mechanical integrity of the basement membrane is critical for gastrulation and extension of the primitive streak. Similar basement membrane perforations have been observed at branch tips in the developing salivary gland and are thought to be important for permitting branch growth (Harunaga et al., 2014).

Alterations to tissue mechanical properties can also serve as physical signals that influence the behavior of neighboring tissues. Neural crest cells in the Xenopus embryo sense stiffening of the underlying head mesoderm tissue via cell-matrix adhesions (Barriga et al., 2018). This stiffening is required for neural crest development, and is itself promoted by morphogenetic movements: convergent extension of the head mesoderm beneath the neural crest cells increases cell density, thus leading to increased stiffness (Barriga et al., 2018). Similar to the example above, stiffening of the underlying endoderm can drive mesenchymal-to-epithelial transition in heart progenitor cells during Xenopus embryogenesis (Jackson et al., 2017). Finally, proliferation generates a stiffness gradient in the developing brains of Xenopus embryos that is sensed by the retinal ganglion cell axons that migrate along the brain surface (Thompson et al., 2019). Formation of the gradient leads to axon turning, directing them to their destination at the optic tectum.

Growing tissues that are subject to compressive forces from the ECM or from neighboring tissues can fold to relieve the resultant strains. In the developing eye of the chick embryo, the ECM constrains the growing optic vesicle and surface ectoderm, forcing them to invaginate and form the concave optic cup (Oltean et al., 2016). During intestinal morphogenesis in the chick embryo, the small intestine grows faster than the attached dorsal mesentery, resulting in compressive forces along the length of the intestinal tube that lead to looping of the gut into stereotyped patterns (Savin et al., 2011). The wavelength of gut looping depends on differential growth rates between the intestine and the dorsal mesentery, which are regulated by BMP signaling (Nerurkar et al., 2017), and can also be explained by the differences in stiffness between these two connected tissues (Table 1) (Savin et al., 2011). When it is isolated from the surrounding mesenchyme, the murine airway epithelium also undergoes predictable buckling behavior with a wavelength that depends on in-plane compressive forces that are proportional to its growth rate (Table 1) (Varner et al., 2015).

Compressive forces that lead to buckling are often driven by tissue growth, and can also be influenced by differences in tissue stiffness or contractility. Constrained growth and mechanical forces imposed by surrounding smooth muscle drive epithelial morphogenesis in the avian intestine (Shyer et al., 2013) and in the murine airway epithelium (Goodwin et al., 2019; Kim et al., 2015). In each of these cases, a uniformly-growing epithelial tube achieves complex topologies (folds and zigzags in the intestine, branches and bifurcations in the lung) due to spatiotemporally patterned differentiation of stiff smooth muscle layers from the surrounding mesenchyme.

Forces from the growth of neighboring tissues direct morphogenesis and can also influence differentiation. For example, smooth muscle differentiation in the chick intestine drives epithelial buckling and is itself regulated by strain that results from growth of the epithelial tube (Huycke et al., 2019). Circumferential and longitudinal smooth muscle layers differentiate sequentially during intestinal morphogenesis. Continuous strain from growth of the epithelium aligns the first layer of smooth muscle cells circumferentially, and subsequently, periodic contractions of this first layer align the second, outer layer of smooth muscle longitudinally (Fig. 2A).

Figure 2. Extrinsic mechanical forces driving morphogenesis.

(A) During chick embryogenesis, growth of the intestinal epithelium generates continuous strain to align the first circumferential layer of smooth muscle cells. Subsequently, periodic contractions of the circumferential smooth muscle layer generate cyclic strain to align the outer longitudinal layer of smooth muscle cells. (B) Pressurized microlumens form between cells of the early mouse embryo and, as they combine, break embryonic symmetry and establish a single fluid-filled cavity within the blastocyst. (C) Blood flow drives morphogenesis of the zebrafish heart valve by activating the shear-sensitive transcription factor KLF2 via mechanosensitive ions channels. Downstream flow-responsive signaling through FGF and WNT is required for maintaining the integrity of the myocardial wall and for driving tissue remodelling. Flow also generates patterns of wall shear stress that predict the direction of endocardial cell convergence towards the AVC.

Static and dynamic pressure

Fluid-filled lumens play diverse mechanical and biochemical roles during development, and several experimental techniques have been used to quantify pressure within them (Chan and Hiiragi, 2020). Lumens form when cells release ions and solutes into the extracellular space, generating an osmotic gradient that directs fluid flow. The fluid within the lumen is contained by tight junctions between cells of the tissue surrounding them, and its expansion generates hydrostatic pressure that exerts stress on the cells surrounding the lumen (Table 1). The mechanisms by which cells sense pressure have yet to be fully elucidated, and likely vary depending on the context. Pressure may deform the cell cortex, leading to mechanotransduction via actomyosin networks or mechanosensitive ion channels. Osmotic gradients generated by fluid-filled lumens can also affect cytoplasmic fluid content and cell volume, thus changing the pH, salt content, and concentration of proteins within the cytoplasm (Chan and Hiiragi, 2020; Guo et al., 2017).

Recently, several studies have beautifully delineated roles for fluid pressure in lumen coalescence, junctional maturation, and cell allocation during blastocyst development (Chan et al., 2019; Dumortier et al., 2019; Leonavicius et al., 2018; Ryan et al., 2019). Pressure build-up in the blastocyst also helps the embryo to hatch from the zona pellucida, a thick membrane that surrounds the embryo prior to implantation (Leonavicius et al., 2018). Prior to blastocyst formation, cells of the mouse embryo produce actin-coated vesicles to secrete fluid into the extracellular space (Ryan et al., 2019). This process produces many microlumens that gradually coalesce to form a single fluid-filled cavity (the blastocoel). The process of lumen coalescence has been likened to hydraulic fracturing: build up of fluid in microlumens between cells mechanically disrupts local cell-cell contacts (Dumortier et al., 2019). Smaller, more pressurized microlumens discharge their contents into neighboring microlumens and, gradually, all microlumens resolve to form the blastocoel (Fig. 2B). As the blastocoel forms and fluid pressure within it increases, trophectoderm cells become stretched and experience higher cortical tension, leading to tight junction maturation and recruitment of the mechanosensory protein vinculin to tight junctions (Chan et al., 2019). Accumulation of keratins in the outer cells of the blastocyst may also help the embryo to withstand the increasing pressure in the blastocoel (Lim et al., 2020).

The effects of pressure and tension are inextricably linked in the earliest stages of mouse embryogenesis and cooperate to pattern the embryo. Asymmetries in tension at cell-cell junctions direct lumen coalescence: fluid in lumens at interfaces with higher tension flows into those at interfaces with lower tension, resulting in final blastocoel positioning at the interface between the less contractile trophectoderm cells and the more contractile ICM cells, rather than between ICM cells (Dumortier et al., 2019). Modulating either pressure or cortical tension also affects blastocoel size, which in turn shifts the balance between inner cell and trophectoderm cell allocations: smaller cavities favor the ICM at the expense of the trophectoderm, whereas larger cavities position more cells within the trophectoderm and fewer within the ICM (Chan et al., 2019). Finally, sorting of the two cells types that comprise the ICM (the epiblast and primitive endoderm) is coupled to lumen expansion from embryonic day (E) 3.5 to E4 and primitive endoderm specification is sensitive to luminal volume (Ryan et al., 2019). The effects of pressure alone on cell-fate specification within the ICM remain unclear; however, manipulating fluid volume changes the availability of soluble factors important for specifying cell fate.

Pressure continues to regulate development at later stages of embryogenesis. During post-implantation mouse development, embryos are subject to external pressures from uterine smooth-muscle contraction. Some of the pressure is absorbed by Reichert’s membrane, a specialized basement membrane that encases embryos immediately following implantation, and the fluid-filled space it creates around the developing embryo, which preserves overall embryo morphology (Ueda et al., 2020). During morphogenesis of the chick brain, the neural tube becomes filled with cerebrospinal fluid (CSF) that exerts pressure on the neuroepithelium (Garcia et al., 2019). Combined with morphogen gradients, this pressure drives growth of the left and right cerebral hemispheres; experimentally decreasing CSF pressure prevents hemisphere outgrowth, whereas increasing pressure leads to enlarged hemispheres. During zebrafish inner ear morphogenesis, hydrostatic pressure increases as the epithelial cells of the otic vesicle change shape and secrete fluid into the cavity (Hoijman et al., 2015); the resultant change in pressure in turn regulates size of the otic vesicle (Mosaliganti et al., 2019). Additionally, patterned actomyosin accumulation along the luminal surface of the otic vesicle allows some cells to resist deformation due to pressure, leading to localized thinning of the epithelium and affecting the shape of the cavity (Hoijman et al., 2015; Mosaliganti et al., 2019).

During development of the mouse lung, static and dynamic pressure drive the formation of complex tissue architecture and influence cell differentiation. Throughout embryogenesis, the lumens of the airways are filled with fluid, and the entire organ is encased within the fluid-filled thoracic cavity; the pressure difference between these two compartments generates transmural pressure across the organ. Embryonic lungs isolated at the early stages of branching morphogenesis can be cultured ex vivo in a “microfluidic chest cavity” designed to apply transmural pressure (Nelson et al., 2017). Under higher transmural pressure, lung explants undergo more branching and have gene expression profiles that more closely resemble those of lungs that develop within the embryo than lungs cultured under lower transmural pressure. Visualizing branching morphogenesis under pressure in this culture system revealed that synchronized airway smooth muscle contractions every ~12 hours preceded near-simultaneous epithelial bifurcations. This observation suggests that smooth muscle contractions could displace luminal fluid towards branch tips, leading to local changes in pressure that induce branching events.

Luminal pressure also drives differentiation of alveolar cell types during the late stages of lung development (Li et al., 2018). Fetal breathing moves luminal fluid towards branch tips, where alveolar progenitors reside. Some of these cells protrude into the mesenchyme and constrict their apical surfaces and are thus protected from changes in fluid pressure. These cells preferentially give rise to the cuboidal surfactant-producing alveolar type II cells, while their neighbours (which feel the full force of fluid pressure) preferentially give rise to the very thin, elongated, gas-exchanging alveolar type I cells.

Fluid flow and shear forces

Cells that line fluid-filled lumens can be subject not only to hydrostatic or dynamic pressure, but also to forces associated with fluid flow. Flow generates shear forces along the walls of the lumen, and the magnitude of these forces scales with fluid velocity (Table 1). A wide range of fluid velocities are present throughout the embryo, from weaker flows generated by beating cilia to very fast flows generated by the beating heart. Mechanosensitive ion channels are key components of flow sensing, and primary cilia have also been suggested to act as force sensors that bend when exposed to shear stress to initiate downstream signaling cascades (Hyman et al., 2017; Spasic and Jacobs, 2017). However, the role of primary cilia is debated, and the entire cell surface could respond to shear forces caused by flow (Ferreira et al., 2019).

Early embryogenesis and establishment of the body plan are orchestrated in part by a small group of cells called the organizer (in amphibians) or the node (in mammals). In particular, left-right patterning is established at the node: 200 to 300 motile cilia at the center of the node generate unidirectional fluid flow, and primary cilia in the cells at the periphery of the node are required to sense this flow (Shinohara et al., 2012; Yoshiba et al., 2012). Flows are highest at the right side of the embryo, increase over the course of development, and reach a velocity of 2-4 μm/s. Whether flow is sensed chemically or mechanically remains a matter of debate (Cartwright et al., 2020). Flow causes spikes in intracellular calcium, supporting the idea that cilia engage in mechanosensing; however, examining this response with higher temporal and spatial resolution revealed that calcium waves do not originate from primary cilia, suggesting that they are not mechanosensing in this context (Delling et al., 2016). It is clear, however, that the mechanisms that detect flow are highly sensitive: mutants in which only two cilia are beating (causing a fluid velocity of 1 μm/s that decays towards the periphery of the node) still generate sufficient flow for left-right patterning (Shinohara et al., 2012).

Midway through embryogenesis, once the heart has formed and started beating, blood flow throughout the developing vascular network becomes a powerful force for tissue morphogenesis (Hyman et al., 2017). Zebrafish embryos with impaired heart development survive much longer than analogous mouse models, making them an excellent model system for studies of cardiovascular development. Flow velocities in the developing zebrafish heart are 3-4 orders of magnitude greater than those at the node (Boselli and Vermot, 2016). Prior to the development of the heart valve, blood flow through the heart frequently reverses direction, leading to oscillatory flow and shear stresses on the valve precursor cells of the endocardium (Fig. 2C). Oscillatory blood flow is sensed by mechanosensitive ion channels expressed in the endocardium, and activates the expression of the shear-responsive gene Kruppel Like Factor 2 (klf2); each of these components are required for valve development (Heckel et al., 2015; Vermot et al., 2009).

The mechanisms by which flow and KLF2 drive heart morphogenesis are being uncovered in the zebrafish and in the mouse. KLF2-dependent FGF signaling from endocardial cells to their neighboring cardiomyocytes is required to prevent their extrusion and to maintain myocardial wall integrity (Rasouli et al., 2018). In murine heart valve development, shear forces acting through KLF2 activate paracrine signaling via Wingless-related integration site 9 (WNT9) from the endocardium to the underlying mesenchymal cells to drive tissue remodelling (Goddard et al., 2017). Similar morphogenetic events may also occur in the embryonic zebrafish heart, where blood flow and KLF2 are also required for the expression of Wnt9b in the endocardium.

Shear stress due to blood flow also predicts patterns of convergent extension in the developing zebrafish heart (Boselli et al., 2017). Labelling endocardial cells with photoconvertible fluorescent markers revealed that they converge towards the atrioventricular canal (AVC), but only in the presence of shear forces caused by blood flow. However, convergence of the endocardial cells towards the AVC cannot be predicted by flow patterns, which are overall unidirectional. A mechanical model of blood flow through a periodically deforming heart tube (which effectively recapitulates flow patterns) predicts that the highest wall shear stresses are experience by the AVC, suggesting that patterned shear stresses may provide the directional cue for endocardial cell convergence.

Flow also regulates development of organ systems beyond the heart and vasculature. In the developing zebrafish spinal cord, motile cilia drive bidirectional flow of CSF through the central canal that is mechanically sensed by CSF-contacting neurons via the cation channel Polycystic kidney disease 2-like 1 (PKD2L1) (Sternberg et al., 2018). Here, flow velocities are slightly faster than at the node, reaching a maximum of ~5 μm/s, and bidirectionality of the flow is attributed to the presence of motile cilia primarily on the ventral surface of the canal (Thouvenin et al., 2020). CSF within the central canal of the spinal cord is continuous with that bathing the ventricles of the developing brain; separating these compartments by photoablation leads to increased body axis curvature, suggesting that flow of CSF and/or its constituent particles from the brain to the spinal cord is important for embryonic development. Additionally, PKD2L1 is required for development of a straight spine (Sternberg et al., 2018), suggesting that CSF flow-sensing by CSF-contacting neurons broadly regulates morphogenesis of the spinal cord.

Conclusions and outlook

Developmental mechanobiology is a rapidly growing field and new, important questions are being asked. In closing, we wish to highlight two areas that might be of particular interest for future studies of the mechanics of development. Firstly, what powers all of this force generation? There is increasing interest in how regulation of metabolism is coupled to force generation and mechanosensing during development (Perestrelo et al., 2018). Metabolic reprogramming related to the activity of the mechanosensor Yap is critical for the first cell-fate specification in early mouse embryos (Chi et al., 2020) and for neural crest migration in avian embryos (Bhattacharya et al., 2020). Classic metabolomics approaches may help to explore this area, but given the inherent complexity of developing tissues, adapting newer techniques in spatial metabolomics may be especially valuable in answering this question.

Secondly, it is now well-appreciated that the nucleus itself is a mechanosensor (Elosegui-Artola et al., 2017; Kirby and Lammerding, 2018). This behavior is evident in cell culture and has been implicated in cancer, but has not yet been shown to be relevant to development. The nuclear lamina can mechanically protect the genome and this might play a role in heart development (Cho et al., 2019). Applied strain is resisted by lowering tension in the nuclear envelope and remodelling heterochromatin to prevent DNA damage in cell culture models of epithelial tissues, and there is evidence that this phenomenon may occur within epithelial folds experiencing high levels of tension during murine digit morphogenesis (Nava et al., 2020). However, mouse models of lamin mutations are able to complete embryogenesis and show only partial perinatal lethality, suggesting either that lamins play a minor mechanical role during early development or that their mechanical functions are redundant (Stewart et al., 2007). Molecular tension sensors have successfully been employed to measure forces at the cell cortex in the developing embryo (Lemke et al., 2019; Tao et al., 2019) – similar sensors that permit the measurement of forces at the interface between the cytoskeleton and the nucleus could yield important insights into this question. Additionally, improved imaging capabilities and biophysical measurements that can be applied to intact tissues will undoubtedly elucidate the role of nuclear mechanosensing in development.

Abbreviations:

AVC

atrioventricular canal

CSF

cerebrospinal fluid

ECM

extracellular matrix

EVL

epithelial enveloping layer

FGF

fibroblast growth factor

ICM

inner cell mass

KLF2

Kruppel like factor 2

PKD2L1

Polycystic kidney disease 2-like 1

PSM

presomitic mesoderm

WNT

Wingless-related integration site