\subsection{\textbf{Keratins coordinate tissue spreading by balancing spreading forces with tissue material properties.}}

Suyash Naik1, Yann-Edwin Keta2, Kornelija Pranjic-Ferscha1, Édouard Hannezo1, Silke Henkes2,* and Carl-Philipp Heisenberg1,*

1 Institute of Science and Technology (ISTA), Am Campus 1, 3400 Klosterneuburg, Austria
2 Lorentz Institute for Theoretical Physics, LION, Leiden University, P.O. Box 9504, 2300 RA Leiden, The Netherlands
* email: shenkes@lorentz.leidenuniv.nl and heisenberg@ist.ac.at

\subsubsection{Abstract}

For tissues to spread, they must be deformable while maintaining their structural integrity. How these opposing requirements are balanced within spreading tissues is not yet well understood. Here, we show that keratin intermediate filaments function in epithelial spreading by adapting tissue mechanical resilience to the stresses arising in the tissue during the spreading process. By analysing the expansion of the enveloping cell layer (EVL) over the large yolk cell in early zebrafish embryos in vivo, we found that keratin network maturation in EVL cells is promoted by stresses building up within the spreading tissue. Through genetic interference and tissue rheology experiments, complemented by a vertex model with mechanochemical feedback, we demonstrate that stress-induced keratin network maturation in the EVL increases tissue viscosity, which is essential for preventing tissue rupture. Interestingly, keratins are also required in the yolk cell for mechanosensitive actomyosin network contraction and flow, the force-generating processes pulling the EVL. These dual mechanosensitive functions of keratins enable a balance between pulling force production in the yolk cell and the mechanical resilience of the EVL against stresses generated by these pulling forces, thereby ensuring uniform and robust tissue spreading.


\subsubsection{Introduction}

Epithelial cell layer spreading is a core feature of multiple developmental and disease-related processes. In \textit{Drosophila} development, for instance, the spreading of the epidermis leads to the closure of an opening at the dorsal side of the embryo \cite{Saias2015DecreaseCellVolume,Young1993NonmuscleMyosinNeeded}. Likewise, in wound healing, the epidermal cell layer spreading and fusion close the wound \cite{Jacinto2000DynamicActinAdhesion,Losick2013PolyploidFusionWound}. Various cellular processes have been proposed to contribute to epithelial cell layer spreading, including cell spreading, cell migration, oriented cell division, and cell intercalation \cite{Brown1981CornealSpreading,Lecuit2007OrchestratingSizeShape}. These processes can generate the mechanical forces driving active tissue expansion and determine the aptitude of tissues to undergo spreading.

Zebrafish embryo morphogenesis is initiated by the spreading of the blastoderm over the large yolk cell in a process named epiboly \cite{Kimmel1995ZebrafishStages}. During epiboly, the EVL, a simple squamous epithelial cell layer formed at the surface of the blastoderm, undergoes massive spreading to eventually engulf the entire yolk cell at the end of epiboly (Figure1A) \cite{Bruce2016ZebrafishEpibolyReview,Lepage2010ZebrafishEpibolyMech}. EVL spreading has been shown to depend on the formation and contraction of a large actomyosin band positioned within a thin cytoplasmic layer at the surface of the yolk cell, the yolk syncytial layer (YSL) \cite{Behrndt2012ForcesEpithelialSpreading}. This actomyosin band within the YSL forms around the entire circumference of the yolk cell close to where the leading edge of the EVL contacts the YSL, and its contraction and flow are thought to generate the mechanical forces pulling the EVL over the yolk cell (Figure 1A) \cite{Behrndt2012ForcesEpithelialSpreading,Schwayer2019ZO1PhaseSeparation}. Both active spreading of EVL cells and oriented EVL cell divisions have been implicated in facilitating EVL spreading by modulating EVL surface tension \cite{Morita2017PhysicalBasisSpreading,Campinho2013TensionOrientedDivisions}. However, EVL morphogenesis not only relies on the generation and transmission of active forces within the EVL and YSL, but also on changes in the material properties of the tissue resisting such forces \cite{Petridou2019TissueRheologyEmbryos}. Yet, the molecular and cellular mechanisms that determine EVL material properties and how they are spatiotemporally coupled to changes in active force production remain unclear. Understanding these mechanisms requires a closer examination of the cytoskeletal components within epithelial cells, particularly those that have been implicated in modulating tissue material properties.

Keratin intermediate filaments are the most abundant and diverse cytoskeletal components in epithelial cells \cite{Laly2021KeratinRigiditySensing,Loschke2015KeratinNetworkRegulation}. They form bundled filaments from keratin type I and type II heterodimers laterally associating in an antiparallel fashion into apolar tetramers that again align and anneal longitudinally into unit-length filaments \cite{Herrmann1999EarlyIFAssemblyUnits,Herrmann2007IFsToNanomechanics}. Keratin filaments are semi-flexible, stable and highly elastic, different from the rather stiff and rigid actin and microtubule filaments \cite{Lorenz2019LateralCouplingIFMechanics,Kreplak2004IFAssemblyMechanisms}. They can self-assemble into intricate subcellular networks, the precise organisation of which depends on their functional adaptation in different cell types \cite{Bragulla2009KeratinStructureFunctions}. Generally, the intracellular keratin network can organise into a rim network, supporting plasma membrane integrity and connecting desmosomal contacts, and a spoke network, surrounding the nucleus and transferring information from the cell exterior to the nucleus \cite{Quinlan2017RimSpokeIFHypothesis,Acehan2008PlakoglobinIFAnchorage,Kroger2013KeratinsAdhesionPKC}. In polarised simple epithelia, such as the EVL, keratin networks are typically positioned near the apex where they are thought to function in resisting mechanical and chemical stresses and maintaining epithelial apicobasal polarity \cite{Windoffer2021QuantitativeKeratin3D,Salas2016KeratinBarrierPolarity,Kayser2012AssemblyKineticsKeratin}. Keratins have been shown to be sensitive to mechanical forces by reorganizing and changing their mechanical properties upon stress application \cite{Lorenz2019LateralCouplingIFMechanics,Latorre2018ActiveSuperelasticity,Duque2024RuptureStrengthMonolayers}. Yet, how keratin network mechanosensitivity functions in epithelial tissue morphogenesis remains unsettled \cite{Lim2020KeratinsAsymmetricInheritance,Nahaboo2022KeratinMembranesMouse}.

The role of epithelial keratins in development and disease is only beginning to be understood \cite{Fuchs1995KeratinsSkin,Homberg2014IRCMBBeyondExpectations}. Keratin mutations can cause diseases that lower the resilience of epidermal tissues to mechanical stress \cite{Pan2013ExpandingSignificanceKeratin}. Moreover, studies in mouse embryos have shown that keratins are required for trophectoderm specification and extra-embryonic tissue growth and expansion \cite{Lim2020KeratinsAsymmetricInheritance,Nahaboo2022KeratinMembranesMouse}. Here we show that keratin intermediate filaments are required for EVL spreading during zebrafish epiboly. They function in this process by balancing EVL tissue viscosity with the external pulling forces mediating EVL spreading. This balancing function of keratins enables the EVL to undergo uniform spreading without rupturing.

\subsubsection{Results}

\textbf{Keratins are specifically expressed within the EVL and YSL during gastrulation.}
\newline
To explore the function of keratins during early zebrafish embryogenesis, we analysed the expression of different keratins in embryos from early blastula to late gastrula stages (4.5--8.5~hpf). Previous studies have shown that 13 type~I keratins and 3 type~II keratins are specifically expressed within the developing EVL during this period \cite{Ho2022KeratinGeneUpdate,Farrell2018SingleCellTrajectoriesZfish}. To identify the temporal keratin expression profiles within the EVL during epiboly, we used RT\mbox{-}qPCR to map the expression of three keratin type~II (keratin~4,~5,~8) and one keratin type~I (keratin~18), abundantly expressed within the EVL \cite{Farrell2018SingleCellTrajectoriesZfish,Pei2007MaternalFoxH1Zfish}. We found all of these keratins to be expressed already at early blastula stages, with their expression continuously increasing until the end of gastrulation (Figure~Supplement~1A). The expression was restricted to the epithelial cells as evidenced by fluorescence \textit{in situ} hybridization for keratin~8 mRNA (Figure~Supplement~1B). To determine the spatial distribution of keratins within the epibolizing embryo, we took advantage of \textit{Tg}(krt18\!:\!krt18\text{-}GFP) embryos expressing GFP\mbox{-}tagged keratin~18 under its endogenous promoter. Keratin~18 expression was first detected in EVL progenitor cells within the early gastrula (4.0~hpf) arranged in short, bundled and unconnected filaments located predominantly at the apical surface of these cells (Figure~1B, Video~1 and Figure~Supplement~1C). Keratin~18 continued to be selectively expressed within EVL cells until mid\mbox{-}gastrulation (5.5~hpf), when some additional expression was also detected within the forming YSL directly adjacent to the place where the leading edge of the EVL contacts the YSL (Figure~1B and Figure~Supplement~7A). During this period and continuing until the end of gastrulation, the apical network of keratin~18 filaments within EVL cells became increasingly dense and interconnected (Figure~1B), as evidenced by a continuous increase in keratin~18\mbox{-}GFP intensity and network density between 4.5 and 8.5~hpf (Figure~1C and~D, and Video~1). This increase in keratin network intensity and density was accompanied by EVL cells increasing the apical area during epiboly (Figure~1E), suggesting a close temporal correlation between keratin network maturation and EVL cell spreading (Figure~1F). Initially, keratin filaments were predominantly localized to the apical surface of the EVL cells (Figure~1B and Supplement~1C), but from 6~hpf onwards, additional keratin~18 expression was detected in a bundle\mbox{-}like configuration along the apical junction of EVL cells (Figure~1B and~G), consistent with previous observation of keratins showing both junctional (``rim'') and apical (``spoke'') localization \cite{Quinlan2017RimSpokeIFHypothesis,Lim2020KeratinsAsymmetricInheritance,Nahaboo2022KeratinMembranesMouse}.

\textbf{Keratin network maturation is mechanosensitive.}
\newline
EVL epiboly movements are driven by a large actomyosin cable forming within the YSL and pulling the margin of the EVL over the yolk cell \cite{Behrndt2012ForcesEpithelialSpreading}. Given that keratin networks can reorganise under stress \cite{Duque2024RuptureStrengthMonolayers,Sivaramakrishnan2009ShearStressKeratinPKCz}, we speculated that the observed maturation of the keratin network within EVL cells might be facilitated by EVL tissue tension building up during the course of epiboly \cite{Behrndt2012ForcesEpithelialSpreading,HernandezVega2017PolarizedCorticalTension}. To test this possibility, we analysed whether and how EVL network maturation is affected in embryos where EVL tension is either increased or decreased. To modulate EVL tension, we expressed a constitutively active form of RhoA (\textit{CA}\,RhoA) specifically within the YSL, promoting YSL actomyosin contractility and pulling, or a constitutively active form of myosin phosphatase (\textit{CA}\,Mypt) leading to reduced YSL contractility and pulling \cite{Schwayer2019ZO1PhaseSeparation,Jayashankar2013PP1bZfishMyosinPhosphatase}. In \textit{CA}\,RhoA\mbox{-}expressing embryos, keratin~18 expression intensity and network density in EVL cells prematurely increased during the course of epiboly, while in \textit{CA}\,Mypt\mbox{-}expressing embryos, keratin expression and network maturation were delayed (Figure~2A--D, Video~2--5 and Figure~Supplement~2B). This keratin mechanosensitivity was detectable both when network maturation was analysed as a function of developmental time or degree of epiboly progression (Figure~2A--D and Figure~Supplement~2B--C), suggesting that the effect of EVL tension on keratin network maturation is not just a secondary consequence of changes in EVL epiboly movements (Figure~Supplement~2D).

To more directly assess the effect of EVL tension on keratin network maturation, we locally increased EVL tension in 5 and 7~hpf embryos by aspirating the EVL using a micropipette and analysing resultant changes in EVL network maturation. We found that keratin filaments showed increased accumulation in EVL cells upon aspiration (Figure~2F--H, Video~8), further supporting the notion that keratin network maturation within the EVL cells is promoted by EVL tension building up during the course of epiboly.

To investigate the mechanisms underlying keratin network mechanosensitivity, we examined whether keratin mRNA expression within the EVL is responsive to reduced pulling forces from the YSL. Using qPCR, we compared keratin~4,~5,~8 and~18 expression levels in control embryos and embryos expressing \textit{CA}\,Mypt within the YSL, which attenuates actomyosin contraction\mbox{-}mediated pulling on the EVL. Notably, this comparison revealed no significant differences in keratin~4,~5,~8 and~18 expression (Figure~Supplement~2I), arguing against a force\mbox{-}dependent transcriptional regulation of keratin within the EVL.

Interestingly, in our micropipette aspiration assay, increased keratin expression in the EVL was spatiotemporally accompanied by a marked enhancement of actin network assembly (Figure~Supplement~2F). Given the well\mbox{-}established mechanosensitivity of the actomyosin cytoskeleton \cite{Chanet2017ActomyosinMechanosensingShape,He2016FlowDependentMyosinRecruitment} and its known physical interactions with the keratin network \cite{Bragulla2009KeratinStructureFunctions,Werner2004EBSMutationsKeratinDynamics,Kolsch2009ActinDependentKeratinPrecursors}, this observation raises the possibility that keratin mechanosensitivity in the EVL may depend on actin network dynamics.

To test this hypothesis, we treated embryos with low concentrations of Cytochalasin~D (CytoD) to impair actin network assembly in EVL cells and assessed keratin upregulation upon micropipette aspiration. Strikingly, CytoD\mbox{-}treated embryos showed a markedly reduced keratin response to aspiration (Figure~2H), suggesting that proper actin network assembly is required for keratin mechanosensitivity in the EVL.

To more directly assess whether actomyosin contractility affects keratin network assembly, we injected \textit{CA}\,Mypt mRNA into a single blastomere at the 64\mbox{-}cell stage to generate EVL clones with reduced contractility. We found that downregulation of actomyosin contractility resulted in a corresponding decrease in keratin network assembly within these clones (Figure~2A--D and Figure~Supplement~2G--H), supporting a causal role of actomyosin tension in promoting keratin assembly.

Together, these findings demonstrate that the keratin cytoskeleton in EVL cells is mechanosensitive and that this mechanosensitivity is dependent on the mechanosensitive assembly of the actin network.

\textbf{Keratins regulate EVL tissue properties.}
\newline
Keratins have previously been implicated in resisting mechanical stresses in epithelial cells. Thus, the mechanosensitive coupling between EVL keratin network maturation and pulling force generation within the YSL might constitute a mechanism to protect the EVL tissue against excessive deformations by mechanically strengthening it proportionally to the force exerted on it.

To test this possibility, we knocked down/out the expression of the two keratin type~II genes (keratin~4 and~8) primarily expressed within EVL cells \cite{Farrell2018SingleCellTrajectoriesZfish}, reasoning that in the absence of keratin type~II expression, no keratin dimers can be formed within the EVL and thus keratin network formation should be defective. Ubiquitous knock\mbox{-}down/out of keratin~4 and~8 expression by using morpholinos and CRISPR\mbox{-}Cas9 or interference with keratin network formation by overexpressing a dominant\mbox{-}negative version of keratin~18 (DNkrt18) \cite{Colombelli2013LaserNanosurgeryReview}, led to strongly diminished keratin~18 expression and network formation within EVL cells (Figure~3A--C, Video~6--7 and Figure~Supplement~3A--B). Loss of the keratin network in all these perturbations led to delayed EVL epiboly movements (Figure~3D, Video~6 and~7 and Figure~Supplement~3C) and frequent rupturing of the EVL towards the end of gastrulation, causing embryo lethality (Figure~Supplement~3D--E). Rupture of the EVL was typically preceded by a reduction in junctional localization of E\mbox{-}cadherin (adherens junctions), Occludin\mbox{-}B (tight junctions), and plakoglobin\mbox{-}A (desmosomal junctions) (Figure~Supplement~3F--H), suggesting that keratins are essential for maintaining EVL tissue integrity by enabling proper formation and stabilization of cell\mbox{-}cell junctions between EVL cells.

To understand how keratins function in EVL epiboly movements, we analysed changes in EVL cell shapes and rearrangement in response to the pulling forces from the YSL during epiboly. Comparing wild\mbox{-}type with keratin~4/8 morphant embryos revealed that EVL cells in wild\mbox{-}type embryos coordinately elongated along the animal\mbox{--}vegetal (AV) axis of the gastrula, the axis of EVL spreading, while no such coordinated cell elongation was detectable in morphant embryos (Figure~4A--B). This points at the possibility that tissue material properties, determining force transduction and mechanical resilience, might be altered by keratin network formation.

To investigate this possibility, we measured the material properties of EVL tissue at early and mid\mbox{-}gastrulation (5 and 7~hpf) using micropipette aspiration. Analysis of the flow profile of the EVL into the pipette revealed a linear response, consistent with viscous properties, which allowed us to assess the viscosity of EVL tissue. Comparing the viscosity of EVL tissue between early and mid\mbox{-}gastrulation stages showed a significant increase in viscosity at mid\mbox{-}gastrulation, coinciding with the maturation of the keratin network (Figure~4C). This suggests that keratin network maturation during epiboly contributes to the increase in EVL tissue viscosity. To test this function of keratins more directly, we performed EVL micropipette aspiration experiments in wild\mbox{-}type and keratin~4/8 morphant embryos. At early gastrulation, when differences in keratin network formation between wild\mbox{-}type and morphant embryos were still relatively small (Figure~1B), we observed a slight reduction in viscosity in keratin~4/8 morphant embryos compared to wild\mbox{-}type (Figure~4C). In contrast, at mid\mbox{-}gastrulation (Figure~4C, Video~8), when the differences in keratin network organization between morphant and wild\mbox{-}type embryos became more pronounced (Figure~1B), viscosity was significantly reduced in the morphant embryos. This supports the idea that keratin network maturation enhances EVL tissue viscosity.

To test whether this effect is due to keratin expression specifically within EVL cells, we knocked down keratin~4/8 expression within the YSL by injection of keratin~4/8 morpholinos directly into the forming YSL at the 512\mbox{-}cell stage. Interestingly, this did not cause any detectable changes in coordinated EVL cell elongation along the AV axis, suggesting that keratins control EVL cell shape changes in a tissue\mbox{-}autonomous manner (Figure~Supplement~4A--B). To further challenge this conclusion, we developed an EVL wound\mbox{-}healing assay, allowing us to monitor autonomous EVL spreading during wound closure \cite{Bi2016MotilityDrivenGlass,Farhadifar2007EpithelialPacking}. For wounding the EVL, we ablated 3--5 neighbouring cells at random positions within the EVL in mid\mbox{-}gastrulation stage embryos (7~hpf) and observed how the coordinated spreading of the neighbouring cells extruded the ablated cells. In control embryos upon ablation, a supracellular actin cable formed at the leading edge of EVL cells neighbouring the ablated cells (Figure~4D, Video~9--10). This was accompanied by the highly coordinated movement and spreading of EVL cells towards the site of cell ablation, eventually leading to the extrusion of the ablated cells and closure of the wounding site (Figure~4G). During this process, not only the leading edge EVL cells elongated towards the wounding site, but also cells further away from the ablation site, resulting in a highly coordinated long\mbox{-}range tissue spreading detected by tissue flow and cell alignment analysis (Figure~4D--G). In contrast, EVL oriented cell elongation and tissue spreading were largely restricted to EVL cells directly neighbouring the wounding site in keratin~4/8 morphant and crispant embryos (Figure~4D--G and Figure~Supplement~4C--E). This supports the notion that keratin expression within the EVL promotes tissue viscosity, leading to an increased length (hydrodynamic length) by which the tissue deforms in response to forces pulling at its margin.

\textbf{Keratins are required for mechanosensitive actomyosin contraction within the YSL.}
\newline
Given that keratin expression within the EVL is mechanosensitive, we hypothesised that this behaviour might constitute a feedback mechanism balancing EVL tissue viscosity with the forces pulling on its margin, thereby setting the rate of EVL epiboly movements. To address this hypothesis, we developed a vertex model \cite{YketaHenkesCellsGitHub,Wang2018Keratin6CollectiveMigration} of the EVL, enabling us to relate EVL tissue spreading dynamics to mechanosensitive keratin expression. To this end, we wrote the energy, which penalises deviations of the areas $A_i$ and perimeters $P_i$ of the cells $i$ from their target values $A_{i,0}$ and $P_{i,0}$, respectively, as
\[
U=\sum_i \left[ \frac{\Gamma_i}{2A_0}\,\bigl(A_i-A_{i,0}\bigr)^2 \;+\; \frac{\Gamma_i}{2}\,\bigl(P_i-P_{i,0}\bigr)^2 \right],
\]
where $\Gamma_i$ is the cell stiffness constant. In order to make the tissue viscoelastic, we relaxed the target area $A_{i,0}$ over a characteristic viscous relaxation timescale $\tau_i$ following
\[
\tau_i\,\frac{dA_{i,0}}{dt}=-(A_{i,0}-A_i),
\]
while enforcing a minimum area $A_0$, and scaled $P_{i,0}$ to work at a constant shape parameter $s_0 = P_{i,0}/\sqrt{A_{i,0}} = 3.72$, a value where the tissue is predicted to be still rigid \cite{YketaHenkesCellsGitHub}.

We next incorporated keratin into our model (Figure~5D). As illustrated in Figure~1B, keratin forms an intracellular network that is initially floppy, but reaches a percolation threshold at approximately $K_{\mathrm{th}}=150$. Capturing the full complexity of its behavior would require simulating the coupled dynamics of actin and keratin networks, including keratin polymerization and depolymerization as well as actomyosin contractility. To simplify the analysis, we instead implemented a mesoscale approximation in which the keratin network acts in parallel with other mechanical components of the cell. Above the percolation threshold, keratin contributes proportionally to both the cell’s effective stiffness and relaxation time with
\[
\Gamma_i=\Gamma\!\left(1+\beta\,\Delta K_i\right),\qquad \tau_i=\tau\!\left(1+\beta\,\Delta K_i\right),
\]
where $\Delta K_i=\max\!\left(0,\,K_i-K_{\mathrm{th}}\right)$ and $\beta$ is the mechanical feedback constant that determines the mechanical effect of the keratin concentration above the threshold $\Delta K_i$.

To include keratin mechanosensitivity, we further assumed that its assembly from the cytosol increases with the stress applied on the tissue. To account for this, we used a simple, linear model of the evolution of the keratin concentration,
\[
\tau_K\,\frac{dK_i}{dt}=\alpha\,\max\!\bigl(0,\,p_i\bigr)-K_i,
\]
where keratin dissociates with timescale $\tau_K$ and $\alpha$ is a coupling constant (note that other nonlinear rheology models have also been used to model the effect of keratin \cite{Duque2024RuptureStrengthMonolayers}). Mechanosensitivity in our model arises through biaxial stress, or equivalently, the in\mbox{-}plane pressure $p_i$ defined as
\[
p_i=\frac{1}{V_0}\!\left(\frac{\Gamma_i}{A_0}\,(A_i-A_{i,0})\,A_i \;+\; \Gamma_i\bigl(P_i-s_0\sqrt{A_{i,0}}\bigr)P_i\right),
\]
where $V_0$ is the conserved 3D cell volume (see SI, Section~2). This formulation captures the pressure response arising from deviations in both cell area and perimeter relative to their preferred values. In this framework, EVL spreading corresponds to a substantial increase in cell area $A_i$, accompanied by dramatic thinning of individual cells. At steady state, this mechanosensitive feedback yields a linear relationship between keratin concentration and pressure, expressed as $K_i=\alpha P_i$ under the assumption of positive pressure. A summary of this mechanochemical feedback loop is presented in Figure~5E.

To estimate the parameters for simulating EVL spreading, we used the EVL aspiration experiments described above (Figure~5A--B, Figure~2F--H and~4C). First, we separately fitted the low\mbox{-}keratin initial response and the high\mbox{-}keratin release curve in the aspiration experiments using a modified viscoelastic Maxwell model to obtain estimates of dissociation timescale $\tau_K$, pressure coupling $\alpha$, cell stiffness constant $\Gamma$, and relaxation timescale $\tau$ (Figure~5C, Figure~Supplement~5A--D and SI Section~1). Then we used these values together with a mean\mbox{-}field version of the mechanical model (see SI Section~4) to compute tissue height and keratin expression in the aspirated tissue (Figure~5F--G and Figure~Supplement~6A--D). We chose $\beta=0.005$ for the wild\mbox{-}type model tissue such that the increase of keratin levels from $K_i=0$ to $K_i=550$ with a percolation threshold $K_{\mathrm{th}}=150$ leads to a 2\mbox{-}fold increase of $\Gamma$ and $\tau$ (Figure~Supplement~5E--F). Please see Table~S1 for the full list of fitted parameters.

Using these parameters in our extended vertex model (see simulation library \cite{YketaHenkesCellsGitHub}), we first simulated the EVL wound\mbox{-}closure behaviour in wild\mbox{-}type and keratin\mbox{-}deficient embryos. For this, we initialised a disordered tissue patch under tension, and then created a model wound by deleting several cells and adding a contractile cable to the wound perimeter (Figure~5H--J, Video~14; for details see SI Section~3). We observed that junction tension
\[
t_i=\Gamma_i\,(P_i-P_{i,0})
\]
was higher and more widely distributed around the wound in wild\mbox{-}type ($\beta=0.005$) compared to keratin\mbox{-}deficient ($\beta=0$) tissues (Figure~5I), while the time required to close the wound remained largely unchanged between these conditions (Figure~5J). This closely matched our experimental observations where keratin\mbox{-}deficient embryos exhibited a more localized distribution of junctional tension near the wound site (Figure~4D--G), while the wound\mbox{-}closure rates were indistinguishable between wild\mbox{-}type and keratin\mbox{-}deficient embryos (Figure~4H). These results support the hypothesis that keratin contributes to a mechanochemical feedback loop within the EVL, modulating tension propagation without significantly altering the kinetics of wound closure.

To test whether we can also simulate the behaviour observed for EVL spreading during epiboly, we simulated epiboly by representing the EVL tissue as a disordered circular packing of $N=529$ cells, where the outer vertices are pulled outwards (Figure~5K and Video~15). As the actual embryo is spherical, we defined the effective height of the model tissue as
\[
z=\frac{\sum_i A_i}{2\pi R},
\]
which is the height of its projection on a sphere of radius $R=350~\mu\mathrm{m}$, and we defined the tissue’s velocity $v_z$ as the time derivative of this height. To mimic the pulling of the EVL margin by actomyosin contraction and flow within the YSL, we applied an outward force $F_{\mathrm{YSL}}$ on the edge of the simulated tissue with an amplitude which increases linearly with time, following the experimentally determined pulling\mbox{-}force evolution within the YSL \cite{Behrndt2012ForcesEpithelialSpreading}. We then measured the mean keratin expression $K$ and edge velocity $v_z$ as a function of time and pulling forces (Figure~5N--P). By ramping the pulling force from $0$ at the beginning to $0.57~\mu\mathrm{N}$ (a value equivalent to the pulling force used in the aspiration experiments) at the end of the simulation, we obtained keratin expression values and tissue edge velocities matching our experimental observations (Figure~1C, 3D and~5O). Interestingly, we also found, similar to our experimental observations (Figure~5M), that keratin expression increased to heterogeneous levels in individual EVL cells in our simulations (Figure~5L), suggesting that this heterogeneous keratin expression is mechanically regulated.

To determine whether our model can also account for the experimentally observed changes in EVL dynamics when pulling forces within the YSL were either increased or decreased, we analysed the response of keratin expression and tissue edge velocity to variations in pulling forces ($F_{\mathrm{YSL}}$) in our simulations (Figure~5N,~P). Consistent with experimental observations, mean keratin expression increased when $F_{\mathrm{YSL}}$ was upregulated and decreased when $F_{\mathrm{YSL}}$ was downregulated (Figure~5N). Moreover, in line with the observed scaling of keratin expression, and thus tissue viscosity, edge velocity in our simulations was not significantly affected by changes in $F_{\mathrm{YSL}}$ (Figure~5P). The origin of this rheostat\mbox{-}like behaviour is that the fixed point of the nonlinear keratin--strain\mbox{-}rate dynamics is nearly independent of the strain rate (see SI Section~4). However, these predictions did not align with our experimental findings at later stages of gastrulation, where both increasing and decreasing $F_{\mathrm{YSL}}$---by modulating actomyosin contractility within the YSL---slowed EVL epiboly movements (Figure~Supplement~2D). This suggests that changes in keratin expression, and consequently EVL tissue viscosity, cannot fully compensate for alterations in $F_{\mathrm{YSL}}$ throughout gastrulation. A similar discrepancy between model predictions and experimental results emerged when comparing tissue edge velocity between wild\mbox{-}type and keratin\mbox{-}deficient embryos. While our simulations predicted a significant increase in edge velocity in keratin\mbox{-}deficient embryos compared to wild\mbox{-}type (Figure~5O)---as expected for a less viscous and more deformable EVL---experimental data showed a decrease in edge velocity in keratin\mbox{-}deficient embryos (Figure~3D).

Interestingly, in our simulations, the reduction in EVL tissue edge velocity observed in keratin\mbox{-}deficient embryos could only be explained by a decrease in the pulling force $F_{\mathrm{YSL}}$ (Figure~5P). This led us to speculate that keratins may have additional functions within the YSL in regulating $F_{\mathrm{YSL}}$, a possibility supported by previous findings that have implicated keratins in actomyosin network organization and mechanosensation \cite{Karsch2020KeratinNetworkIntegrity,Lulevich2010SingleCellMechanicsKeratinocytes}. To test this possibility, we analysed dynamic actomyosin reorganization within the YSL in the presence and absence of keratins. In wild\mbox{-}type embryos, retrograde flows of actin and myosin within the YSL led to the formation of a contractile actomyosin band, previously shown to generate the pulling forces that drive EVL spreading \cite{Behrndt2012ForcesEpithelialSpreading}. However, in embryos where keratin~4/8 was specifically knocked down within the YSL, these actomyosin flows were severely diminished and less aligned (Figure~6A, D, F).

To investigate the functional interaction between keratins and the actomyosin network within the YSL, we analyzed the coordination of YSL actin and keratin network dynamics during EVL epiboly movements. Using particle image velocimetry (PIV) analysis, we observed both actin and keratin retrograde flows within the YSL, with keratin flows being more spatially confined to the EVL--YSL boundary where the keratin network was predominantly localized (Figure~Supplement~7C--C$'$). Moreover, actin flows further away from the EVL--YSL boundary were strongly reduced when reaching the region of keratin localization close to the boundary, suggesting that actomyosin contraction\mbox{-}driven flows compress the keratin network by physically interacting with it as previously reported \cite{Wong2003K6WoundRepair}.

To assess whether this interaction impacts force transmission within the actin network---potentially by keratins providing a rigid scaffold that enhances actin network crosslinking---we performed UV\mbox{-}laser ablation of the YSL actin network in embryos with or without keratin expression in the YSL. Strikingly, the actin recoil velocity and hydrodynamic length following laser cuts was significantly reduced in \textit{krt4/8} morphants compared to controls (Figure~Supplement~7D--F), indicating decreased force transmission within the actin network in the absence of keratin.

Given this keratin\mbox{-}dependent effect on actin network rheology, and previous reports linking keratins to actomyosin mechanosensation \cite{Karsch2020KeratinNetworkIntegrity,Lulevich2010SingleCellMechanicsKeratinocytes}, we next asked whether keratin within the YSL might be required for the YSL actin network responding to tension generated at the EVL--YSL boundary. To address this, we performed micropipette aspiration of the YSL and monitored mechanosensitive changes in the actomyosin network in both wild\mbox{-}type and keratin loss\mbox{-}of\mbox{-}function embryos. In wild\mbox{-}type embryos, pulling on the YSL induced a local increase in actin intensity near the site of aspiration (Figure~6B,~E), suggesting that YSL actomyosin contraction and flow is upregulated by tension generated at the EVL--YSL boundary. In contrast, embryos with specific knockdown of keratin~4/8 in the YSL failed to show this upregulation (Figure~6B,~E), suggesting that this mechanosensitive response of the actomyosin network depends on keratin network formation within the YSL.

Collectively, these findings suggest that keratins are required for the generation of tension within the YSL that drives EVL epiboly movements, likely by supporting actin network crosslinking and its mechanosensitive response to applied forces.

\subsubsection{Discussion}

Our findings suggest that keratins function in EVL spreading during epiboly by coupling tissue contractility and tension to tissue viscosity and connectivity. During epiboly, actomyosin contraction and flow within the YSL generate the forces pulling the margin of the EVL towards the vegetal pole of the gastrula. These pulling forces not only induce stress within the EVL, but also within the YSL to which the EVL is mechanically coupled at its margin. In the EVL, build-up of stress triggers keratin network maturation, which again increases EVL tissue viscosity and promotes cell-cell junction formation and maintenance, thereby ensuring that the tissue remains intact when stress rises during epiboly. In the YSL, keratins are essential for transmitting forces within the actin network, promoting stress-dependent actin accumulation, and enabling efficient contractions required to pull the EVL over the yolk cell. By fulfilling this dual mechanosensitive role — linking EVL viscosity to YSL contractility — keratins ensure robust and efficient EVL spreading during epiboly.

Keratins have previously been shown to be stress-responsive and display important functions in mechanical tissue resilience and spreading \cite{Nahaboo2022KeratinMembranesMouse, Herrmann2007IFsToNanomechanics, Karsch2020KeratinNetworkIntegrity, Lulevich2010SingleCellMechanicsKeratinocytes}. While spreading necessitates malleability, mechanical resilience is facilitated through properties such as tensile strength. Various studies have linked keratins to both these functions in culture cells and mouse embryos \cite{Nahaboo2022KeratinMembranesMouse, Wong2003K6WoundRepair}; however, it remains unclear exactly how keratins could mediate these contrasting roles in a systemic manner. Our findings suggest a mechanism by which to reconcile the functions of keratins in ensuring tissue integrity \cite{Homberg2014IRCMBBeyondExpectations, Karsch2020KeratinNetworkIntegrity, Stenvall2021K8DeletionColon} and promoting tissue spreading \cite{Nahaboo2022KeratinMembranesMouse, Deek2016MechanicsSoftKeratinNetworks}. The ability of keratins to resist mechanical stress increases as their organisation progressively transforms from an immature, disconnected form into a dense, interconnected cellular network, which ultimately develops into a supracellular, tissue-scale network through the association of keratins with desmosomal junctions \cite{Quinlan2017RimSpokeIFHypothesis}. This process links cellular keratin networks and stabilises desmosomal proteins \cite{Acehan2008PlakoglobinIFAnchorage, Buchau2022DesmosomeHyperadhesionKeratin, Kowalczyk1998DesmosomesReview}. To ensure that this increasing resistance against deformation of the EVL does not stall EVL spreading and epiboly movements, keratins also facilitate tension-dependent actin network accumulation within the YSL, thereby adapting mechanical pulling force production within the YSL to EVL tissue viscosity resisting its deformation.

Previous studies have suggested that keratins can interact with the actomyosin cytoskeleton \cite{Herrmann2007IFsToNanomechanics, Kim2012ModelingActinKeratinSpreading}, although the biochemical basis and functional significance of this interaction during early development remain poorly understood. Keratin–actin interactions are thought to occur either directly \cite{Werner2004EBSMutationsKeratinDynamics, Herrmann2007IFsToNanomechanics} or indirectly via large cytolinker proteins such as plakins and plectin \cite{Bouameur2014PlakinsReview, Inaba2020KeratinsPlakinsMicroridges}. Disruption of the actin cytoskeleton has been shown to impair keratin network organization and stability \cite{Kolsch2009ActinDependentKeratinPrecursors}. Conversely, keratin loss can perturb actin network architecture during wound healing \cite{Wong2003K6WoundRepair}, as well as impair actin stress fiber formation and cell polarization in response to local mechanical forces acting on C-cadherins \cite{Weber2012CadherinKeratinMechano}. Our results provide direct evidence that keratin and actin networks interact functionally: keratins are required for actin mechanosensation within the YSL, and conversely, actin is necessary for keratin mechanosensation within the EVL. Whether these interactions are purely mechanical — for example, providing structural support for network assembly and remodeling — or also involve biochemical signaling pathways that regulate polymerization and architecture remains to be determined.

Notably, keratin intermediate filaments are thought to be absent in insects \cite{Herrmann2011IFHistoryInsects}, suggesting that tissue morphogenesis and spreading in these animals can occur in the absence of keratin function. In embryogenesis of the insect \textit{Tribolium castaneum}, for instance, the extraembryonic serosa, a simple squamous epithelial cell layer, undergoes massive spreading during epiboly \cite{Jain2020FluidizationGapClosureInsect}. Similar to EVL epiboly movements in zebrafish, serosa spreading is mediated by forces pulling on its leading edge \cite{Jain2020FluidizationGapClosureInsect}. Interestingly, this pulling leads to a regionalization of the serosa tissue into a solid-like dorsal portion with little cell rearrangements and a fluid-like ventral portion consisting of cells undergoing intercalations \cite{Jain2020FluidizationGapClosureInsect}. In contrast, no such clear regionalization can be observed in the zebrafish EVL with very little cell intercalations occurring throughout the tissue except some cells at the EVL margin withdrawing from the leading edge at very late stages of EVL epiboly. This different response of the EVL and serosa tissues to pulling forces might be due to the presence and absence of keratin expression within the respective tissues, pointing to the intriguing possibility that the function of keratins for homogeneous tissue spreading has become dispensable in insects. How this function of keratins in epithelial tissues has been adapted to specific organismal settings, and why keratins became expendable in many insect species remains to be explored.

While keratins belong to the most abundant and diverse cytoskeletal components in epithelial cells, remarkably little is yet known about the mechanisms by which they function in epithelial tissues. Our findings identify a critical role of keratins in promoting tissue viscosity and contractility in response to tissue tension. This ensures robust tissue spreading by balancing tissue integrity and expansion.
\newpage
\subsubsection{Figures}
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/Fig1_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Keratin network maturation in the EVL.} }

    \label{Figure1Chapter3}
\end{figure}

\smallskip
\noindent\textbf{Figure~\ref{Figure1Chapter3} Keratin network maturation in the EVL.} 

    (A) Left: schematic representation of a zebrafish embryo at 60\% epiboly stage (6 hpf) showing the expression and localization of keratin within the EVL and YSL (together with actin). Arrows mark the direction of EVL epiboly movements. Right: cross-section of the region outlined in the left panel marking the expression and localization of keratin, actin (within the YSL) and junctional components linking the margin of the EVL to the YSL.
    
    (B) Maximum intensity projection images of keratin (top and middle rows) and actin (Utrophin; bottom row)) expression in Tg(krt18:krt18-GFP) (keratin) and Tg(acbt2:utr-mCherry) (actin) embryos showing the progression of keratin expression within the EVL and YSL during epiboly (4 - 9.5 hpf).
    
    (C) Averaged keratin intensity within the EVL in Tg(krt18:krt18-GFP) embryos as a function of time during epiboly (4 - 9.5 hpf). N = 3 experiments, n = 5 embryos. Error bars as ribbons SD of mean.
    
    (D) Averaged density of the keratin network in Tg(acbt2:utr-mCherry, krt18:krt18-GFP) embryos as a function of time during epiboly (4 - 9.5 hpf). N = 3, n = 6 embryos. Error bars as ribbons SD of mean.
    
    (E) Average apical cell area of individual EVL cells in Tg(acbt2:utr-mCherry, krt18:krt18-GFP) embryos as a function of time during epiboly (4 hpf-8.5 hpf). N = 3, n = 4 embryos. Error bars as ribbons SD of mean.
    
    (F) 3-dimensional (3D) plot of keratin intensity, network density, and EVL cell area as a function of time during epiboly. N = 3, n = 4 embryos. Spread of the surface (width) represents data spread, indicating their variability (SD of intensity and area).
    
    (G) Correlation of junctional and apical keratin intensity measured in individual EVL cells in Tg(acbt2:utr-mCherry, krt18:krt18-GFP) embryos at 60\% epiboly stage (6 hpf) N = 3, n = 3 embryos.
\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/Fig2_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Pulling forces promote keratin expression within the EVL.} }

    \label{Figure2Chapter3}
\end{figure}

\smallskip
\noindent\textbf{Figure ~\ref{Figure2Chapter3} Pulling forces promote keratin expression within the EVL}

(A) Schematic showing actomyosin contraction and flows within the YSL providing the mechanical forces pulling the EVL over the yolk cell during epiboly (for details see10) in a zebrafish embryo at 4 hpf (left) and 6 hpf (right). Green arrows, EVL epiboly movements; black arrows, actomyosin contraction within the YSL.
    
(B) Maximum intensity projection images of keratin expression in representative  Tg(acbt2:utr-mCherry, krt18:krt18-GFP) control embryos (YSL injection of 0.2\% phenol red, top row) and embryos injected with 100 pg CAMypt (middle row) and 50 pg CARhoA (bottom row) into either the YSL at 3.3 hpf (CAMypt) or marginal cells at 3.3 hpf (CARhoA) at sphere stage (4 hpf, left column), shield stage (6 hpf, right column), and  90\% epiboly stage (9.5hpf, right column). Scale bar: 100 µm.
    
(C) Plot of average keratin intensity as a function of time (hpf) in Tg(krt18:krt18-GFP) control (green, N = 3, n = 5 embryos), CAMypt (orange, N = 3, n = 5 embryos) and CARhoA mRNA injected embryos (pink, N = 3, n = 4 embryos) as described in (B). Error bars as ribbons SD of mean.
    
(D) Plot of average density of keratin network as a function of time (hpf) in Tg(actb2:utr-mCherry, krt18:krt18-GFP) control (green, N = 3, n = 6 embryos), CAMypt (orange, N = 3, n = 4 embryos) and CARhoA mRNA injected embryos (pink, N = 3, n = 3 embryos) as described in (B). Error bars as ribbon SD of mean of individual cells per replicate.

(E) 3-dimensional (3D) plot of keratin intensity, network density and EVL cell area as a function of time (hpf) in Tg(actb2:utr-mCherry, krt18:krt18-GFP) control (viridis, N = 3, n = 3 embryos), caMypt (orange, N = 3, n = 3 embryos) and caRhoA mRNA injected embryos (White to Pink, N = 3, n = 3 embryos) as described in (B). Spread of the surface (width) represents data spread, indicating their variability (SD of intensity and area).

(F) Schematic showing pipette aspirations of the EVL of a 70\% epiboly (7 hpf) stage embryo where the regions of interest within the pipette and outside of it are marked as yellow boxes.

(G) Maximum intensity projection images of keratin localization and intensity within the EVL before (left) and after (right) aspiration with a pipette in a representative Tg(krt18:krt18-GFP) embryo. White lines outline the boundary of the pipette. Scale bar: 10 µm.

(H) Box plot of keratin intensity within the pipette before (green) and after (orange) EVL aspiration in control (N = 4, n = 33 embryos), and Cytochalasin D (CytoD; 25 nM)-treated Tg(actb2:utr-mCherry, krt18:krt18-GFP) embryos at 5 and 7 hpf (N = 3, n = 25 embryos). Boxes represent quartiles for the data, dots outliers (pvalues: ***<0.001 Wilocoxon test).
\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/Fig3_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Loss of keratin expression diminishes EVL epiboly movements.} }

    \label{Figure3Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{Figure3Chapter3} Loss of keratin expression diminishes EVL epiboly movements}

    (A) Maximum intensity projection images of keratin expression in representative Tg(krt18:  krt18-GFP) embryos at sphere stage (4 hpf, left column), shield stage(6phf, middle column) and bud stage (10 hpf, right column) with insets on the right lower corner showing single cells with their boundary marked by a red line injected at the one-cell stage with 2 ng control MO (top row), 1 ng krt4 plus 1 ng krt8 MO (middle row), or with TraCr krt4 and krt8 gRNA (krt4/8 crispant F0; bottom row). Scale bar: 100 µm.
    
    (B) Plot of averaged density of keratin network in individual EVL cells as a function of time (hpf) during epiboly in Tg(actb2:utr-mCherry, krt18:krt18-GFP) embryos injected at the one-cell stage with 2 ng control MO (top row; green, N = 4, n = 4 embryos), 1 ng krt4 plus 1 ng krt8 MO (middle row; orange, N = 4, n = 4 embryos), or with TraCr krt4 and krt8 gRNA (krt4/8 crispant F0; bottom row; pink, N = 2, n = 6 embryos). Error bars as ribbon SD of the mean of individual cells per replicate.
    
    (C) 3-dimensional (3D) plot of keratin intensity, network density and area of EVL cells as a function of time (hpf) during epiboly in Tg(actb2:utr-mCherry, krt18:krt18-GFP) embryos injected at the one-cell stage with 2 ng control MO (top row; green, N = 4, n = 4 embryos), 1 ng krt4 plus 1 ng krt8 MO (middle row; orange, N = 4, n = 4 embryos), or with TraCr krt4 and krt8 gRNA (krt4/8 crispant F0; bottom row; pink, N = 2, n = 6 embryos). Spread of the surface (width) represents data spread, indicating their variability (SD of intensity and area).
    
    (D) Plot of EVL epiboly movement speed as a function of time (hpf) during epiboly starting at sphere stage (4 hpf) until late epiboly stages (9 hpf) in Tg(actb2:utr-mCherry, krt18:krt18-GFP) embryos injected at the one-cell stage with 2 ng control MO (top row; green, N = 3, n = 4 embryos), 1 ng krt4 plus 1 ng krt8 MO (middle row; orange, N = 3, n = 4 embryos), or with TraCr krt4 and krt8 gRNA (krt4/8 crispant F0; bottom row; yellow, N = 2, n = 4 embryos). Error bars as SD of the mean of replicates.

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/Fig4_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Mechanical force percolation within the EVL is dependent on keratin expression.} }

    \label{Figure4Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{Figure4Chapter3} Mechanical force percolation within the EVL is dependent on keratin expression}

    (A) Plots of EVL cell orientations with ellipses representing shape descriptors (long and short axis) of individual EVL cells with the line in the middle marking the orientation of the long axis at consecutive stages during epiboly (4.5, 6, 9, 12 hpf) in curvature corrected Tg(actb2:utr-mCherry, krt18:krt18-GFP) embryos injected at the one-cell stage with 2 ng control MO (top row) or 1 ng krt4 plus 1 ng krt8 MO (middle row). Each cell is colour-coded according to the orientation of the axis (hsv) as shown in the colour bar at the bottom (Red: AV axis orientation, blue: dorsoventral/DV orientation).
    
    (B) Rose plot of EVL cell orientations in curvature corrected Tg(actb2:utr-mCherry,krt18:krt18-GFP) embryos at 6 hpf injected at the one-cell stage with 2 ng control MO (B; N = 4, n = 93 cells) or 1 ng krt4 plus 1 ng krt8 MO (C; N = 4, n = 84 cells).
    
    (C) Bar plots of tissue viscosity measured at the EVL margin using micropipette aspiration at 4-6 hpf and 6-7.5 hpf in Tg(actb2:utr-mCherry, krt18:krt18-GFP) injected at the one-cell stage with 2 ng control MO (B; N = 5, n = 37 embryos) or 1 ng krt4 plus 1 ng krt8 MO (C; N = 5, n = 37 embryos).
    
    (D) Representative quiver plots of EVL tissue recoil flow velocities after cell ablation/wounding (wound centre marked by a red dot) in Tg(actb2:utr-mCherry) embryos at 6 hpf injected with 2 ng control MO (top) or 1 ng krt4 plus 1 ng krt8 MO (bottom). The arrows show the local velocity coloured according to the magnitude (viridis).
    
    (E) Average radial recoil velocity of the tissue flow plotted as a function of distance from the wound centre (0 µm) in Tg(actb2:utr-mCherry) embryos injected with 2 ng control MO (top) or 1ng krt4 plus 1 ng krt8 MO (bottom) at successive time points after cell ablation (from yellow to blue; N = 4, n = 23 embryos).
    
    (F) Left panels: Max intensity projection images of EVL cells in Tg(actb2:utr-mCherry) embryos injected with 2 ng control MO (top) or 1 ng krt4 plus 1 ng krt8 MO (bottom) at 6.5 hpf. Images were overlaid with lines representing their orientation (line angle) and alignment (colour-coded according to the alignment angle; reversed viridis). Right upper panel: schematic showing how cell alignment was determined by measuring the angle from the wound centre ($\theta$1, green lines) and the cell longest cell axis ($\theta$2, blue lines). The alignment was calculated by determining the magnitude of the difference between the angle from the wound centre and the cell longest cell axis (|$\theta$1-$\theta$2|) in successive rows of cells around the wound centre. Right lower panel: bar plot (bottom) of EVL cell alignment in Tg(actb2:utr-mCherry) embryos injected with 2 ng control MO (green) or 1 ng krt4 plus 1 ng krt8 MO (blue) upon control MO injection (green) in successive cell rows (cell 1 - cell 3) around the wound center. (N = 4, n = 23 embryos).

    (G) Plot of tissue recoil velocity as a function of distance from the wound center in Tg(actb2:utr-mCherry) embryos injected with 2 ng control MO (green) or 1 ng krt4 plus 1 ng krt8 MO (blue). N = 4, n = 23 embryos.
    
    (H) Plot of wound area as a function of time upon ablation of EVL cells in Tg(actb2:utr-mCherry) embryos injected with 2 ng control MO (green) or 1 ng krt4 plus 1 ng krt8 MO (orange). N = 4, n = 23 embryos.

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/Fig5_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Vertex model of EVL spreading in the presence and absence of keratin expression.} }

    \label{Figure5Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{Figure5Chapter3} Vertex model of EVL spreading in the presence and absence of keratin expression}

    (A) Plot of extension length of the aspirated EVL tissue in Tg(krt18:Krt18GFP) embryos as a function of time upon micropipette aspiration. Colormap represents developmental time starting at the beginning of epiboly (4 hpf, blue) to later stages (8 hpf, orange).

    (B) Plot of keratin intensity measured inside the pipette in the aspirated EVL tissue in Tg(krt18:Krt18GFP) embryos as a function of time upon micropipette aspiration. Colormap represents developmental time starting at the beginning of epiboly (4 hpf, blue) to later stages (8 hpf, orange).
    
    (C) Representative plot of extension length as a function of time fit to the modified viscoelastic Maxwell model (inset, schematic representing the model) used to measure parameters for the vertex model.
    
    (D) Schematic representing a singular model EVL cell showing keratin filaments (green) at two distinct states of the network activity, dependent on the keratin $K_i$ formation: unpercoalted (left, $K_i$<$K_{th}$, mechanically inactive) and percolated (right, $K_i$>$K_{th}$, mechanically active) with $K_{th}$ being the threshold value of K above which the keratin mechanochemical feedback loop becomes activated.
    
    (E) Schematic diagram of the mechanochemical feedback loop built into the model representing the feedback from pulling forces from the YSL ($F_{YSL}$) stretching the tissue leading to increased pressure $p$ in the tissue that enhances keratin $K$ formation. This increase in keratin $K$ feedsback into the area $A$ of the cells and increases tissue stiffness $\Gamma$ and relaxation time $\tau$.  This increase in tissue stiffness $\Gamma$ and relaxation time $\tau$, in turn, feeds back on the pressure $p$ in the tissue.
    
    (F) Plot of model aspirated tissues heights as a function of time  measured in the mean field model. Mean tissue height (dark red line) is represented as solid line over curves representing estimated parameter variability shown as lines with higher transparency (light red lines).
    
    (G) Plot of model keratin intensities in aspirated tissues as a function of time in the mean field model. Mean keratin intensity (dark green line) is represented as a solid line over curves representing estimated parameter variability shown as lines with higher transparency (light green lines).
    
    (H) Representative model tissue ablations showing keratin expression at t=30s in a EVL model tissue with periodic boundary conditions and mechanochemical feedback model parameter $\beta$=0.05. At t=0s 6 cells are removed to create a hole at the center of the tissue. A tension of 3 $\mu$N is applied on the edges surrounding the removal site to simulate a wound healing process. Colormap on the side represents the keratin intensities of the cells as a function of keratin $K$ with percolated keratin network ($K_i$$>$$K_{th}$) appearing in green and unpercoalted ($K_i$<$K_{th}$) in red-yellow.
    
    (I) Plot of edge tension at t=30s around the ablated wound site in keratin-deficient (below; $\beta$=0) and in wild-type (top; $\beta$=0.005) EVL model tissues. Colormap on the right represents the edge tension. Rainbow: 1.0 $\mu$N to 0.0 $\mu$N).
    
    (J) Plot of wound area (shown in I) as a function of time after ablation in keratin-deficient ($\beta$=0; green) and wild-type with ($\beta$=0.005; orange) EVL model tissues.
    
    (K) Representative vertex model of the EVL tissue at mid epiboly depicting intensity of keratin $K$ (green, percolated and red-yellow, unpercolated) and the orientation of the cell elongation shown as a line in the cell centre (red line). The pulling force by the YSL applied at the edge is represented as red arrows pointing outwards. Stage of pulling corresponds to approximately 7.3 hpf in the zebrafish developing embryo.
    
    (L) Histograms of keratin intensity distributions in simulated tissues as a function of simulated developmental time shown as a colormap of the lines (2.1h violet to 8.3h red.
    
    (M) Histograms of keratin intensity distributions observed within the individual EVL cells in a representative Tg(krt18: Krt18GFP) embryo as a function of development time during epiboly shown as a colormap of the lines (4.7 hpf violet to 9.1 hpf red). 
    
    (N) Plot of mean keratin intensity in the model EVL tissue as a function of time at increasing pulling forces from the YSL $F_{YSL}$  (color-coded). The threshold level of keratin $K_{th}$ is represented as dotted line above which the mechanochemical feedback becomes active in the model.
    
    (O) Plot of mean tissue edge velocity in the presence of wild-type pulling force $F_{YSL}$ as a function of time in keratin-deficient ($\beta$=0; orange) and wild-type ($\beta$=0.005; green) EVL model tissues. 
    
    (P) Plot of mean tissue edge velocity in the presence of three different pulling forces ($F_YSL$): wild-type forces (orange and purple lines), and forces lower (green and pink lines) and higher (blue and brown lines) than the wild-type value. Solid lines represent wild-type ($\beta$=0.005; green, orange and brown lines) and dotted lines keratin-deficient ($\beta$=0; blue, purple and pink lines) EVL model tissues. 

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/Fig6_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Actin flow alignment within the YSL is dependent on keratin expression.} }

    \label{Figure6Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{Figure6Chapter3} Actin flow alignment within the YSL is dependent on keratin expression}

    (A) Maximum intensity projections of the keratin network at the EVL-YSL boundary of Tg(krt18:Krt18GFP) embryos at shield (left column), 75\% epiboly (middle column) and 95\% epiboly (right column) injected with 2 ng control MO (top row) or 1 ng krt4 plus 1 ng krt8 MO (bottom row) into the YSL at sphere stage (3.3 hpf). Scale bar: 25 µm.
    
    (B) Maximum intensity projections of the actin network at the EVL-YSL boundary of Tg(actb2:Utrophinmcherry,krt18:KeratinGFP) embryos at 6 hpf injected with 2 ng control MO (top row) or 1 ng krt4 plus 1 ng krt8 MO (bottom row) into the YSL at sphere stage (3.3 hpf) before (Pre Asp, left column) and after (Post Asp, middle column) micropipette aspiration. Scale bar: 25 µm. Right column: Representative maximum intensity projections of actin and keratin in Tg(actb2:Utrophin-mcherry,krt18:Keratin18GFP) at 3.3 hpf injected with 2 ng control MO (top panel) or 1 ng krt4 plus 1 ng krt8 MO (bottom panel). Images are overlaid with quiver plots of retrograde actin flows within the YSL. Scale bar: 50 µm.
    
    (C) Plot of EVL epiboly movement speed as a function of time (hpf) during epiboly in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos injected with 2ng control MO (green) or 1 ng krt4 plus 1 ng krt8 MO (orange) into the YSL at sphere stage (3.3 hpf). Error bars as ribbon SD of mean. N = 4, n = 8 embryos.
    
    (D) Plot of retrograde actin flow velocity in the YSL as a function of distance from the EVL-YSL boundary in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos at shield stage (6 hpf) injected with 2 ng control MO (green) or 1 ng krt4 plus 1 ng krt8 MO (orange) into the YSL at sphere stage (3.3 hpf). Error bars as ribbon SD of mean. N = 3, n = 6 embryos.
    
    (E) Box plot of actin intensity in a fixed region of interest (ROI) within the YSL close to the point of micropipette aspiration before (pre Asp) and after (post Asp) aspiration in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos at 3.3 hpf injected with 2 ng control MO (green boxes) or 1 ng krt4 plus 1 ng krt8 MO (orange boxes)  into the YSL at sphere stage (3.3 hpf). N = 4, n = 13 embryos. Boxes represent quartiles of the data and error bars the spread (pvalues: ***<0.001, **<00.1 paired t test).
    
    (F) Box plot of actin flow alignment within the YSL at 6 hpf in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos injected with 2 ng control MO (green boxes) or 1 ng krt4 plus 1 ng krt8 MO (orange boxes)  into the YSL at sphere stage (3.3 hpf). Boxes represent quartiles of the data and error bars the spread. N = 3, n = 6 (***<0.001, paired t test).

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.65\linewidth]{graphics/FigS1_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Keratin expression and localization during epiboly} }

    \label{FigureS1Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS1Chapter3} Supplement 1: Keratin expression and localization during epiboly}  
    
    (A) Bar plot of fold change of keratin 18, keratin 8, keratin 4, and keratin 5 expression in embryos at 1K (3.3hpf), 30\% epiboly (4.5 hpf), shield (6hpf) and bud (9hpf) stages measured by qPCR. 

    (B) Maximum Intensity projections of krt8 mRNA fluorescence via in situ hybridization in WT embryos at 9 hpf. Scale bar: 25 $\mu$m

    (C) Maximum Intensity projections of keratin network in Tg(krt18: Krt18GFP) embryos at shield (6hpf), 75\% epiboly (8.5 hpf) and bud (10 hpf) stages injected of 50pg Krt4-mcherry RNA at the one-cell stage. Left column, keratin 18 (green); middle column, keratin 4 (red); right column, keratin 4 and 18 colocalization (white; co-localization index R below threshold 0.005). Scale bar: 100 $\mu$m

    (D) Maximum Intensity projections of keratin network in Tg(krt18: Krt18GFP) embryos at shield (6hpf), 75\% epiboly (8.5 hpf) and bud (10 hpf) stages injected of 50pg Krt4-mcherry RNA at the one-cell stage. Scale bar: 25 $\mu$m

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/FigS2_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Tension-dependent regulation of keratin expression within the EVL} }

    \label{FigureS2Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS2Chapter3} Supplement 2: Tension-dependent regulation of keratin expression within the EVL}

(A) Maximum intensity projection images of keratin expression in Tg(actb2: Utrophin- mcherry, krt18:Krt18GFP) embryos at sphere (4 hpf, left column), shield (6 hpf, right column), and 90\% epiboly (9.5hpf, right column) stages injected with (0.2\% phenol red, control, top row), 100pg caMypt (middle row), or 50 pg caRhoA (bottom row) directly into the YSL at 3.3 hpf (control, caMypt) or into marginal cells at 3 hpf (caRhoA) Scale bar: 25 $\mu$m

(B) Plot of average keratin intensity as a function of epiboly stages in Tg(krt18:KrtGFP) control (green, N=3, n= 5 embryos), caMypt (orange, n=3, n=5 embryos) and caRhoA mRNA injected embryos(pink, N=3, n=4 embryos) at 3.3 hpf into the YSL as described in Figure 2(A- C). Error bars as ribbons SD of mean.

(C) Plot of average density of keratin network as a function of epiboly stages in Tg(actb2:Utrophin-mcherry, krt18:Krt18-GFP) control (green, N=3, n= 6 embryos), caMypt (orange, n=3, n=4 embryos) and caRhoA mRNA injected embryos (pink, N=3, n=3 embryos) at 3.3 hpf into the YSL as described in (B). Error bars as ribbon SD of mean of individual cells per replicate.

(D) Plot of EVL epiboly movement speed as a function of time (hpf) during epiboly starting at sphere stage (4hpf) until late epiboly stages (9 hpf) in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos injected at 3.3hpf with control (green, N=3, n= 6 embryos), caMypt (orange, n=3, n=4 embryos) and caRhoA mRNA (pink, N=3, n=3 embryos). Error bars as ribbon SD of mean of individual cells per replicate.

(E) Maximum intensity projection images of keratin expression in individual primary cells from Tg(krt18:Krt18GFP) embryos dissociated at 4 hpf (left) and 6.5 hpf (right). The outline of the cell is shown as the red dotted line. Scale bar: 15 $\mu$m

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/FigS3_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Effect of dominant negative keratin 18 expression on keratin network formation and EVL epiboly} }

    \label{FigureS3Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS3Chapter3} Supplement 3: Effect of dominant negative keratin 18 expression on keratin network formation and EVL epiboly}

(A) Maximum intensity projection images of keratin expression in Tg(actb2: Utrophin- mcherry, krt18:Krt18GFP) at sphere (4hpf, left column), shield (6phf, middle column) and bud (10 hpf, right column) stages injected with 0.2\% phenol red (control, top row) or 150 pg dnKrt18 RNA (bottom row) at one-cell stage. Scale bar: 100 $\mu$m

(B) Maximum intensity projection images of keratin expression in Tg(actb2: Utrophin- mcherry, krt18:Krt18GFP) at sphere (4hpf, left column), shield (6phf, middle column) and bud (10 hpf, right column) stages injected with 0.2\% phenol red (control, top row) or 150 pg dnKrt18 RNA (bottom row) at one-cell stage. Scale bar: 25 $\mu$m

(C) Plot of EVL epiboly movement speed as a function of time (hpf) during epiboly starting at sphere (4 hpf) until late epiboly (9 hpf) stages in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos injected with 0.2\% phenol red (control, green) or 150 pg dnKrt18 RNA (orange) at one-cell stage. (N=2, n=6 embryos).

(D) Maximum intensity projection images of EVL during rupture at 13.5 hpf in Tg(actb2: Utrophin-mcherry, krt18:Krt18GFP) embryos injected with 1ng krt4 plus 1ng krt8 MO at one cell stage showing the EVL before (top) and after the EVL ruptures (bottom).

(E) Bar plot of quantification of death rate in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos injected at the one-cell stage with 2ng control MO (top row; green, N=4, n=4 embryos), 1ng krt4 plus 1ng krt8 MO (middle row; orange, N=4, n=4 embryos), or with TraCr krt4 and krt8 gRNA (krt4/8 crispant F0; bottom row; yellow, N=2, n=6 embryos).

(F) Exemplary plot of tissue extension as a function of EVL tissue aspiration time, displaying a linear response upon aspiration and retraction(green). Plot of fluorescence intensity of keratin as a function of aspiration in the same embryo measured in the medial plane of the pipette in the same embryo (orange)

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/FigS4_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Effect of dominant negative keratin 18 expression on keratin network formation and EVL epiboly} }

    \label{FigureS4Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS4Chapter3} Supplement 4: Effect of dominant negative keratin 18 expression on keratin network formation and EVL epiboly}

(A) Exemplary plots of EVL cell orientations with ellipses representing shape descriptors (long and short axis) of individual EVL cells with the line in the middle marking the orientation of the long axis at consecutive stages during epiboly (4.5, 6, 9, 12 hpf) in curvature corrected Tg(actb2:Utrophin-mcherry, krt18:Krt18GFP) embryos injected at high stage (3.3 hpf) directly into the YSL to interfere with keratin network formation within the YSL specifically. Each cell is colour-coded according to the orientation of the axis (hsv) as shown in the colour bar at the bottom (Red: AV axis orientation, blue: dorsoventral/DV orientation).

(B) Rose plot of EVL cell orientations in curvature corrected Tg(actb2:Utrophinmcherry,krt18:Krt18GFP) embryos at 6 hpf injected at high stage (3.3 hpf) into the YSL with 2 ng control MO (B; N=4, n=93 cells) or 1ng krt4 plus 1ng krt8 MO (C; N=4 n= 84 cells).

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/FigS5_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Micropipette aspiration experimental data and Maxwell model} }

    \label{FigureS5Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS5Chapter3} Supplement 5: Micropipette aspiration experimental data and Maxwell model}

    (A) Scatterplot of measured time scales $\tau_r$ (blue, square) and $\tau_r'$ (green, rhombus) as a function of time during development in the EVL aspiration experiments obtained from fits of the height relaxation to equation (S4) during the aspiration ($\tau_r$, blue) and release phases ($\tau_r'$,green), respectively, using the method described in (Figure 5A-C). Scatter plot of measured time scale $\tau_\kappa$ for the relaxation of the keratin $K(T)$ as a function of time during development as depicted in (Figure 5B) after pipette release (yellow, circle), from a fit to exponential form $e^{-\tau/\tau_\kappa }$. Line plots depict the average with confidence intervals for  $\tau_r$ (blue), $\tau_r'$ (green) and  $\tau_\kappa$ (yellow).
    
    (B) Scatterplot of measured EVL tissue aspiration velocities after aspiration (square, blue), after aspiration but before release (circle, yellow), after release (rhombus, green) as extracted by linear fits to height curves $l(t)$, where $\tau_{release}$ is the time at which the pressure is removed. Line plots depict the average with confidence intervals for  $l(t>0)$ (blue), $l \tau_{release^+})$ (green) and  $l(\tau_{release}^-)$ (yellow).
    
    (C) Scatterplot of tissue relaxation time scales $\tau$ during aspiration (square, blue) and $\tau'$ during release (circle, yellow) as a function of time during development measured from fits to equation (S4) of the EVL aspiration experiments used to extract Maxwell model parameters (equations (S5)). Line plots over the scatter depict the average with confidence intervals shaded around the line.
    
    (D) Scatterplot of tissue elastic constant $\kappa_k$ as a function of time during development, measured from fits to the tissue extension velocities in equation (S5). Line plots over the scatter depict the average with confidence intervals shaded around the line. 
    
    (E) Scatterplot of substrate friction coefficient $\zeta$  as a function of time during development measured from fits to the tissue extension velocities in equation (S5). Line plots over the scatter depict the average with confidence intervals shaded around the line.
    
    (F) Scatterplot of internal tissue tension $\tau$ as a function of time during development measured from  fits to the tissue extension velocities in equation (S5). Line plots over the scatter are the average with confidence intervals shaded around the line.

    See SI section 1 for model details.


\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/FigS6_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Time and velocity scales for the EVL micropipette aspiration experiment and parameters of the Maxwell model} }

    \label{FigureS6Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS6Chapter3} Supplement 6: Time and velocity scales for the EVL micropipette aspiration experiment and parameters of the Maxwell model}

    (A) Scatter plot of measured time scales $\tau_r$ (blue, square) and $\tau_r'$ (green, rhombus) in the EVL aspiration experiments from fits of the height relaxation to equation (S4) during the aspiration and release phases, respectively, using the method described in Figure Supplement 5C. Measured time scale $\tau_\kappa$ for the relaxation of the keratin $\kappa(T)$K(T) as depicted in (figure supplement 5B) after pipette release (yellow, circle), from a fit to exponential form $e^{-\tau/\tau_\kappa} $. Line plots depict the average with confidence intervals for  $\tau_r$ (blue), $\tau_r'$ (green) and  $\tau_\kappa$  (yellow).

    (B) Scatter plot of measured velocities in the EVL aspiration experiments as extracted by linear fits to the height curves $l$(t) (figure supplement 5A) where $\tau_{release}$ is the time where the pressure is removed. 
    
    (C-F) Fits to equation (S4) of the EVL aspiration experiments during aspiration and release inverted to extract Maxwell model parameters (equations (S5)), i.e. tissue relaxation time scales $\tau$ during aspiration and $\tau'$ during release (C), tissue elastic constant $\kappa$ (D), substrate friction coefficient $\gamma$ (E), and internal tissue tension T (F). Line plots over the scatter are the average with confidence intervals shaded.

    See SI section 1 for model details.

\newpage
\begin{figure}[htbp]
    \centering
    \includegraphics[width=0.85\linewidth]{graphics/FigS7_thesis.pdf}
    \captionsetup{width=0.95\linewidth}
    \caption{\textbf{Actin-keratin flows within the YSL } }

    \label{FigureS7Chapter3}
\end{figure}
\noindent\textbf{Figure ~\ref{FigureS7Chapter3} Supplement 7: Actin-keratin flows within the YSL }

    (A) Z-plane reslice image of a cross-sectional view of the YSL at the EVL margin in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos. EVL and YSL are outlined by a white dotted line, and keratin filaments within the YSL are marked by white arrows. Scale bar: 15 $\mu$m. 

    (B) Plot of average flow velocity of keratin (green) and actin (orange) within the YSL in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos measured using PIV as a function of distance from the EVL-YSL boundary.

    (C) Plot of a representative particle image velocimetry flows of keratin filaments (virdis) and actin cortex (plasma) at the EVL-YSL boundary in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos measured using PIV. The dotted box outlines the region of interest represented in (C’) showing a high-magnification view of the keratin flow field (viridis) at the EVL-YSL boundary. Scale bar: 1 $\mu$m/min.
    
    (D) Box plot of the hydrodynamic length of the YSL actin network measured by UV-laser cutting of the actin cortex in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos at shield stage (6.5 hpf) injected with control MO (green, N = 4, n = 15 embryos) or 1 ng krt4 plus 1 ng krt8 MO (orange, N = 4, n = 17 embryos) into the YSL at high stage (**<00.1 paired t test). 
    
    (E) Box plot of initial recoil velocities of the actin cortex after laser ablations in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos injected with control MO (green, N = 4, n = 15 embryos) or 1 ng krt4 plus 1 ng krt8 MO (orange, N = 4, n = 17 embryos) in the YSL at high stage (**<00.1 paired t test).
    
    (F) Plot of actin intensity recovery after UV-laser cutting as a function of time measured within a region of interest at the cut within the YSL actin cortex in Tg(actb2:Utrophinmcherry, krt18:Krt18GFP) embryos at shield stage (6.5 hpf) injected with control MO (green, N = 4, n = 15 embryos) or 1 ng krt4 plus 1 ng krt8 MO (orange, N = 4, n = 17 embryos) into the YSL at sphere stage (3.3 hpf). 


\newpage
\subsubsection{Video legends}
These legends correspond to the movies available as supplementary material that can be downloaded from \href{https://seafile.ist.ac.at/d/d45a8a27d35a4f40902c/}{Videos}

\href{https://seafile.ist.ac.at/f/d253f37bd3244278b6cd/}{Video 1: Keratin expression within the EVL during epiboly}
Time-lapse of keratin expression (right) and network organization (left) in Tg(krt18:Krt18GFP) embryos during epiboly (4 - 9.5 hpf). Lateral view. Frame rate 11.25 min/frame. Scale bar: 100 (left) and 30 $\mu m$ (right). 

\href{https://seafile.ist.ac.at/f/ed8f06f62057401cbdfd/}{Video 2: Keratin expression within the EVL in embryos with reduced YSL pulling force}
Time-lapse of keratin expression within the EVL upon reduced pulling forces from the YSL in a representative  Tg(krt18:Krt18-GFP) control embryo injected with 0.2\% phenol red (left) and an embryo injected with 100 pg CAMypt into the YSL (right) imaged from from 4.5-11 hpf. Lateral view. of the embryos Frame rate 10 min/frame. Scale bar: 100 $\mu m$.

\href{https://seafile.ist.ac.at/f/54c7a6b586034f6591b1/}{Video 3: Keratin network maturation in EVL cells of embryos with reduced YSL pulling force}
Time-lapse of keratin network maturation in EVL cells upon reduced pulling forces from the YSL  in a representative  Tg(krt18:Krt18-GFP) control embryo injected with 0.2\% phenol red (left) and an embryo injected with 100 pg CAMypt into the YSL (right) imaged from 4.5-7.1 hpf. Lateral view near the EVL-YSL boundary. Frame rate 10 min/frame. Scale bar: 25 $\mu m$.

\href{https://seafile.ist.ac.at/f/abe6c0bfa3e1430f8e5d/}{Video 4: Keratin expression within the EVL of embryos with enhanced YSL pulling force}
Time-lapse of keratin expression upon increased pulling forces from the YSL in representative Tg(krt18:Krt18-GFP) control embryo injected with 0.2\% phenol red (left) into the YSL and 50 pg CARhoA into marginal blastomeres at 3.3 hpf (right) imaged from 4.5-10.3 hpf. Lateral view. Frame rate10 min/frame. Scale bar: 100 $\mu m$. 

\href{https://seafile.ist.ac.at/f/c6e473cb3ee6488ba813/}{Video 5: Keratin network maturation in EVL cells of embryos with enhanced YSL pulling force}
Time-lapse of keratin network maturation upon increased pulling forces from the YSL in a representative Tg(krt18:Krt18-GFP) control embryo injected with 0.2\% phenol red (left) and an embryo injected with 50 pg CARhoA into marginal blastomeres at 3.3 hpf (right) imaged from 4.5-7.5 hpf. Lateral view near the EVL-YSL boundary. Frame rate 10 min/frame. Scale bar: 25 $\mu m$. 

\href{https://seafile.ist.ac.at/f/0b9daedeca404e6c932b/}{Video 6: Keratin expression within the EVL of embryos with reduced keratin type II expression}
Time-lapse of keratin expression in representative Tg(krt18:Krt18-GFP) embryos injected at the one-cell stage either with 2 ng control MO (control, left) or 1 ng krt4 plus 1 ng krt8 MO (right) imaged from 4-15.75 hpf. Lateral view. Frame rate 10.25 min/frame. Points of rupture in keratin morphant embryos are marked with asteriks. Scale bar: 100 $\mu m$.

\href{https://seafile.ist.ac.at/f/96081207f36f4defb84a/}{Video 7: Keratin network maturation in EVL cells of embryos with reduced keratin type II expression}
Time-lapse of keratin expression in representative Tg(krt18:Krt18-GFP) embryos injected at the one-cell stage either with 2 ng control MO (control, left) or 1 ng krt4 plus 1 ng krt8 MO (right) imaged from 4.5-8.4 hpf. Lateral view. Frame rate10 min/frame. Scale bar: 25 $\mu m$.

\href{https://seafile.ist.ac.at/f/a52ca387ebe34ac0bbb7/}{Video 8: Changes in keratin expression upon EVL aspiration}
Time lapse of EVL aspiration in a representative Tg(krt18:Krt18-GFP) embryo using a 60 µm pipette imaged by brightfield (left) and confocal (keratin, green, right) microscopy. Z-plane in the centre of the pipette. Frame rate 1 sec/frame. Scale bar: 25 $\mu m$.

\href{https://seafile.ist.ac.at/f/08a27eb22abc4b43bb6d/}{Video 9: Changes in keratin expression during EVL wound closure after cell ablation}
Time lapse of EVL in a representative Tg(actb2:Utrophin-mcherry, krt18:Krt18-GFP) embryo imaged before (pre) and after (post) UV laser-mediated cell ablation showing keratin (right, green) and actin (left, orange). Frame rate 20 sec/frame. Scale bar: 25 $\mu m$.

\href{https://seafile.ist.ac.at/f/42886b6f3cbe4f59b684/}{Video 10: Wound closure in control and keratin-deficient embryos}
Time lapse of EVL response in representative Tg(actb2:Utrophin-mcherry) embryo imaged before (pre) and after (post) UV laser-mediated cell ablation injected at the one-cell stage either with 2 ng control MO (control, left) or 1 ng krt4 plus 1 ng krt8 MO (right). Frame rate 20 sec/frame. Scale bar: 25 $\mu m$. 

\href{https://seafile.ist.ac.at/f/c647a1fae1794b6ebcce/}{Video 11: E-cadherin expression in control and keratin-deficient embryos}
Time lapse of E-cadherin expression in Tg(cdh1-YFP)xt17 embryos injected at the one-cell stage either with 2 ng control MO (control, left) or 1 ng krt4 plus 1 ng krt8 MO (right) starting at the beginning of epiboly (4 hpf) to failure of EVL integrity (14 hpf). Frame rate 10 min/frame. Scale bar: 25 $\mu m$. 

\href{https://seafile.ist.ac.at/f/83eead7bd7d142c2805b/}{Video 12: Occludin-b expression in control and keratin-deficient embryos}
Time lapse of Occludin b expression in Tg(oclnb-GFP)pd1126 embryos injected at the one-cell stage either with 2 ng control MO (control, left) or 1 ng krt4 plus 1 ng krt8 MO (right) starting at the beginning of epiboly (4 hpf) to the end of epiboly (10 hpf). Frame rate 10 min/frame. Scale bar: 25 $\mu m$. 

\href{https://seafile.ist.ac.at/f/824dfd1fd3844bf18bca/}{Video 13: Jup-a expression in control and keratin-deficient embryos}
Time lapse of Jup-a expression in Tg(her4.1:jupa-EGFP)  embryos injected at at the one-cell stage either with 2 ng control MO (control, left) or 1 ng krt4 plus 1 ng krt8 MO (right) starting at the beginning of epiboly (4 hpf) to the end of epiboly (10 hpf). Frame rate 10 min/frame. Scale bar: 25 $\mu m$.

\href{https://seafile.ist.ac.at/f/8954c87e50154122aec8/}{Video 14: Simulated EVL wound closure in model with and without keratin feedback onto mechanics}
Time lapse of simulated wound closure following ablations in EVL model tissues without keratin mechanical feedback (‘keratin deficient’, top) and with keratin mechanical feedback (‘control’, bottom). Keratin concentration $K_i$ in the cell is shown by colour ($K_i$: 0-150 red-yellow, $K_i$: 150-350 green). Tensile stress in the tissue $t_i$ from the junctional tension of the cells around the wound is shown by the colour of the cell edges (rainbow $t_i$: 0.0 – 0.6).

\href{https://seafile.ist.ac.at/f/caa88eeb19764574b48e/}{Video 15: Simulated EVL tissue undergoing epiboly} 
Time lapse of simulated EVL model tissue being stretched by radially outward forces increasing with time applied on the margin of the tissue, simulating the forces generated by the YSL on the EVL edge in the embryo. Keratin intensity $K_i$ shown as the colour of the cells (left) increases with time in response to the stress experienced by the tissue under this force, starting from unpercolated levels ($K_i$< $K_{th}$, red-yellow) and progressing in each cell over the threshold keratin levels to percolated levels ($K_i$> $K_{th}$, green). Elevation of keratin levels in each cell over the threshold level is stochastic due to the heterogeneity in the mechanical effects on the disordered tissue. Tensile stress in the tissue $t_i$ is shown by the colour of the cell edges  (rainbow $t_i$: 0.0 - 0.6).
