Epithelial fusion is mediated by a partial epithelial-mesenchymal transition

Epithelial fusion is mediated by a partial epithelial-mesenchymal transition | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Epithelial fusion is mediated by a partial epithelial-mesenchymal transition View ORCID Profile Varsha N. Tamilkumar , Harsha Purushothama , View ORCID Profile Raj K. Ladher doi: https://doi.org/10.1101/2025.04.21.649757 Varsha N. Tamilkumar 1 National Centre for Biological Sciences, Tata Institute for Fundamental Research , GKVK PO, Bellary Road, Bangalore, India , 560065 2 The University of Trans-Disciplinary Health Sciences and Technology (TDU) , Attur Layout, Bangalore, India , 560064 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Varsha N. Tamilkumar Harsha Purushothama 1 National Centre for Biological Sciences, Tata Institute for Fundamental Research , GKVK PO, Bellary Road, Bangalore, India , 560065 Find this author on Google Scholar Find this author on PubMed Search for this author on this site Raj K. Ladher 1 National Centre for Biological Sciences, Tata Institute for Fundamental Research , GKVK PO, Bellary Road, Bangalore, India , 560065 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Raj K. Ladher For correspondence: rajladher{at}ncbs.res.in Abstract Full Text Info/History Metrics Supplementary material Preview PDF ABSTRACT Epithelial fusion is a fundamental morphogenetic process critical for the closure and compartmentalisation of developing organs. While widely studied in systems such as neural tube and palatal closure, the cellular transitions that enable fusion remain poorly understood. Here, we investigate epithelial fusion during chick otic vesicle (OV) closure and identify a transient population of cells at the epithelial interface that mediate this process. These otic edge (OE) cells exhibit distinct morphology, reduced apicobasal polarity, and dynamic junctional remodelling, including altered distribution of ZO-1, CDH1, and RAC1. Notably, OE cells lack basal contact and display high sphericity, consistent with a partial epithelial-to-mesenchymal transition (EMT) phenotype. Transcriptomic profiling of microdissected tissues reveals that OE cells constitute a transcriptionally distinct population, enriched for EMT regulators, ECM remodelling genes, and WNT pathway components. Among these, the transcription factors Grhl2 and Sp8 were specifically expressed at the OE and exhibited opposing roles in epithelial identity. CRISPR-Cas9 mediated knockdown of either gene led to disrupted CDH1 localisation, loss of OE cell morphology, and failure in epithelial segregation. These results suggest that epithelial fusion requires a regulated, hybrid EMT state that balances junctional plasticity with tissue cohesion. Our findings demonstrate that fusion-competent epithelial cells are not merely passive participants but actively modulate their shape, polarity, adhesion, and genetic identity to enable morphogenesis. INTRODUCTION Epithelial fusion is a key morphogenetic process during organogenesis, enabling the shaping and compartmentalisation of developing tissues. During fusion, the edges of two epithelia are brought into close apposition, and through transient remodelling of junctional components, cells from opposing edges establish new contacts. This process converts what was once a continuous epithelial sheet into two distinct epithelia ( Pai et al., 2012 ). Epithelial fusion is a conserved mechanism across species and occurs in a variety of developmental contexts, including Drosophila gastrulation and trachea formation, as well as vertebrate processes such as optic fissure closure, otic vesicle formation, body wall closure, palatogenesis, and neural tube closure ( Alvarez and Navascués, 1990 ; Geelen and Langman, 1979 ; Gestri et al., 2018 ; Nikolopoulou et al., 2017 ). Failures in epithelial fusion are associated with several common birth defects, including coloboma, cleft palate, neural tube defects, and omphalocele ( Ray and Niswander, 2012 ). Studies on fusion events such as optic fissure closure, palatal shelf fusion, and neural tube closure have provided critical insights into the molecular and cellular processes that govern fusion. A recurring theme in these systems is the importance of the epithelial edge, the interface between two distinct epithelial domains. For instance, in neural tube closure, this is the boundary between neural and non-neural ectoderm; in palatogenesis, between the nasal and oral periderm; and in optic fissure closure, between the neural retina and retinal pigmented epithelium. Fusion can proceed from the apical surface, as in neural tube and palatal shelf closure, or from the basal side, as seen in optic fissure fusion ( Chan et al., 2020 ; Lan et al., 2015 ; Nikolopoulou et al., 2017 ). As epithelial edges converge, the leading-edge cells undergo significant remodelling to mediate fusion. In several systems, including wound healing, optic fissure closure, and Drosophila amnioserosa closure, these cells display a distinct character; they form actin-based protrusions, show reduced apical-basal polarity, altered intercellular junctions, and often lie adjacent to discontinuities in the basement membrane. These traits are suggestive of a transient loss of epithelial identity, a phenotype commonly referred to as a partial epithelial-to-mesenchymal transition (partial EMT) ( Bahri et al., 2010 ; Futterman et al., 2011 ; Gestri et al., 2018 ; Nikolopoulou et al., 2017 ). EMT is not a binary switch but a continuum of stable intermediary states with varying degrees of epithelial and mesenchymal features ( Nieto et al., 2016 ). In the context of epithelial fusion, the moderation of epithelial traits does not always involve the acquisition of mesenchymal properties; often, it is simply the loss of some epithelial characteristics, such as tight junction integrity or apical-basal polarity ( Bahri et al., 2010 ; Gestri et al., 2018 ; Nagai et al., 2022 ; Nikolopoulou et al., 2017 ). This partial EMT phenotype confers the flexibility required for leading-edge cells to undergo junctional remodelling and interact with the ECM, features essential for successful fusion. Interestingly, similar moderated epithelial states are observed in disease contexts such as cancer progression and fibrosis, where cells require plasticity to migrate or invade ( Aiello et al., 2018 ; Cheung et al., 2013 ). The inner ear is derived from the otic placode, a thickened epithelial region of the embryonic surface ectoderm. During development, the placode invaginates, bringing the border between the otic and non-otic ectoderm into close proximity. Once this contact is established, the otic vesicle (OV) pinches off from the overlying ectoderm and becomes internalised within the cranial mesenchyme ( Ladher, 2017 ; Sai and Ladher, 2015 ). Fusion between the apposed epithelial edges facilitates this segregation. Given the parallels with other epithelial fusion systems, we hypothesised that cells mediating otic vesicle fusion transiently adopt a less epithelial, more fluidic state that enables fusion and tissue segregation. In this study, we characterise the cellular and molecular attributes of the otic epithelial edge (OE) cells involved in fusion. Morphologically, these cells are distinct from neighbouring epithelial populations, displaying altered shape and polarity. Through protein localisation and transcriptomic analyses, we identify features that mark OE cells as a specialised population within the continuous epithelial sheet. Furthermore, using CRISPR-Cas9–mediated mosaic knockdown of the transcription factors Grhl2 and Sp8, we show that perturbing the balance of epithelial and mesenchymal programs disrupts adherens junction dynamics and the characteristic cell shape changes required for fusion. These phenotypes were accompanied by a failure in epithelial segregation, suggesting that a regulated cell state transition, controlled by a network of transcription factors, is essential for otic vesicle fusion. Our transcriptomic profiling and in situ hybridisation studies support the existence of this regulatory network and its role in shaping fusion-specific cell behaviour. MATERIALS AND METHODS Chicken Eggs Fertilized Kaveri chicken eggs were obtained from the Central Poultry Development Organisation and Training Institute, Hessaraghatta, India. Eggs were stored at 25°C until use and subsequently incubated at 37°C in a humidified incubator. Embryos were staged according to the Hamburger and Hamilton (1951) developmental staging system. Immunohistochemistry (IHC) Cryosections were washed three times in 0.1% Tween-20 in PBS (PBST) for 15 minutes each. Blocking was performed at room temperature (RT) for 1 hour in PBST containing 3% heat-inactivated goat serum and 2 mg/mL bovine serum albumin (BSA). Sections were then incubated with primary antibodies (see Table 1 for details and dilutions), diluted in the blocking solution, for approximately 36–40 hours at 4°C. View this table: View inline View popup Table 1: List of antibodies and their dilutions Following primary incubation, sections were washed three times in PBST (15 minutes each), and re-blocked for 1 hour at RT. Secondary antibodies and fluorophore-conjugated phalloidin (see Table 1 for details and dilutions) were diluted in blocking solution and applied to the sections for 3 hours at RT. After incubation, sections were washed again three times in PBST for 15 minutes each. Nuclei were counterstained with DAPI (7–10 minutes), followed by a final wash in PBS for 10 minutes. Coverslips were mounted using Fluoroshield mounting medium. Images were acquired using an Olympus FV3000 confocal microscope equipped with high-sensitivity detectors, using 10× and 63× objectives, at a resolution of 2048 × 2048 pixels. Scanning electron microscopy (SEM) Embryos were accessed using paper windows and immediately fixed in primary fixative containing 2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 M sodium cacodylate buffer at 4°C for 24 hours. Following fixation, embryos were washed three times in 0.1 M sodium cacodylate buffer on ice. Post-fixation was performed in 1% osmium tetroxide (OsO□), chilled on ice, for 2 hours. To eliminate residual yolk and paper fibres, embryos were rinsed three times with filtered Milli-Q water. Dehydration was initiated with two washes in 50% ethanol for 20 minutes each, followed by a graded ethanol series (70%, 90%, 95%, 99.5%, and 100%) for 15 minutes per step. Final dehydration involved two 20-minute washes in 100% absolute ethanol. Dehydrated embryos were subjected to critical point drying (CPD) using a Leica EM CPD300, following the standard “sludge worm” protocol. Dried specimens were sputter-coated with a 10 nm gold layer using an Emitech K550X Sputter Coater. Imaging was performed on a Zeiss Merlin Compact VP SEM system. For imaging the cellular morphology within the otic vesicle, embryos were carefully bisected through the otic vesicle using a scalpel prior to primary fixation. Imaris-surface module Transverse cryosections from HH16 and HH17 stage chick embryos were stained with fluorescently labelled phalloidin to visualise cortical F-actin, which demarcates the approximate cell boundaries. Z-stacks were acquired from these sections and processed using the Imaris (Bitplane) surface tool to reconstruct the three-dimensional boundaries of individual cells of interest manually. Cell outlines were traced across the z-stacks to generate surface-rendered objects. Three distinct epithelial cell populations were analysed: (1) pseudostratified cells within the otic vesicle, (2) rounded interfacial cells at the fusion edge, and (3) squamous cells of the surface ectoderm that are continuous with the otic vesicle epithelium. The shape descriptor “sphericity” was used to distinguish these populations quantitatively. Statistical significance among the three groups was assessed using one-way ANOVA. Protein localisation analysis using ImageJ Sections of HH16 and HH17 stage chick embryos were immunostained for various proteins using the immunohistochemistry protocol described above. To quantify the spatial distribution of protein localization at the cellular level, a single optical section from each z-stack was selected, and individual cell boundaries were manually traced using the freehand line tool in Fiji (ImageJ). Protein intensity along the cell periphery was extracted using the Plot Profile function. The coefficient of variation (CV) of pixel intensity values along the circumference was calculated for each cell. The CV, defined as the standard deviation divided by the mean intensity along the cell boundary, served as a metric to assess heterogeneity in protein distribution. A lower CV indicated a more uniform (diffuse) localization, whereas a higher CV reflected a punctate or spatially restricted distribution. This method allowed us to quantitatively compare the expression patterns of proteins across the three epithelial cell types analyzed: pseudostratified otic vesicle cells, rounded edge cells, and squamous surface ectoderm cells. Whole-Mount In Situ Hybridisation All solutions used were treated with 0.1% diethyl pyrocarbonate (DEPC) to inactivate RNases. DEPC was added to a final concentration of 0.1% and incubated at 37°C overnight. Solutions and components that could not be autoclaved were prepared using DEPC-treated water to ensure RNase-free conditions. Chick embryos were fixed in 4% paraformaldehyde (PFA) for 2 hours at room temperature (RT), followed by thorough washing in DEPC-treated PBS (DEPC-PBS) to remove residual fixative. The allantoic membrane was removed, and embryonic cavities were punctured to facilitate reagent penetration. Embryos were dehydrated through a graded methanol/PBS series (25%, 50%, 75%, 90%, 95%, 99%) and washed twice with 100% methanol before storage at –20°C in 100% methanol. For in situ hybridisation, embryos were rehydrated gradually into PBST (PBS with 0.1% Tween-20) on ice. Proteinase K treatment was performed at a concentration of 10 µg/mL in DEPC water, with the incubation time corresponding to the embryo’s Hamburger-Hamilton (HH) stage (e.g., HH17 embryos were treated for 17 minutes) on ice. Following digestion, embryos were washed twice with PBST for 5 minutes each on ice and post-fixed in 4% PFA with 0.2% glutaraldehyde in PBST for 20 minutes at RT. Three 5-minute washes followed post-fixation in PBST. Embryos were then transferred into pre-hybridisation solution and allowed to sink before being incubated in fresh pre-hybridisation solution at 65°C for 1 hour. Digoxigenin (DIG)-labelled RNA probes were added to a final concentration of 1 µg/mL, and hybridisation was carried out overnight at 65°C. Post-hybridisation washes included a series of high-stringency washes at 65°C, followed by aqueous washes in MABT (Maleic acid buffer with Tween-20). Blocking was performed using the Roche Blocking Reagent according to the manufacturer’s instructions. The blocking solution was supplemented with 20% heat-inactivated goat serum before incubation with anti-DIG-AP antibody (Roche) at a dilution of 1:2000, overnight at 4°C. Embryos were washed extensively with MABT over a full day at RT to reduce non-specific staining. Colour was developed in NTMT buffer using NBT/BCIP substrate at RT without agitation. Alternatively, BM-Purple substrate (Roche) was used for colourimetric detection. After colour development, embryos were equilibrated in 60% glycerol for optical clearing and imaged using a stereomicroscope. DIG-labelled RNA probes were synthesised using Roche reagents and primers listed in Table 2 , following the manufacturer’s protocol. View this table: View inline View popup Download powerpoint Table 2: List of primers used to generate WhISH antisense probes. Tissue Dissection and RNA Preparation All dissection tools and Petri dishes were treated to eliminate RNase contamination. Tools were baked at 200°C for 5 hours, and all surfaces, including microscope stages and workbenches, were cleaned thoroughly with RNaseZap (Thermo Fisher Scientific). PBS used during dissection was DEPC-treated and autoclaved before use. Dispase (2.4 U/mL) was prepared in DEPC-treated PBS, aliquoted, and stored at –20°C. Dispase was used to remove mesenchymal cells adhering to the surface ectoderm and otic vesicle; however, the otic edge tissue was not treated with dispase due to its small size and susceptibility to damage. Embryo dissections were performed using electrolytically sharpened tungsten needles, which were frequently sterilised by passing through an open flame. Embryos were pinned to a Sylgard-coated dish using dissection pins to ensure stability during tissue isolation. The otic edge was dissected first. A tungsten needle was used to perforate the area around the otic pore from the dorsal side, followed by perpendicular perforations to isolate the intact otic pore region completely. The dissected tissue was immediately transferred to a sterile 1.5 ml tube containing 1 ml of TRI reagent (Invitrogen) for RNA preservation. Next, the otic vesicle (excluding the edge) was dissected by gently nudging the structure without damaging the surrounding tissue. The isolated otic vesicle was incubated in a droplet of dispase solution for 10 minutes in a sterile 35 mm dish to remove residual mesenchymal cells. Following digestion, the tissue was transferred to a 1.5 mL tube containing 1 mL of TRI reagent. The surface ectoderm was then dissected using a single tungsten needle to gently displace the tissue, followed by using two needles in a scissor-like manner to excise a small region of epithelium. The dissected tissue was incubated in dispase solution for 10 minutes and then transferred into a 1.5 mL tube containing TRI reagent. For each RNA sample, eight tissue pieces were pooled into a single tube and stored at –20°C until RNA extraction. Total RNA was isolated using the TRI reagent protocol, and the quality and concentration of RNA were assessed using 1 µL of the sample for each of the following: Bioanalyzer (Agilent) for RNA integrity and Qubit fluorometer (Invitrogen) for concentration measurement. RNA samples were resuspended in RNase-free water and stored at –80°C until cDNA library preparation. CRISPR-Cas9 Construct The control-sgRNA sequence 5-GCACTGCTACGATCTACACC-3 was adopted as previously used elsewhere and cloned into BsmB1-flanked cloning region of pcU6.1 sgRNA (Addgene: 92395) ( Gandhi et al., 2017 ). The guide sequences for Grhl2 and SP8 were designed using CRISPOR webtool and were cloned into pcU6.1 sgRNA ( Concordet and Haeussler, 2018 ). The Grhl2-sgRNA used was 5-TGGCAGCTCCACGCCAATTACGG-3 and SP8-sgRNA was 5-CACGCAGTCACGTCGGCGAGCGG-3. CRISPR-Cas9 Electroporation Fertilized Kaveri chicken eggs were incubated for 50 hours to obtain Hamburger-Hamilton stage 15 (HH15) embryos ( Fig. 2 .2). DNA injection and electroporation were performed directly into the developing otic vesicle. Download figure Open in new tab Fig. 1 Three cell populations in otic vesicle closure. A : Scanning Electron Micrographs of bisected OV across stages from (i) HH14, (ii) HH15, (iii) HH16 (iv) HH17 and (v) HH17+ The pseudostratified otic vesicle (OV) cells are shaded in orange and marked by an asterisk. The cuboidal surface ectoderm (SE) cells are shaded in green and marked by arrowheads. The round otic edge (OE) cells are shaded in blue and are marked by arrows. B: Marking the shape of cells at different stages of epithelial fusion in OV sections. The cells are colour-coded according to their sphericity, with blue marking a lower sphericity and red marking a higher sphericity. (i) When the edges are apart, (ii) When the edges are just meeting, (iii) When the edges are fusing and (iv) When OV closure is complete and has segregated from the SE. C: One-way ANOVA of sphericity of cells. D: Population of round cells in the otic edge across fusion stages. 1 &2: HH15-HH15+, when edges are apart. 3 &4: HH16 - HH16+ when edges are pushed closer and fusion is beginning. 5 &6: HH17-17+ when fusion has occurred and remodelling has started. 7: HH18, when the fusion is over and otic vesicle has segregated from the surface ectoderm. Scale bars-10µm. Download figure Open in new tab Fig. 2: Expression profile of adhesion and polarity proteins at different stages of epithelial fusion. For A, C, and E: The first panel is a section where the edges are apart, the second panel is when fusion and remodelling are happening, the third is a higher magnification of the second panel, and the fourth panel is when fusion and segregation are complete. Red asterisk marks the round edge cells. For B, D and F: The graphs depict the coefficient of variation (CV) of fluorescence intensity of different proteins in three cell populations; cuboidal surface ectoderm SE, pseudostratified otic vesicle cells PS and round otic epithelial edge cells OE. Significance was tested using one way-ANOVA. A: Tight junction protein ZO1. B: CV of ZO1. C: Apical polarity marker, Rac1 D: CV of Rac1. E: Adherens junction marker, Cdh1. F: CV of Cdh1. G: HH16 staged embryos electroporated with constitutively active eGFP and stained for laminin. Red asterisk marks a round cell expressing GFP and not attached to the basal lamina. H: Whole mount imaging of the closing otic vesicle, stained for Rac1, to visualize filopodia, as marked by a red asterisk. Scale bar for sections and whole mount-10µm. Scale bar for magnified insets-5 µm. Microcapillary glass needles were loaded with the DNA injection mix consisting of 5 µg/µL plasmid DNA, 30% sucrose, and 0.05% Fast Green to visualize the injection volume. The vitelline membrane over the embryo was carefully removed, and the DNA mix was gently injected into the lumen of the otic vesicle using a micromanipulator, avoiding physical contact with the embryonic tissue. Electroporation was carried out immediately after injection using platinum electrodes on either side of the otic vesicle (Supplementary Fig. 2). Five square-wave pulses (14 V, 50 ms duration, 100 ms intervals) were applied using a square-pulse electroporator. Following electroporation, embryos were cultured for 8 hours in albumin agar plates at 37°C before fixation or further processing. RESULTS Cells in the otic edge are distinctly shaped To characterise the cellular features of the otic vesicle (OV) during closure we first used SEM to observe this process. En face views of the closing OV showed fusion starting from around HH16 and covering a period of around 8 hours until HH17+, when the OV has just closed. To understand its cellular features, we imaged slices through the OV as it closes ( Fig 1A ). Surface ectodermal (SE) cells showed a cuboidal morphology while otic vesicle cells were pseudostratified. At the border of these two domains, we detected cells that showed a more rounded phenotype and, in some cases, were detached from the basal lamina or below the apex of the epithelium. To further understand and characterise the cell shape found at the otic edge, we used phalloidin to stain for cortical actin in thick sections of the closing otic vesicle. This allowed us to segment individual cells and assess their shape ( Fig 1B ). Cells with a sphericity index close to 1 were identified in the otic edge cells ( Fig 1C ). To ask when these round cells first appeared, we assessed the different stages of otic vesicle closure. The rounded otic edge (OE) cells first appeared at HH16, 8 hours before vesicle closure, and their numbers increased until vesicle fusion and the formation of the otocyst. By HH18, 2hrs after fusion, OE cells were not detected ( Fig 1D ). Otic Edge Cells show reduced apicobasal polarity To understand the rounded edge cells further, we investigated the localisation of markers of epithelial polarity during closure. To quantify polarised expression, we developed a measure of localisation along the periphery of cells, the coefficient of variation (CV). Here, values close to zero show invariant intensity across the periphery, whereas values close to one show high variation. Zonula Occludens-1 (ZO-1) is a tight junction component found at the extreme apical end of the lateral domain ( Gumbiner, 1987 ; Stevenson et al., 1986 ). ZO-1 is restricted to the apical regions in both SE and OV cells ( Fig 2A ), with an average CV of around 0.68±0.66 and 0.72±0.15 respectively. In edge cells, however, ZO-1 is down-regulated, with a CV of 0.26±0.067, indicative of unpolarized expression ( Fig 2B ). The small GTPase Rac1 is known to regulate actin assembly and is localised to epithelia ( Eaton et al., 1995 ; Ridley et al., 1992 ). Rac1 is apically localised in both SE and OV cells with a CV of 0.65±0.19 and 0.91±0.39, respectively ( Fig 2C & 2D ). Previous studies had identified a role for Rac1 in filopodia formation in neural border cells prior to neural tube closure ( Rolo et al., 2016 ). Consistent with this, we observe Rac1-positive filopodia in otic edge cells as the edges are brought together ( Fig 2H ). As the edges fuse, Rac1 is down-regulated in otic edge cells. Moreover, apical polarity is diminished and shows a CV of 0.36±0.1 in OE cells ( Fig 2D ). Cdh1 (E-cad) is a component of adherens junctions in epithelia ( Takeichi, 2014 ; Yoshida-Noro et al., 1984 ). We find that it is localised to the lateral domain of SE and OV cells with a CV of 0.62±0.15 and 0.58±0.2, respectively ( Fig 2E & 2F ). A population of cells in the edge show a loss of this lateral restriction, with a CV of 0.43±0.125. The radialisation of Cdh1 expression led us to probe further the epithelial properties of otic edge cells. Epithelia are characterised by their association with the basal lamina ( Matlin et al., 2017 ). We thus asked if round cells in the otic edge were in contact with the basal lamina. We electroporated the otic edge with a low concentration of a plasmid encoding GFP. This filled some cells, enabling us to assess their shape and position. Using immunostaining for laminin, we found that a proportion of round cells do not contact the basal lamina. In contrast, SE and OV cells remain in contact with the basal lamina ( Fig 2G ). Fusion generates interstitial cells between the surface ectoderm and the otic vesicle During the final stages of otic vesicle closure, the apposed epithelia establish contact and fuse to close the otic pore. Importantly, the two epithelia segregate so that the otic vesicle pinches off the surface ectoderm. Soon after fusion and when remodelling has just begun, we noted a population of interstitial cells between the SE and OV ( Fig 3A-C ). This population is transient, and no longer detectable after HH17+. Download figure Open in new tab Fig. 3: Interstitial cells aid in fusion and segregation. A: SEM of HH17 otic vesicle, the red asterisk shows the site of fusion. B: SEM of HH17+ otic vesicle, the red asterisk shows a population of mesenchymal cells at the site of fusion and remodelling. C: Round interstitial cells marked in HH17+ otic vesicle section. D: Cleaved caspase3 marks cells undergoing apoptosis. The red asterisk marks caspase3 positive cells in the edge and at the site of fusion. E: Laminin, a basal membrane marker. The red asterisk marks diffused expression of laminin at the site of fusion. F: Interstitial cells show a diffused expression of Cdh1 at the site of remodelling, as marked by diffused laminin expression (red asterisk). G: Interstitial cells stained for Rac1 show a polarized expression and seem to be secreting laminin (red asterisk). H: Otic vesicle section stained for Pax2, (i)during fusion and (ii) during remodelling. Scale bar: sections-10µm. To ask if the interstitial cells underwent cell death, we investigated the activity of cleaved caspase 3, a marker of apoptosis ( Porter and Janicke, 1999 ). Staining is observed at the junction of the surface ectoderm and otic vesicle from HH16 ( Fig 3D i,ii). Later, we observe apoptosis in only some of the interstitial cells ( Fig 3D iii-v). This may suggest that interstitial cells have re-integrated into either the SE or OV. The interstitial cells must re-establish their polarity after segregation. To investigate this process, we revisited the expression patterns of key polarity determinants - Rac1, Cdh1, and the ECM component laminin. At early stages of otic vesicle development, laminin is expressed in a contiguous, fibrillar pattern ( Fig 3E i,ii). As fusion progresses, this expression becomes more diffuse ( Fig 3E iii,iv), before returning to a fibrillar organisation by the end of segregation ( Fig 3E v). Notably, some interstitial cells secrete laminin and exhibit polarised Rac1 expression, suggesting a role in orienting neighbouring cells ( Fig 3G iv,v). In contrast, Cdh1 maintains a diffuse expression pattern throughout segregation ( Fig 3F iv,v), which may be crucial for allowing cells to reform new junctions and commit to specific fates. To understand whether edge cells can switch lineage, we examined the otic fate marker Pax2 ( Fig 3H ) ( Christophorou et al., 2010 ; Freter et al., 2012 ; Hidalgo-Sanchez et al., 2000 ; Hutson et al., 1999 ). We found that it is transiently expressed during fusion in the edge cells during fusion, but is completely absent from the site following segregation, consistent with previous observations in mouse models ( Burton et al., 2004 ). These findings suggest that a coordinated interplay between polarity proteins, ECM remodelling, and junctional complexes drives cell sorting during otic vesicle closure. Edge cells have a distinct transcriptomic profile Altered junctional and polarity proteins appear to be critical for the neighbor exchange observed during epithelial fusion and the partial EMT phenotype, as reported in other systems ( Aiello et al., 2018 ; Arnoux et al., 2008 ; Bahri et al., 2010 ; Huang et al., 2012 ; Shaw and Martin, 2016 ). This partial EMT state is thought to be maintained by a finely balanced expression of transcription factors promoting either epithelial maintenance or mesenchymal transition ( Saitoh, 2023 ). Among these are Grhl2/3, Zeb2, and Snail2 ( Nieto et al., 2016 ). We thus investigated their expression during otic vesicle fusion. While Grhl2 and Zeb2 transcripts were detected at the site of fusion in HH17 embryos, Snail2 expression was restricted to the adjacent mesenchyme ( Fig. 4A ). Interestingly, Grhl2 was localised to the Pax2-negative region of the otic vesicle ( Fig. 3H iii), and its expression persisted at the otic pore edges from HH14 through the completion of fusion ( Fig 4I ). In contrast, Zeb2 expression was more transient and spatially restricted to the edge during the fusion stage ( Fig 4A iii). The co-expression of epithelial-stabilising Grhl2 and EMT-inducing Zeb2 suggests that cells at the fusion site may exist in a hybrid EMT state. Download figure Open in new tab Fig. 4. Transcriptomic state of the edge cells. A: Whole mount and section of HH17 otic vesicle WhISH for the genes Grhl2 , Snail2 and Zeb2 . B: Side view of HH17 chick embryo showing the three types of tissue taken for bulk-RNA sequencing. C: The Multi-Dimensional Scaling (MDS) plot shows quadruplicates of the same tissue clustering together, and the three tissue types segregated well away from each other. This confirms the quality of the sample collection. D: Heatmap of top 30 DEG in OE with respect to OV and SE respectively E-N: WhISH sections of HH17 embryos of some gene targets identified from RNA-seq data for three stages as mentioned in Fig.2 . The gene names are mentioned in the respective figure panel. Scale bar: sections-10µm. To explore this further, we performed bulk mRNA sequencing of microdissected tissues from HH17 embryos, isolating the otic epithelium (OE), otic vesicle (OV), and surface ectoderm (SE) ( Fig 4B ). PCA and differential gene expression (DGE) analyses confirmed that the OE cells represent a transcriptionally distinct population from either OV or SE ( Fig 4C ). Reactome analysis of DGE between the edge region (OE) and surrounding tissues (SE and OV) identified several enriched pathways, including gastrulation, WNT signalling, ECM remodelling, and organogenesis (Supplementary Fig. 1A, Supplementary Fig. 1B), as summarised in the heatmaps ( Fig 4D i, ii). Some targets were then validated by whole-mount in situ hybridisation (WhISH) ( Fig. 4E–N ), with expression domains predominantly confined to the OE, although occasionally extending into SE or OV. Dach1 is a known inhibitor of EMT via Snail1 suppression in breast cancer ( Zhao et al., 2015 ) and Six1 suppression in liver cancer ( Cheng et al., 2018 ). Its expression was restricted to the OE and SE. Consistent with this reciprocal relationship of Dach1 and Six1, our DGE data revealed upregulation of Dach1 and downregulation of Six1 in OE versus OV ( Fig. 4F ). This corresponds with findings in later inner ear development, where knockdown of Dach1 in cochlear epithelium induces EMT and dysplasia ( Miwa et al., 2019 ). Several transcription factors implicated in developmental patterning and EMT were spatially restricted within the otic region. Dlx5/6 are known regulators of vestibular fate in OV and in play a role in craniofacial patterning ( Depew et al., 2002 ; Robledo and Lufkin, 2006 ). As previously observed ( Groves and Bronner-Fraser, 2000 ), Dlx5 showed dorsal OV expression ( Fig. 4G ). Msx1 is involved in known to drive apoptosis during limb development ( Lallemand et al., 2009 ) and EMT during palate morphogenesis ( Bendall and Abate-Shen, 2000 ; Levi et al., 2006 ). It is restricted to the OE ( Fig. 4K ), a region also undergoing apoptosis ( Fig. 3H iii). Genes involved in signalling cross-talk and cell-ECM interaction were also differentially expressed at the fusion site. Epha4 mediates cell migration via TGFβ signalling in cancer ( Hachim et al., 2017 ) had a transient, restricted expression in the OE ( Fig. 4H ). Itgb3 is a key integrin mediating ECM interaction and known for its role in metastasis ( Kovacheva et al., 2021 ). It was expressed in the OE and the dorsal otic vesicle ( Fig. 4J ). Bambi, a negative regulator of TGFβ and BMP signalling induced by WNT signalling ( Lin et al., 2008 ; Sekiya et al., 2004 ) was also transiently expressed at the OE ( Fig. 4E ). Wnt3a is known to induce EMT ( Qi et al., 2014 ), showed persistent expression in the OE even after fusion. This suggests additional roles in inner ear patterning, likely in dorsal-ventral patterning of the otocyst ( Riccomagno et al., 2005 )( Fig. 4N ). The transcription factor SP8 acts downstream of Wnt3, Fgf10, and Bmpr1A during mouse development ( Bell et al., 2003 ). It is also implicated in inner ear development in Xenopus ( Chung et al., 2014 ) and EMT in hepatoblastoma ( Wagner et al., 2020 ). It showed restricted OE expression ( Fig. 4M ). Rac3, a regulator of actin cytoskeleton dynamics and invasive behaviour in breast cancer ( Gest et al., 2013 ), was expressed at the fusion site, further supporting a role in cell remodelling ( Fig.4L ). Overall, we observed a spatially restricted and balanced expression of transcription factors and signalling molecules that either promote or inhibit WNT, TGFβ, and BMP signalling. This balance likely sustains the partial EMT state of OE cells. Notably, TGFβ signalling has been implicated in epithelial fusion in other developmental contexts ( Iwata et al., 2011 ; Knickmeyer et al., 2018 ; Nakajima et al., 2000 ). While many of these genes are well-studied in cancer and adult tissues, their coordinated expression here suggests a developmental program that mimics, but does not fully execute, EMT during embryonic epithelial fusion. Molecular perturbation of epithelial fusion Based on their expression patterns and putative roles in epithelial dynamics and EMT regulation, Grhl2 and SP8 were selected for functional perturbation. Grhl2 is a key transcription factor that regulates epithelial integrity by controlling the expression of Cdh1 and Claudin4 ( Werth et al., 2010 ). During embryogenesis, its role appears context-dependent; while some studies suggest it suppresses EMT in cancer cell lines, others highlight a more ambiguous involvement ( Cieply et al., 2012 ; Jolly et al., 2016 ; Wang et al., 2023 ; Werner et al., 2013 ). Mouse models with Grhl2 knockouts exhibit neural tube (NT) and organ defects, varying with the specific allele ( Pyrgaki et al., 2011 ). In zebrafish, Grhl2b mutants show inner ear phenotypes resembling human non-syndromic deafness DFNA28 ( Han et al., 2011 ). Neural tube defects in mouse Grhl2 knockouts have been linked to Cdh1-dependent mechanisms ( Nikolopoulou et al., 2019 ). We detected Grhl2 expression at the otic edge prior to fusion, and this expression was downregulated after segregation in the OE. SP8, a transcription factor involved in WNT signalling and limb and NT development, has also been implicated in inner ear formation ( Chung et al., 2014 ). To investigate the functional roles of Grhl2 and Sp8, we performed CRISPR-Cas9-mediated mosaic knockdowns (KDs) via electroporation. GFP-positive cells indicated successful uptake of CRISPR constructs. In embryos electroporated with control gRNAs, CDH1 expression appeared similar to wild-type patterns ( Fig 5A, B ). However, embryos electroporated with Grhl2-sgRNA displayed misregulated CDH1 expression at the fusion site. Aberrant CDH1 accumulation was detected within OE cells, regardless of GFP positivity ( Fig 5C ). Moreover, segregation between the OV and SE was incomplete ( Fig 5C iv,v). In contrast, SE cells that were GFP-positive exhibited complete loss of CDH1 ( Fig 5C iv, v), reminiscent of the switch to CDH2 seen in Grhl2 KO mice ( Nikolopoulou et al., 2019 ). Download figure Open in new tab Fig. 5: Genetic perturbation of epithelial fusion. Different stages of epithelial fusion are depicted here: (i) Edges are close, (ii) Edges establish contact, (iii)Edges are fusing, (iv) Site of fusion is remodeling, (v) Fusion site is segregating. Sections are stained with DAPI (cyan), CDH1 (magenta) and GFP (green) in A: WT embryos. B: Embryos electroporated with control-sgRNA C: Embryos electroporated with Grhl2-sgRNA. The yellow asterisk marks the accumulation of Cdh1 in the edge cells and the red asterisk marks the loss of Cdh1 in the surface ectoderm cells. D: Embryos electroporated with Sp8-sgRNA. The yellow asterisk marks the accumulation of Cdh1 in the edge cells and the red asterisk marks the loss of Cdh1 in the surface ectoderm cells. E: Sphericity of OE, PS and SE cells in electroporated embryos. Significance tested by one-way ANOVA. Scale bar: sections-10µm. Sp8 mosaic knockdowns also resulted in ectopic CDH1 accumulation at the OE and loss of expression in the SE ( Fig 5D ). Furthermore, the sphericity of cells in the OE and SE was significantly reduced compared to control embryos ( Fig 5E ), suggesting altered mechanical properties or junctional remodelling. Although the CRISPR knockdowns were mosaic, the phenotypes observed were tissue-wide. This broader effect could be due to the mechanical and signalling integration properties of CDH1, which transmits cortical actin tension between neighbouring cells ( Lecuit and Yap, 2015 ). Interestingly, the pseudostratified morphology of cells mediating fusion remained largely unperturbed. This is consistent with the absence of Grhl2 and Sp8 expression in the OV, suggesting their primary function is restricted to the OE and SE. These findings support a model in which Grhl2 and Sp8 contribute to cell shape regulation, CDH1 localisation, and ECM remodelling, all of which are essential for proper epithelial fusion and segregation. DISCUSSION Epithelial fusion is a critical event in embryogenesis, with failures resulting in a range of developmental disorders such as neural tube defects, cleft palate, and body wall closure defects ( Copp et al., 2015 ; Ray and Niswander, 2012 ). While significant work has focused on genetic contributions to fusion events, particularly in single-gene mutation models, these do not fully capture the complexity of fusion failure in human birth defects. Our study takes a complementary approach by investigating the dynamic cell behaviours, junctional remodelling, and ECM interactions that occur during otic vesicle fusion, using the chick embryo as a model system. One of the key findings from our study is the identification of a transient population of round cells located at the boundary between the surface ectoderm (SE) and OV. These cells display distinct morphological and molecular features that evolve across fusion stages. Immunostaining for CDH1, ZO1, and RAC1 revealed dynamic changes in their expression ( Fig. 2 ), indicating a process of junctional remodeling. Epithelial remodeling in this context likely involves neighbor exchange and transient loss of stable cell-cell contacts, mediated through endocytic trafficking of adhesion proteins. Previous studies have shown junction remodelling is mediated by intracellular trafficking ( Levayer and Lecuit, 2013 ; Roeth et al., 2009 ; Sigismund et al., 2021 ), often in response to mechano-chemical stimulus; for instance, in the Drosophila wing, p120-catenin is released under tension, promoting CDH1 endocytosis ( Iyer et al., 2019 ). Similar tension-mediated mechanisms may operate during otic fusion, where tissue stress, arising from the buckling of the otic edge may enhance CDH1 turnover. This process could be further regulated by Rac-family GTPases, which link CDH1 endocytosis with actin cytoskeleton remodelling ( Baum and Georgiou, 2011 ) and is consistent with the observed expression of Rac1 and the specific expression of Rac3 in fusing cells. The final phase of fusion is marked by ECM remodelling, particularly changes in laminin distribution ( Fig. 3 E-G). Initially fibrillar and continuous, laminin becomes diffuse as fusion proceeds. It is reassembled into distinct basement membranes as the epithelia segregate. This remodelling resembles epiboly during gastrulation, where tissue thinning occurs through radial intercalation of deeper cells into superficial layers ( Keller and Hardin, 1987 ; Trinkaus and Lentz, 1967 ). A similar unpolarized intercalation is also likely necessary for the round OE cells to integrate into either the OV or SE. Such intercalation may be driven by junctional remodeling and cell sorting mechanisms. Two primary hypotheses explain sorting at adherens junctions: differences in surface adhesion ( Foty and Steinberg, 2013 ) and differential cell stiffness regulated by cadherins ( Krens and Heisenberg, 2011 ). The diffuse expression of junctional proteins in round cells may regulate stiffness and allow a more fluid and labile cell state, allowing junctions with either SE or OV neighbours to readily form. Rac1 plays a central role in coordinating polarity and ECM assembly. Its expression, along with F-actin, was detected in cells secreting laminin ( Fig 3G ), suggesting it orchestrates actin-mediated basal matrix organisation ( O’Brien et al., 2001 ). In contrast, scanning electron microscopy revealed a subset of mesenchyme-like cells near the OE, which may participate in basal lamina breakdown ( Fig 3B ), akin to periocular mesenchyme involvement in optic fissure closure ( Gestri et al., 2018 ). The requirement for such cells in OV closure remains to be tested in mesenchyme-free cultures. Notably, Rac1 is expressed in premigratory cell populations ( Kee et al., 2007 ). Its downregulation by the proteoglycan, Syndecan4 is necessary for initiating migration ( Matthews et al., 2008 ). The close homologue Rac3 is specifically expressed in the otic edge ( Fig 4L ), further suggesting a role for Rac family members in regulating the cell state transitions required for fusion. Transcriptomic analysis of the OE region revealed differential expression of genes involved in ECM remodelling, gastrulation, and Wnt signalling (Supplementary Fig 1A & B), processes also enriched during optic fissure fusion ( Hardy et al., 2019 ). However, interpreting chick bulk RNA-seq data remains challenging due to limited genome annotation and GO mapping resources. Functional dissection of candidate genes required targeted KD, allowing us to distinguish roles in differentiation versus epithelial fusion. We focused on two transcription factors with opposing roles: Grhl2, an epithelial maintenance factor, and Sp8, an EMT inducer. Grhl2 zebrafish mutants exhibit enlarged otic vesicles and malformed semicircular canals, structures arising from the dorsal OV where fusion occurs ( Han et al., 2011 ). Sp8 mutants in Xenopus display broad defects in the auditory and vestibular systems ( Chung et al., 2014 ). Mosaic knockdown of either gene led to CDH1 accumulation in cells at the fusion interface and a loss of round cells, replaced by pseudostratified or squamous morphologies ( Fig. 5 ). These data suggest that proper junction turnover and partial EMT are disrupted under knockdown conditions, resulting in impaired force transmission and cell shape change. Accumulation of CDH1 likely increases cortical stiffness, reducing the ability of cells to undergo necessary morphological transitions. This is supported by prior studies showing that Grhl2 overexpression stiffens epithelia and impedes morphogenetic movements ( Nikolopoulou et al., 2019 ). Interestingly, despite the mosaic nature of our knockdown, fusion was disrupted tissue-wide. This non-cell-autonomous effect may be mediated through either chemical signalling or mechanical coupling of junctional tension across epithelial sheets. Importantly, the epithelia did not segregate. This could be attributed to a lack of a fluidic state of the round cells, that we postulate are required for proper epithelial fusion. Proper localization of junctions is important to accommodate cell shape changes without compromising their function ( Levayer and Lecuit, 2013 ; Roeth et al., 2009 ). Thus, it is possible that Grhl2 and Sp8 are key regulators of the partial EM phenotype of OE round cells, and by regulating CDH1 localisation, they regulate epithelial fusion. Future studies should explore the additional candidates identified in our transcriptome, such as Mlc9 , Exoc4 , Rac3 , and β -catenin , to unravel the broader gene regulatory network underlying otic fusion. Additionally, live imaging and biophysical measurements of junctional tension could provide deeper insight into the emergent behaviours driving epithelial fusion and segregation during inner ear development. Supplementary Fig. 1: Pathway over-representation using reactome analysis A. OE vs OV comparisons Top pathways overrepresented in the differential gene expression analysis. A: Colour key for the reactome. The yellow to blue colour indicates the pathways with high to low over representation. B: Muscle contraction C: WNT signalling D: Gastrulation and Kidney development. E: Nervous system development and F: ECM organisation. B: OE vs SE comparisons Top pathways over represented in the differential gene expression analysis. A: gastrulation B: Early embryonic development C: Nervous system development D: Transcription E: ECM organisation F: Cell junction organisation G: Muscle contraction H: Colour key for the reactome. The yellow to blue colour indicates the pathways with high to low overrepresentation. Supplementary Fig.2: Schematic for electroporation. After electroporation, the embryos were kept on thin-albumin agar plates and incubated at 37°C. ACKNOWLEDGEMENTS This work was supported by the Department of Atomic Energy, Government of India, Project Identification No. RTI 4006, and grants from ANRF-SERB (CRG/2018/001235), the Royal National Institute for Deaf People (IPG programme) and TIFR Infosys-Leading Edge Grant. We acknowledge the support of the Electron Microscopy Facility and Central Imaging Facility at NCBS. We thank Rolf Eriksson for his valuable inputs in bulk mRNA sequencing. We also thank Dr. Awadesh of the NGS facility in NCBS, for performing the mRNA sequencing. Shivangi Pandey and Nidhi Parikh helped in the standardisation of the in-situ hybridisation protocol. Raman Kaushik helped in the design of the CRISPR-gRNA. We thank CSIR for JRF-SRF Fellowship to VNT. 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