Mechanical forces stimulate Golgi export

Mechanical forces stimulate Golgi export | 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 Mechanical forces stimulate Golgi export View ORCID Profile Chandini Bhaskar Naidu , View ORCID Profile Javier Vera Lillo , Sabine Bardin , View ORCID Profile Anabel-Lise Le Roux , View ORCID Profile Nicolas Mateos , View ORCID Profile Jessica Angulo-Capel , View ORCID Profile Adam Wolowczyk , View ORCID Profile Pere Roca-Cusachs , View ORCID Profile Maria F. Garcia-Parajo , View ORCID Profile Franck Perez , View ORCID Profile Bruno Goud , View ORCID Profile Jean-Baptiste Manneville , View ORCID Profile Stéphanie Miserey , View ORCID Profile Felix Campelo doi: https://doi.org/10.1101/2025.09.02.673725 Chandini Bhaskar Naidu 1 Intracellular Transport: Engineering and Mechanisms Laboratory, Institut Curie, PSL Research University, Sorbonne Université, Centre National de la Recherche Scientifique , UMR 144, Paris, France 2 Laboratoire Matières et Systèmes Complexes, UMR 7057, CNRS and Université Paris Cité , CNRS, UMR7057, 10 rue Alice Domon et Léonie Duquet, F-75013, Paris cedex 13, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Chandini Bhaskar Naidu Javier Vera Lillo 3 ICFO-Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Javier Vera Lillo For correspondence: javier.vera{at}icfo.eu Jean-Baptiste.Manneville{at}u-paris.fr Stephanie.Miserey{at}curie.fr felix.campelo{at}upf.edu Sabine Bardin 1 Intracellular Transport: Engineering and Mechanisms Laboratory, Institut Curie, PSL Research University, Sorbonne Université, Centre National de la Recherche Scientifique , UMR 144, Paris, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site Anabel-Lise Le Roux 4 Institute for Bioengineering of Catalonia (IBEC), The Barcelona Institute of Technology (BIST) , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Anabel-Lise Le Roux Nicolas Mateos 3 ICFO-Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Nicolas Mateos Jessica Angulo-Capel 3 ICFO-Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jessica Angulo-Capel Adam Wolowczyk 3 ICFO-Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Adam Wolowczyk Pere Roca-Cusachs 4 Institute for Bioengineering of Catalonia (IBEC), The Barcelona Institute of Technology (BIST) , Barcelona, Spain 5 University of Barcelona , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Pere Roca-Cusachs Maria F. Garcia-Parajo 3 ICFO-Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology , Barcelona, Spain 6 Institució Catalana de Recerca i Estudis Avançats (ICREA) , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Maria F. Garcia-Parajo Franck Perez 1 Intracellular Transport: Engineering and Mechanisms Laboratory, Institut Curie, PSL Research University, Sorbonne Université, Centre National de la Recherche Scientifique , UMR 144, Paris, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Franck Perez Bruno Goud 1 Intracellular Transport: Engineering and Mechanisms Laboratory, Institut Curie, PSL Research University, Sorbonne Université, Centre National de la Recherche Scientifique , UMR 144, Paris, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Bruno Goud Jean-Baptiste Manneville 2 Laboratoire Matières et Systèmes Complexes, UMR 7057, CNRS and Université Paris Cité , CNRS, UMR7057, 10 rue Alice Domon et Léonie Duquet, F-75013, Paris cedex 13, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jean-Baptiste Manneville For correspondence: javier.vera{at}icfo.eu Jean-Baptiste.Manneville{at}u-paris.fr Stephanie.Miserey{at}curie.fr felix.campelo{at}upf.edu Stéphanie Miserey 1 Intracellular Transport: Engineering and Mechanisms Laboratory, Institut Curie, PSL Research University, Sorbonne Université, Centre National de la Recherche Scientifique , UMR 144, Paris, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Stéphanie Miserey For correspondence: javier.vera{at}icfo.eu Jean-Baptiste.Manneville{at}u-paris.fr Stephanie.Miserey{at}curie.fr felix.campelo{at}upf.edu Felix Campelo 3 ICFO-Institut de Ciencies Fotoniques, The Barcelona Institute of Science and Technology , Barcelona, Spain 7 Department of Medicine and Life Sciences (MELIS), Universitat Pompeu Fabra (UPF) , Barcelona, Spain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Felix Campelo For correspondence: javier.vera{at}icfo.eu Jean-Baptiste.Manneville{at}u-paris.fr Stephanie.Miserey{at}curie.fr felix.campelo{at}upf.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF ABSTRACT Human cells face a wide range of external mechanical stimuli that vary with cell type, state, and pathological conditions. The rapidly growing field of mechanobiology investigates how cells sense and respond to these forces. While most work has focused on focal adhesions (FAs), plasma membrane, and nucleus as primary mechanosensors, how reciprocal inside-out signals adapt intracellular organelles to extracellular mechanics has remained largely unexplored. Here, we show that extracellular mechanical signals influence the secretory function of the Golgi apparatus. By subjecting adherent cells to various mechanical challenges –cell spreading on surfaces coated with different ligands, altering substrate stiffness, or applying equibiaxial strains to the cells–, we reveal that extracellular forces modulate Golgi transport carrier biogenesis, thereby regulating exocytosis. Together with modulation of Golgi membrane tension, we identify molecular determinants of the underlying mechanotransduction pathway, including microtubule acetylation, diacylglycerol (DAG) production, and protein kinase D (PKD) activity. These findings uncover a bidirectional mechanotransduction axis in which extracellular mechanics tune Golgi secretory output, providing a framework for investigating organelle-based mechanoadaptation in physiology and disease, particularly in cancer and fibrosis where secretion is critical. INTRODUCTION Exocytosis enables cells to deliver molecules to their surface or secrete them into the extracellular space, thereby maintaining plasma membrane (PM) and extracellular matrix (ECM) homeostasis and organization and facilitating interactions with the environment. Newly synthesized secretory and membrane proteins are transported along intracellular routes from the endoplasmic reticulum (ER) through the Golgi complex and to the trans -Golgi network (TGN). At the TGN, cargos are sorted and packaged into transport carriers, which are rapidly directed to the PM along microtubule (MT) tracks. Upon arrival at the cell surface, transmembrane proteins are incorporated into the PM, while soluble cargos are released into the extracellular space through exocytosis. Similar to the highly heterogeneous organization of the PM, exocytic events are spatially regulated, ensuring that secretion is precisely coordinated with cellular architecture and function. Previous studies have demonstrated that many exocytic events occur preferentially near focal adhesions (FAs) ( Stehbens et al., 2014 ; Huet-Calderwood et al., 2022 ; Eisler et al., 2018 ; Fourriere et al., 2019 ; Lachuer et al., 2023 ). In particular, by combining the Retention Using Selective Hooks (RUSH) assay ( Boncompain et al., 2012 ) –which synchronizes anterograde cargo transport– with a Selective Protein Immobilization (SPI) assay, we previously mapped the precise spatial organization of cargo arrival sites at the PM. Interestingly, our findings revealed that exocytosis does not occur randomly across the cell surface but rather at hotspots juxtaposed with FAs ( Fourriere et al., 2019 ). Among the carriers involved in these targeted exocytic events are Golgi-derived, RAB6-positive carriers ( Fourriere et al., 2019 ). RAB6 mediates multiple different Golgi-to-PM export routes, including the trafficking of CARTS (carriers of the TGN to the cell surface), a class of RAB6- and protein kinase D (PKD)-dependent Golgi-to-PM transport carriers ( Wakana et al., 2012 ). FAs are dynamic, force-sensitive molecular platforms localized at the cell membrane. The core components of FAs are integrins, heterodimeric transmembrane receptors that connect the ECM with cytoplasmic components, enabling the transmission of extracellular signals into the cell. These outside-in signals control essential processes such as cell migration, tissue morphogenesis, and PM homeostasis ( Kechagia et al., 2019 ; Kanchanawong and Calderwood, 2023 ) . While substantial progress has been made in understanding how FAs sense and transduce extracellular mechanical stimuli, it is far less understood whether and how intracellular organelles such as the Golgi reciprocally adapt their activity to these cues and thereby contribute to the overall cellular mechanoresponse. The aforementioned observation that membrane regions proximal to FAs act as hotspots for exocytosis suggests a spatial coordination of membrane trafficking and adhesion sites. Exocytosis delivers integrins, ECM components (such as collagens, fibronectin and fibrillin), and signaling molecules to the PM ( Moreno-Layseca et al., 2019 ), thereby sustaining adhesion dynamics. While MTs have long been recognized to target to FAs ( Schmidt and Stehbens, 2024 ; Krylyshkina et al., 2003 ; Seetharaman and Etienne-Manneville, 2019 ), emerging evidence indicates that exocytosis near FAs is spatially orchestrated by MT guidance, RAB GTPase-dependent Golgi-to-PM trafficking, and, potentially, mechanochemical feedback loops. This spatial regulation may provide a mechanism for localizing membrane expansion, modulating membrane tension, and remodeling the ECM and PM proteome –processes critical for adhesion homeostasis during cell spreading and migration. Understanding these dynamics may also offer insights into how dysregulation of these processes contributes to pathological conditions such as cancer invasion. While several mechanosensitive modules, such as FAs, fibrillar adhesions, and adherens junctions at the PM ( Elosegui-Artola et al., 2018 ; Pascalis and Etienne-Manneville, 2017 ), and the nuclear envelope ( Thorpe and Lee, 2017 ; Amar et al., 2021 ; Jahed and Mofrad, 2019 ) have been well characterized, recent studies indicate that other organelles, such as the ER and lysosomes, can respond to mechanical forces ( Phuyal and Baschieri, 2020 ; Rawal et al., 2025 ; Merta et al., 2025 ; Aceitón et al., 2025 ; Ren et al., 2025 ; Naughton et al., 2025 ). Moreover, mechanical forces applied to the PM can trigger Ca 2+ release from the ER via an actomyosin-dependent mechanism ( Kim et al., 2015 ). In the context of intracellular trafficking, neurite mechanical tension enhances active vesicular transport in neurons ( Ahmed and Saif, 2014 ), while COPII-mediated transport from the ER to the Golgi apparatus is regulated by mechanical strain through interactions between the small G proteins Rac1 and Sar1 ( Phuyal et al., 2022 ), and is sensitive to ER membrane viscosity ( Jiménez-Rojo et al., 2025 ). The Golgi apparatus itself is mechanoresponsive: external forces generated by adhesion to the ECM influence lipid metabolism and Golgi mechanics ( Romani et al., 2019 ), and cell–matrix adhesion controls Golgi organization and secretory output via an Arf1- and AXL-dependent pathway ( Singh et al., 2018 ; Joshi et al., 2025 ). Because the actomyosin cytoskeleton ( Miserey-Lenkei et al., 2010 ; Zilberman et al., 2011 ) and MT networks ( Fourriere et al., 2020 ) are closely associated with Golgi membranes, forces transmitted through the cytoskeleton can directly alter Golgi dynamics ( Miserey-Lenkei et al., 2017 ). In addition, using optical-tweezers based microrheology, we showed that directly applying force to the Golgi apparatus delays actin-mediated membrane fission and thereby perturbs post-Golgi trafficking, indicating that Golgi membranes are mechanosensitive ( Guet et al., 2014 ). In this study, we address the impact of extracellular mechanical forces on post-Golgi carrier biogenesis. Our results show that disruption of FA formation decreases Golgi-derived carrier biogenesis, whereas substrate stiffening and mechanical stretching both enhance Golgi export capabilities. Using Halo-Flipper probes, we further show that extracellular mechanical forces are accompanied by changes in Golgi membrane tension. This is paralleled with an increase in MT acetylation and changes in Golgi diacylglycerol (DAG) content. Collectively, our data identifies the Golgi apparatus as a mechanoresponsive organelle, revealing that its export activity is modulated by extracellular mechanical stimuli and is crucial for efficient cell spreading and mechanoadaptation. RESULTS Disruption of FA formation inhibits Golgi-derived carrier biogenesis In non-polarized 2D cell cultures, we and others have shown that exocytic sites are not randomly distributed across the PM but preferentially localize close to FAs in a RAB6-dependent manner ( Fourriere et al., 2019 ). RAB6 is a general regulator of post-Golgi carrier biogenesis, including CARTS ( Wakana et al., 2012 ). However, whether CARTS exocytosis itself is spatially regulated with respect to FAs has not been directly tested. To address this, we mapped the delivery of the CARTS-specific secretory cargo Pancreatic Adenocarcinoma Upregulated Factor (PAUF). HeLa cells transiently co-expressing paxillin-GFP (FA marker) and FM4-mKate2-PAUF (CARTS cargo) were imaged by live-cell total internal reflection fluorescence (TIRF) microscopy. After exiting the Golgi apparatus, PAUF-positive carriers migrated quasi-directionally through the cytoplasm ( Video 1 ) and docked preferentially near FA-rich regions of the plasma membrane ( Fig. S1A ). At these sites, CARTS often paused for several seconds before a sudden loss of fluorescence, consistent with fusion with the PM ( Fig. S1A , panel (i) ). By contrast, outside FA areas, CARTS were absent or observed only transiently, without apparent pausing, suggesting that these carriers were likely in transit toward FA sites for exocytosis ( Fig. S1A , panel (ii) ). Using a surface protein immobilization (SPI) assay at 40 min upon release, CARTS-delivered secreted PAUF accumulated at the periphery of the cell in proximity of FAs ( Fig. S1B,C ), consistent with previous findings combining SPI and RUSH of different types of cargoes ( Fourriere et al., 2019 ). To test whether integrin-mediated adhesion modulates Golgi-to-PM trafficking, we devised two complementary assays: (i) a combined spreading-secretion assay, and (ii) modulation of FA number by substrate stiffness. First, for the combined spreading-secretion assay ( Fig. 1A ), HeLa cells in suspension were plated on glass coated with fibronectin (FN, which promotes FA formation) or poly-L-lysine (PLL). As previously described ( Fourriere et al., 2019 ), PLL-seeded cells failed to assemble FAs ( Fig. S1D ). Cells were co-transfected with GFP-mem (a general PM marker, see Methods) and FM4-mKate2-PAUF to measure both cell spreading (adhered area) and CARTS biogenesis (number of CARTS at 30 min after cargo release from the ER) ( Fig. 1A , see Methods) by confocal imaging. On FN, cells exhibited larger adhesion areas and produced significantly more CARTS than on PLL ( Fig. 1B, C ). CARTS numbers on FN increased over time, peaking at 4 h post-seeding, whereas PLL-seeded cells showed persistently low CARTS counts even after 24 h ( Fig. 1C ). Likewise, adhesion area on FN steadily expanded to 4 h, while on PLL it remained minimal and unaltered over 24 h ( Fig. 1D ). Similar results were obtained when cells were gently lifted with EDTA prior to the spreading-secretion assay, as compared to trypsinization ( Fig. S1E ). Importantly, CARTS number correlated linearly with adhesion area under all conditions, resulting in a constant number of CARTS per adhesion area ( Fig. 1E,F ). These results suggest that integrin activation and FA formation may co-regulate cell spreading and Golgi export in a commensurate manner. Download figure Open in new tab Figure 1. FA formation impairment leads to decreased CARTS formation. ( A ) Schematic representation of the spreading assay. D/D is D/D solubilizer, which dissolves mKate2-FM4-PAUF aggregates, allowing synchronized cargo release from the ER; CHX is cycloheximide. ( B ) Images represent fixed HeLa cells transfected with GFP-mem and the mKate2-FM4-PAUF, seeded over FN or PLL, and acquired by confocal microscopy after the indicated times. Scale bar, 10 µm. ( C-E ) SuperPlots showing individual cell measurements (small, light-colored symbols; N∼10 per biological replicate) and the mean value for each independent biological replicate (larger, black-outlined circles; n=6 on FN, n=3 on PLL). Each color represents a different experimental replicate. Horizontal black lines and error bars indicate the mean and standard error of the mean (SEM) of the n=3 biological replicates. Plots represent the number of CARTS per cell ( C ), the adhesion surface area per cell ( D ), and the number of CARTS per unit adhesion area ( E ). Repeated-measures 2-way ANOVA tests were performed, and P values were obtained using Tukey’s post-hoc multiple comparison test. ( F ) Correlation plot showing the number of CARTS per cell and the corresponding cell spreading area per each measured individual cell across all tested conditions (see legend, cells seeded on FN only). Normality of both variables was tested using the Shapiro-Wilk test, and the results showed non-normality. Each point represents an individual cell. A nonparametric Spearman correlation analysis was performed, with the correlation coefficient being ρ=0.37 and P value < 0.0001. ( G ) Spinning disk confocal microscopy images of fixed HeLa cells stably expressing RUSH-EGFP-CD59 plated on FN-coated glass coverslips, or on stiff (30 kPa) or soft (2 kPa) FN-coated PAA gels. 4h after plating them, cells were incubated for 45 minutes with biotin before fixation to specifically visualize CD59-positive post-Golgi carriers. ( H ) SuperPlot showing quantification of the number of CD59-positive vesicles in cells seeded on substrates of various stiffness as described in ( G ). Second, we performed a substrate stiffness assay to further manipulate FA assembly, as the number of FAs decreases with decreasing substrate stiffness, resulting in minimal FA formation on soft substrates ( Pelham and Wang, 1997 ; Barber-Pérez et al., 2020 ) . HeLa cells were plated for 4 h on FN-coated polyacrylamide (PAA) gels of defined stiffness (from 2 kPa soft gels to 30 kPa stiff gels) or on glass, and FAs were visualized by immunofluorescence microscopy using vinculin staining ( Fig. S1F ). Cells on soft substrates spread poorly and formed almost no vinculin-positive FAs, whereas cells on stiff substrates (30 kPa PAA gels or glass) displayed robust spreading and abundant FAs ( Fig. S1F ). We then assessed Golgi export using the RUSH assay, which enables synchronized trafficking of selected cargos ( Boncompain et al., 2012 ). HeLa cells stably expressing RUSH-EGFP-CD59 (a RAB6-dependent GPI-anchored protein) were plated on soft (2 kPa PAA gels) and stiff (30 kPa PAA gels or glass) FN-coated substrates for 4 hours. Post-Golgi carriers were quantified at 45 min after biotin addition ( Fig. 1G ). Cells on stiff substrates exhibited ∼150–200 carriers per cell, compared with ∼50 carriers per cell on soft substrates ( Fig. 1H ). Overall, our data support the hypothesis that integrin-mediated outside-in signaling required for efficient cell spreading triggers an adaptive response in the secretory pathway. Mechanical stretch enhances Golgi export capabilities Cell adhesion and spreading on integrin-activating substrates generate mechanical forces via integrin engagement and actomyosin contractility ( Geiger et al., 2009 ). We have found that this correlates with increased Golgi export capacity ( Fig. 1 ). To test whether a distinct external mechanical cue produces similar effects, we applied isotropic mechanical strain on cells using an equibiaxial stretching device ( Fig. 2A ) (see Methods; ( Le Roux et al., 2025 )), a mechanical stimulus that mimics ECM stretching. HeLa cells expressing PAUF-mRFP were seeded on FN-coated stretchable PDMS membranes and allowed to adhere for 1 h at 37°C. To synchronize cargo at the TGN, cells were then shifted to 20°C for 2h in the presence of cycloheximide (CHX). After shifting back to 37°C to resume Golgi export, cells were either left unstretched or subjected to a 15% isotropic strain for 30 min before fixation ( Fig. 2B ). Widefield imaging of the PAUF-mRFP signal revealed that mechanical stretch increased the number of cytoplasmic CARTS by ∼ 40% compared with unstretched controls ( Fig. 2C,D ). Measurements of cell adhesion area showed a ∼12% expansion upon stretch ( Fig. 2E ), indicating some adjustment of adhesion area on PDMS. Consequently, the number of CARTS per adhesion area showed a slight but not statistically significant increase ( Fig. 2F ). These data suggest that although cells quickly remodel their adhesion footprint under stretch, the boost in Golgi-derived carrier production persists. Download figure Open in new tab Figure 2. Mechanical strain application on cells increases their secretory capabilities. ( A ) Schematics of the stretch system used to apply a mechanical strain on cells. ( B ) Schematic representation of the pipeline followed in the stretching experiments. ( C ) Representative widefield images of fixed HeLa cells transfected with PAUF-RFP, seeded over FN-coated PDMS membranes, and subjected to no external forces or 15% equibiaxial stretch for 30 min. Scale bars, 10µm. ( D-F ) SuperPlots showing individual cell measurements (small, light-colored symbols; N>45 per biological replicate) and the mean value for each independent biological replicate (larger, black-outlined circles; n=3). Each color represents a different experimental replicate. Horizontal black lines and error bars indicate the mean and SEM of the n=3 biological replicates. Two-sided parametric ratio paired t-test was used. The P values are indicated in the plot. Mechanical forces impact Golgi membrane tension Our previously published work using optical tweezers demonstrated that forces applied directly on Golgi membranes alter their mechanical properties, and that actin depolymerization decreases Golgi rigidity ( Guet et al., 2014 ). More recently, fluorescence lifetime imaging microscopy (FLIM)-based sensors of the Flipper family, such as Halo-Flippers, have been developed as lipid packing and membrane tension reporters ( Strakova et al., 2019 ; Colom et al., 2018 ). Flippers respond to lipid packing, and thereby membrane tension, by shifting between planar and twisted conformations, which can be quantified by FLIM. We therefore tested whether extracellular mechanical forces alter Golgi tension using Halo-Flippers. First, to validate the use of Halo-Flippers at Golgi membranes, we exploited the effect of Latrunculin A on actin depolymerization, which is expected to reduce Golgi tension. RPE1-ManII-Halo stably expressing cells treated with low doses of Latrunculin A displayed compaction of the Golgi apparatus, as previously shown ( Egea et al., 2006 ), and a significant decrease in Halo-Flipper fluorescence lifetime at the Golgi ( Fig. S2A-B ), consistent with reduced membrane tension. These results align with our previous optical tweezers measurements ( Guet et al., 2014 ) and confirm that Halo-Flipper can report changes in Golgi membrane mechanical properties. We then examined the effect of two extracellular mechanical perturbations on Golgi tension. First, RPE1-ManII-Halo cells were seeded on FN-coated glass slides and allowed to spread for 0.5, 1, 2, 3, or 4 h (as outlined in Fig. 1A ). At each time point, cells were post-labeled with JF-646 to mark the Golgi area and Halo-Flipper was imaged by FLIM ( Fig. 3A ). Over the spreading time course, Halo-Flipper fluorescence lifetimes at the Golgi increased, suggesting a progressive rise in Golgi membrane tension that parallels FA assembly and cell spreading ( Fig. 3B ). Second, RPE1-ManII-Halo-expressing cells were seeded for 4h on stiff or soft FN-coated PAA gels (as described in Fig. 1 ) ( Fig. 3C ). FLIM measurements showed significantly lower lifetimes on soft substrates relative to stiff ones ( Fig. 3C-D ), suggesting reduced Golgi membrane tension under low mechanical load. Together, these data link integrin-mediated mechanical inputs, FA formation, Golgi membrane mechanics, cell spreading area, and post-Golgi carrier formation: robust spreading with abundant active FAs correlates with elevated Golgi tension and increased number of post-Golgi carriers, whereas limited adhesion on soft substrates corresponds to decreased Golgi tension and reduced post-Golgi carrier abundance. These findings establish that Golgi membrane mechanics responds to external mechanical forces, pointing to a potential mechanoadaptation of the Golgi function. Download figure Open in new tab Figure 3: Golgi membrane tension responds to mechanical forces. ( A ) Images of live RPE1-ManII-Halo stably expressing cells spread for 30 minutes to 4 hours on FN-coated glass coverslips. The FLIM signal is displayed. To specifically measure FLIM at the Golgi, ManII-Halo was post-labeled with JF-646. For each time point, higher magnification is shown on the right. ( B ) FLIM value expressed as fluorescence lifetime in ns was quantified at the indicated time points. ( C ) Images of live RPE1-ManII-Halo stably expressing cells plated on FN-coated glass coverslips, or stiff (30 kPa) or soft (2 kPa) FN-coated PAA gels. The FLIM signal is displayed. The Golgi apparatus was post-labelled using JF-646. ( D ) FLIM value expressed as fluorescence lifetime in ns was quantified in the different conditions. Mechanical cues drive microtubule acetylation to enhance Golgi export We next investigated how Golgi membranes respond to mechanical cues at the cell surface and which signaling pathways may relay forces to the Golgi apparatus. It is known that extracellular mechanical signals tune tubulin acetylation and GEF-H1/RhoA activation ( Seetharaman et al., 2022 ; Krendel et al., 2002 ), which in turn leads to PLCε activation and diacylglycerol (DAG) production at the TGN ( Eisler et al., 2018 ). DAG is necessary for CARTS biogenesis by recruiting and activating Protein Kinase D (PKD), a kinase required for the fission of a subset of Golgi-to-PM carriers, including CARTS ( Wakana and Campelo, 2021 ). Based on this, we hypothesized that MT acetylation could serve as one of the mediators transmitting mechanical signals from the cell surface to the Golgi membranes. Because astrocytes plated on stiffer substrates show higher levels of acetylated MTs ( Seetharaman et al., 2022 ), we tested whether other mechanical stimuli –cell spreading on different ligand coatings and equibiaxial cell stretch– similarly induce MT acetylation. First, to test whether MT acetylation mediates mechanotransduction from FAs to the Golgi membranes, we compared HeLa cells seeded on FN to those on PLL ( Fig. 4A ). By 1 h on FN, acetylated tubulin appears to be mostly localized in the perinuclear area ( Fig. 4A ). By 4 h, long protruding acetylated MTs extended from the perinuclear region toward the PM ( Fig. 4A ). This pattern was absent in PLL-plated cells at all time points ( Fig. 4A ). Quantification of the acetylated-to-total tubulin ratio revealed a time-dependent increase on FN that plateaued at 4 h, whereas cell on PLL displayed no consistent trend, possibly reflecting large cell-to-cell variability and lack of integrin engagement on PLL substrates ( Fig. 4B ). Next, to emulate externally applied stretch, we subjected HeLa cells to a 15% equibiaxial strain for 30 min using our stretching device ( Fig. 2A ). External stretch suggested a trend toward higher tubulin acetylation relative to unstretched controls ( Fig. 4C ), but the effect was heterogeneous across experiments and, overall, not statistically significant, indicating that additional factors may modulate this response ( Fig. 4D ). Download figure Open in new tab Figure 4. Cell spreading induces tubulin acetylation ( A ) Representative confocal microscopy images of the acetylated and total α-tubulin fractions of fixed HeLa cells seeded for the indicated times on FN- or PLL-coated glass coverslips. Scale bars, 10µm. ( B ) SuperPlot showing individual cell measurements (small, light-colored symbols; N∼10 per biological replicate) and the mean value for each independent biological replicate (larger, black-outlined circles; n=3 replicates). Each color represents a different experimental replicate. Horizontal black lines and error bars indicate the mean and SEM of the n=3 biological replicates. A repeated-measures one-way ANOVA test was performed only on FN-seeded cells. P values using Tukey’s post-hoc multiple comparison test are reported in the figure. ( C ) Representative confocal microscopy images of the acetylated and total α-tubulin fractions of fixed HeLa cells seeded over FN-coated PDMS membranes, and subjected to no external forces of 15% mechanical strain. Scale bars, 10µm. ( D ) SuperPlot showing individual cell measurements (small, light-colored symbols; N∼10 per biological replicate) and the mean value for each independent biological replicate (larger, black-outlined circles; n=3). Each color represents a different experimental replicate. Horizontal black lines and error bars indicate the mean and SEM of the n=3 biological replicates. A two-sided parametric ratio paired t-test was used. The P value is indicated in the plot. To determine whether MT acetylation alone is sufficient to increase the number of Golgi-derived carriers independently of external mechanical forces, we treated HeLa cells with tubacin, a histone deacetylase family member 6 (HDAC6) inhibitor that increases MT acetylation. As reported previously ( Haggarty et al., 2003b ; a), tubacin significantly elevated acetylated tubulin ( Fig. S3A-B ), and significantly increased the number of PAUF-mRFP– positive carriers released from the TGN, without any significant increase in adhesion area ( Fig. 5A-D , Fig. S3C ). Tubacin treatment in RPE1-ManII-Halo-expressing cells did not lead to significant increase in Golgi membrane tension as measured by Halo-Flipper FLIM ( Fig. 5E-F ), suggesting that in addition to tubulin acetylation other signaling pathways may control Golgi membrane tension and function. Download figure Open in new tab Figure 5. Microtubule hyperacetylation promotes CARTS biogenesis. ( A ) Confocal microscopy images of fixed HeLa cells transfected with the PAUF-RFP plasmid, subjected to a treatment with or without Tubacin. Scale bars, 10µm. ( B-D ) SuperPlots showing individual cell measurements (small, light-colored symbols; N∼10 per biological replicate) and the mean value for each independent biological replicate (larger, black-outlined circles; n=3) of ( B ) Number of CARTS per cell, (C) Adhesion area per cell (µm 2 ), and (D) Number of CARTS per adhesion area. A two-sided parametric ratio paired t-test was used. The P value is indicated in the plot. ( E ) FLIM Images of live RPE1-ManII-Halo stably expressing cells before and after overnight treatment with 10 µM of Tubacin. The FLIM signal (lifetime) is displayed. Golgi was post-labelled using JF-646 (intensity shown, JF646 channel). Higher magnification is shown on the right. ( F ) FLIM-measured fluorescence lifetime (in ns) was quantified for each experimental condition. Collectively, these results indicate that both physiological spreading forces as well as externally applied stretch enhance MT acetylation, and that MT hyperacetylation is sufficient to promote Golgi export. Thus, MT acetylation emerges as a key component of the mechanotransduction pathway from the cell surface to the Golgi apparatus, although our data suggests that additional routes likely contribute to the regulation of Golgi membrane tension and function. Future work will be needed to define how these parallel pathways are integrated to coordinate Golgi mechanoresponse and mechanosensitivity. Golgi DAG levels are modulated by mechanical stimuli Having shown that mechanical cues and MT acetylation enhance CARTS formation, we next asked whether these inputs also regulate Golgi diacylglycerol (DAG) levels, since DAG recruits and activates PKD to promote carrier biogenesis downstream of RhoA/PLCε signaling ( Eisler et al., 2018 ). To monitor DAG at the Golgi area, HeLa cells were transfected with the GST-C1a-PKD DAG biosensor ( Maeda et al., 2001 ) and analyzed by immunofluorescence microscopy. First, we compared cells seeded on FN with cells seeded on PLL ( Fig. 6A ). Cells on FN accumulated higher Golgi DAG levels than cells on PLL ( Fig. 6A, B ). In FN-seeded cells, Golgi DAG tended to decline over prolonged spreading, perhaps reflecting DAG incorporation into nascent transport carriers or conversion into other lipids needed for membrane fission ( Campelo and Malhotra, 2012 ). By contrast, cells on PLL showed an increase in Golgi DAG by 24 h, paralleling the rise in tubulin acetylation observed under those conditions ( Fig. 4B ). Second, we compared DAG levels at the Golgi area in cells on soft (2 kPa) versus stiff (30 kPa or glass) FN-coated PAA gels ( Fig. 6C-D ). Consistent with our spreading data, cells on stiff substrates showed higher DAG content at the Golgi area than cells on soft substrates, paralleling increased MT acetylation and Golgi-derived carrier biogenesis under these conditions. Download figure Open in new tab Figure 6. Mechanical cues alter Golgi DAG content. ( A ) Representative confocal microscopy images of fixed HeLa cells transfected with the GST-C1a-PKD DAG sensor, immunostained for TGN46, seeded over FN or PLL for the indicated times. Scale bars, 10µm. ( B ) SuperPlot (N∼10 cells per biological replicate; n=3 biological replicates) of the quantification of ( A ). A repeated-measures 2-way ANOVA test was performed. P values using Tukey’s post-hoc multiple comparison test are reported. ( C ) Representative images of HeLa cells transfected with the GST-C1a-PKD DAG sensor, immunostained with GM130, and seeded on FN-coated glass coverslips, stiff (30 kPa) or soft (2 kPa) PAA gels. ( D ) SuperPlot showing quantification of ( C ). ( E ) Representative confocal microscopy images of fixed HeLa cells transfected with the GST-C1a-PKD plasmid, immunostained with an α-TGN46, and subjected to a treatment with or without 10 µM Tubacin. Scale bars, 10 µm. ( F ) SuperPlot (N∼10 per biological replicate; n=3 biological replicates) showing quantification of ( E ). A two-sided parametric ratio paired t-test was used. The P value is indicated in the plot. Finally, to test whether MT acetylation alone modulates Golgi DAG levels, we treated HeLa cells with tubacin ( Fig. 6E-F ). Whereas tubacin-induced MT hyperacetylation increased the number of cytoplasmic CARTS ( Fig. 5B ), it caused only a modest, non-significant increase in Golgi DAG content ( Fig. 6E-F ). Consistently, Golgi membrane tension remained unchanged upon tubacin treatment ( Fig. 5E,F ), indicating that full DAG accumulation and membrane tension increase at the Golgi membranes likely require additional force-dependent activation of other DAG-synthesizing pathways ( Romani et al., 2019 ). Taken together, these findings position DAG modulation at the Golgi as a potential mechanotransduction pathway linking cell surface mechanical forces to Golgi function. Future work employing more sensitive DAG level measurements, such as targeted lipidomic analyses will be needed to validate and further dissect the dynamics of this lipid-mediated regulation. Golgi-derived export is necessary for cell spreading and mechanoadaptation Our data so far suggest the existence of a critical feedback loop between the cell surface – particularly FAs– and the Golgi apparatus, by which extracellular mechanical cues are transduced to the Golgi to modify its lipid composition, mechanical properties, and secretory capacity. We thus asked whether this feedback loop exists, so this Golgi response is functionally required for proper cell adhesion and spreading. To investigate this, we performed spreading assays in HeLa cells expressing the membrane marker GFP-mem, while selectively impairing Golgi export using different strategies. First, we treated cells with Golgicide A, a general inhibitor of Golgi-to-PM trafficking ( Sáenz et al., 2009 ) . Although initial cell spreading within the first hour was unaffected, a significant reduction in cell adhesion area was evident by 4 h in Golgicide A-treated cells ( Fig. S4A,B ), suggesting that Golgi-derived export becomes an important contributor to continued spreading. We next inhibited PKD using CRT0066101. PKD inhibition decreased the number of cytoplasmic CARTS ( Fig. 7A,B ) and led to a similar phenotype characterized by reduced expansion of the adhesion area at 4 h ( Fig. 7C,D ). In parallel, FLIM measurements in RPE1-ManII-Halo cells showed that CRT0066101 treatment also reduced Golgi membrane tension, as reported by Flipper lifetime ( Fig. 7E,F ). These data are consistent with previous work showing that PKD activity supports directional migration and invasion in breast cancer models ( Borges et al., 2015 ). Hence, disruption of PKD-dependent carrier biogenesis reduces both Golgi membrane tension and cell spreading efficiency. Because PKD also regulates actin dynamics and transcription ( Fu and Rubin, 2011 ), we took a more specific approach to selectively impair CARTS-mediated export. HeLa TGN46 KO cells, which lack the transmembrane cargo adaptor TGN46 that is required for cargo sorting into CARTS ( Lujan et al., 2024 ), exhibited significantly reduced spreading compared to wild-type (WT) cells, a phenotype that persisted for at least 24 h post-seeding ( Fig. 7G, H ). Collectively, our results underscore that the Golgi apparatus actively responds and adapts to extracellular mechanical cues ( Fig. 7I ). The ability of the Golgi apparatus to sense, respond, and secrete in response to extracellular mechanical forces supports proper cell adhesion, spreading, and homeostasis. Our work unveils new concepts in organelle mechanobiology, highlighting key mechanotransduction steps at the Golgi apparatus, a secretory mechanoresponse, and the importance of feedback communication between the PM and internal membranes in maintaining cellular homeostasis. Download figure Open in new tab Figure 7: Golgi-derived export is necessary for cell spreading and mechanoadaptation. ( A ) HeLa cells transfected with the PAUF-RFP plasmid were spread on FN for 4h and subjected to a treatment with (5 µM) or without CRT0066101, a PKD inhibitor, fixed, and imaged by confocal microscopy. Scale bar, 10 µm. (B) SuperPlot showing the number of CARTS per cell for each experimental condition. (C) Representative confocal microscopy images of fixed HeLa cells transfected with the GFP-mem plasmid, subjected or not to a treatment with CRT0066101. Scale bars, 10 µm; representative cell length scales shown. (D) SuperPlots showing individual cell measurements (N∼10 per biological replicate; n=3 biological replicates). A two-sided parametric ratio paired t-test was used. The P value is indicated in the plot. ( E ) Images of live RPE1-ManII-Halo stably expressing cells before and after treatment with 5 µM CRT0066101. The FLIM signal and lifetime are displayed. The Golgi apparatus was post-labelled using JF-646. Higher magnification is shown on the right. (F) SuperPlot showing FLIM values expressed as fluorescence lifetimes in ns was quantified for each experimental condition. (G) Representative confocal microscopy images of fixed HeLa cells (WT or TGN46 KO), transfected with the GFP-mem plasmid, and seeded over FN for the indicated times. Scale bars, 10µm. (H) SuperPlot showing individual cell measurements (N∼10 per biological replicate; n=3 biological replicates) quantifying adhesion area from images in ( G ). A repeated-measures 2-way ANOVA test was performed. P values using Tukey’s post-hoc multiple comparison test are reported. ( I ) Schematic representation of our findings (see text for details). DISCUSSION Our work reveals a mechanotransduction axis linking extracellular forces to the Golgi complex. These forces are transduced to remodel Golgi membranes by tuning their biophysical and biochemical characteristics, and lead to an increase in the number of cytoplasmic Golgi-derived carriers (CARTS as well as CD59-positive carriers) to support adhesion and efficient spreading. Using three distinct mechanical stimuli (spreading forces on integrin-activating substrates, substrate stiffness, and equibiaxial stretch), we show that all of them increase the number of Golgi-derived carriers. In parallel, these cues elevate Golgi membrane tension (measured by Halo-Flipper FLIM) and Golgi DAG levels (measured by a fluorescent DAG biosensor). We provide evidence that MT acetylation acts as a critical downstream effector of mechanical cues and is sufficient to boost post-Golgi carrier formation even in the absence of external forces. Finally, we show that Golgi export –particularly via PKD- and CARTS-dependent routes– is both upregulated by mechanical stimuli and required for sustained cell spreading, suggesting a reciprocal communication between FA establishment, cell spreading, and directed Golgi-to-PM trafficking to exocytic hotspots. Ample evidence supports the idea that exocytosis in adherent cells is not uniformly distributed across the cell surface but occurs at secretion hotspots, typically juxtaposed to FAs ( Stehbens et al., 2014 ; Eisler et al., 2018 ; Huet-Calderwood et al., 2022 ; Fourriere et al., 2019 ). FAs are molecular platforms required for adhesion and for sensing and transducing extracellular mechanical forces ( Kechagia et al., 2019 ; Chastney et al., 2025 ). Integrin β1 itself is delivered from the TGN to the plasma membrane through a RAB6-mediated ( Huet-Calderwood et al., 2022 ) and PKD-dependent pathway ( Yeaman et al., 2004 ), directly linking Golgi export routes to the reinforcement of adhesion complexes. However, whether –and how– these mechanical cues drive adaptation of the secretory machinery has remained largely unexplored. Notably, recent studies show that ER exit sites and ER export can be upregulated upon extracellular mechanical cues by the GTPase Rac1 ( Phuyal and Baschieri, 2020 ; Phuyal et al., 2022 ), and optogenetic and other mechanostimulatory perturbations alter ER export dynamics ( Song et al., 2024 ; Chen et al., 2025 ). In addition, FAs and ECM can regulate the COPII machinery involved in ER to Golgi transport ( Jung et al., 2022 ) . Collectively, these reports point to coordinated, adhesion-dependent regulation across the secretory pathway. Our findings therefore position the Golgi as an active mechanoresponsive organelle, extending mechanotransduction beyond conventional mechanosensor organelles, such as the plasma membrane and nucleus. Our data place MT acetylation downstream of extracellular mechanical inputs and upstream of Golgi functional adaptation, consistent with its role as a tension-sensitive mediator ( Seetharaman et al., 2022 ) and a regulator of PKD-dependent Golgi trafficking ( Eisler et al., 2018 ). By showing that inhibiting tubulin deacetylation (using the HDAC6 inhibitor tubacin) partially phenocopies mechanical stimulation, we identify MT acetylation as both a sensor and an amplifier of force signals, upstream to the GEF-H1/RhoA/PLCε/DAG/PKD axis previously implicated in Golgi export ( Eisler et al., 2018 ). We propose a model ( Fig. 7I ) in which mechanical forces –transduced via a yet-unidentified mechanoreceptor– trigger increased microtubule acetylation, releasing GEF-H1 and sustaining RhoA signaling ( Seetharaman et al., 2022 ). RhoA then reinforces actomyosin contractility through ROCK and also promotes DAG production at the TGN via PLCε, recruiting and activating PKD to drive carrier biogenesis ( Eisler et al., 2018 ) ( Fig. 7I ). Too low Golgi membrane tension may render the TGN inefficient for carrier fission or alternatively may be a consequence of an altered TGN lipidome, so the full extent of this mechanical adaptation warrants further study. Of note, elevated membrane tension is known to inhibit membrane curvature generation ( Dai and Sheetz, 1995 ; Carlsson, 2018 ; Le Roux et al., 2021 ), highlighting the importance of a well-balanced, dynamic control of Golgi membrane tension for secretory function. Importantly, our observations can be placed in the context of opposing effects of membrane tension on endocytic versus exocytic pathways. High plasma-membrane tension is well-documented to inhibit endocytosis ( Apodaca, 2002 ; Kosmalska et al., 2015 ; Wu et al., 2017 ), a block that can be overcome locally by polymerization of actin to provide the force required for vesicle invagination and scission ( Boulant et al., 2011 ; Gauthier et al., 2011 ) . By analogy, actomyosin contractility may not be purely inhibitory but also contribute to promote the biogenesis and fission of post-Golgi carriers by shaping Golgi membrane mechanics and promoting DAG/PKD-dependent scission; thus, actin dynamics can act as a context-dependent permissive or driving force for membrane trafficking depending on the compartment and the direction of membrane flow. Notably, we found that tubacin-induced MT hyperacetylation induced only a modest increase in Golgi DAG and no measurable change in Golgi tension, suggesting that additional regulatory layers and other effectors, such as Lipin-1 ( Romani et al., 2019 ), contribute to full DAG accumulation and tension modulation. We note, however, that Flipper probes report membrane lipid packing/ordering as a proxy for membrane tension, and perturbations that alter Golgi lipid composition (e.g., PKD inhibition or DAG modulation) can alter Flipper lifetimes independently of tension reporters ( Colom et al., 2018 ). Thus, while our FLIM data are consistent with tension changes, interpretation of lifetime shifts as pure mechanical readouts should be cautious and will require complementary validation in future work. The precise molecular links between MT acetylation, Golgi membrane tension, and PKD activation remain to be defined ( Fig. 7I ). By uncovering these connections, our work establishes Golgi-based mechanoadaptation as a key mechanism of cellular regulation and opens important avenues for future investigation. Reciprocal feedback between exocytosis and membrane tension also provides a physiological rationale for the coupling we observed: exocytosis lowers PM tension by adding membrane facilitating further spreading and migration ( Gauthier et al., 2011 ; Pontes et al., 2017 ) . Thus, upregulation of Golgi-to-PM trafficking and secretion at adhesion-proximal hotspots can both respond to and modulate PM mechanics, promoting processes such as migration, tissue morphogenesis, and membrane homeostasis. In pathological contexts, such as cancer invasion, fibrosis, and wound healing, altered ECM mechanics could dysregulate this Golgi mechanoresponse, driving aberrant secretion of matrix components or signaling factors. Notably, delivery of newly synthesized integrin β1 relies on RAB6- and PKD-dependent Golgi export ( Huet-Calderwood et al., 2017 ; Yeaman et al., 2004 ), suggesting that misregulation of this pathway could directly impact adhesion turnover and invasive behavior. Targeting elements of this axis (e.g., HDAC6, PLCε, PKD, TGN46, or other intermediates yet to be identified) may offer novel therapeutic strategies to modulate mechanoresponsive secretion. Finally, extending these studies to polarized cells, 3D matrices, and in vivo models will be essential to understand Golgi-mediated mechanoregulation at the tissue level. In summary, we uncovered a two-way dialogue between the plasma membrane and the Golgi apparatus that is mediated by mechanical forces, microtubule acetylation, and TGN signaling (DAG and PKD). This feedback loop may allow cells to adapt their adhesive and migratory behavior to the mechanical properties of their environment, thus positioning the Golgi as a new mode of cellular mechanoregulation. METHODS Reagents, plasmids, antibodies The Paxillin-pEGFP plasmid was a gift from Rick Horwitz (Addgene plasmid # 15233; http://n2t.net/addgene:15233 ; RRID:Addgene_15233) ( Laukaitis et al., 2001 ). The mKate2-FM4-PAUF plasmid was a gift from Yuichi Wakana (Tokyo University of Pharmacy and Life Sciences, Japan) ( Wakana and Campelo, 2021 ). The Double palmitoylated Neuromodulin (N-terminal)-GFP (GFP-mem) plasmid is used as a general PM marker as it is a fusion protein with a signal for post-translational palmitoylation targeting the fusion protein to the cell membrane, and was a gift from Francesc Tebar’s lab (Universitat de Barcelona, Spain) ( Vidal-Quadras et al., 2011 ) . The plasmid encoding for PAUF-mRFP ( Wakana et al., 2012 ) and C1a-GST-PKD ( Maeda et al., 2001 ) were donated by Vivek Malhotra (CRG, Barcelona, Spain). Commercially available antibodies used were as follows: anti TGN46 (sheep) antibody was purchased from Bio-RAD (AHP500GT). Anti-GM130 (mouse) was from BD Biosciences (610822). Anti-α-tubulin (rabbit) antibody was from Abcam (ab18251) and anti-tRFP (rabbit) antibody was from Evrogen (AB233). Anti-acetylated-α-tubulin mouse (T6793) and anti-Glutathione-S-Transferase (GST) rabbit (G7781) were obtained from Sigma-Aldrich. Secondary antibodies used were Alexa488 (A32766), Alexa555 (A32794), or Alexa647 (A21448) coupled donkey anti-mouse, anti-rabbit and anti-sheep immunoglobulin G (IgG) (Invitrogen). Janelia Fluor probes (JF-646) were from Promega. The following reagents were purchased from Sigma-Aldrich: Fibronectin (FN) (11051407001), Poly-L-Lysine (PLL) (P1524), Tubacin (SML0065), Golgicide A (345862) and Cycloheximide (CHX) (C4859). D/D solubilizer (635054) was obtained from Takara Bio. CRT0066101 was from Tocris Bioscience (4975). Sylgard™ 184 Elastomer KIT was from Dow Inc (24001673921). Cell culture and transfection HeLa cells were cultured in DMEM (Capricorn Scientific GmbH) supplemented with 10% FBS (Invitrogen), 1% penicillin-streptomycin (Gibco) and L-Glutamine (LabClinics) under 5% CO 2 at 37°C. HeLa Cells were transiently transfected using X-tremeGENE™ 9 DNA transfection reagent (Sigma-Aldrich) following the manufactureŕs instructions. Cells were used for the designed experiments ∼16h post-transfection. In experiments where cells were placed at 20°C for 2h to synchronize cargo release from the Golgi apparatus, cell culture medium was supplemented with HEPES 25mM (Sigma-Aldrich). Additionally, cell culture medium was supplemented with CHX 100 µM to inhibit new protein synthesis before cargo release. TGN46 KO HeLa cells were generated as described in ( Lujan et al., 2024 ). RPE1 cells were cultured in DMEM/ F12 supplemented with 10% fetal bovine serum (FBS) under 5% CO 2 at 37°C. RPE1 cells were transfected using Lipofectamine 3000 (Invitrogen) following the manufactureŕs instructions. RPE-ManII-Halo stable cell line was generated as follows: RPE1 cells were transfected with pIRES-ManII-Halo (ManII-Halo sequence from prHom-ManII-Halo, kind gift from D. Toomre, was inserted into pIRES-Neo3) and single clones were then selected following Ampicillin selection. Cell treatments For MT acetylation and DAG production experiments, cells were pre-seeded for 1h on FN-coated coverslips (10µg/ml final concentration, overnight (o/n) incubation) and then treated with 10µM Tubacin or DMSO for 3h continuously. For CARTS formation experiments, cells pre-seeded on FN coverslips were subjected to a pre-treatment of 30 minutes with Tubacin or DMSO, followed by 2h incubation at 20°C in presence of the drug, and 30 minutes CARTS release with Tubacin still present in the samples. For PKD inhibition experiments, cells were pre-seeded over FN-coated coverslips for 30 minutes and then incubated with CRT0066101 5µM or DMSO for 30, 210 minutes or 24h. Combined spreading-secretion assay For cell spreading assays (e.g., Fig. 1A,B ), HeLa cells transiently transfected with the GFP-mem and FM4-mKate2-PAUF constructs or RPE1-ManII-Halo stable cells were split and seeded over a ligand-coated coverslip (FN or PLL). Unless otherwise stated, cells were lifted by incubation with Trypsin (0.05% trypsin, 0.53 mM EDTA) for 3–5 min in a 37°C incubator. When lifted using EDTA, cells were incubated with 10 mM EDTA in Ca 2+ and Mg 2+ free PBS for 10 min in a 37°C incubator. Cells were kept at 37°C, and 30 min before the end of the spreading times, cell culture medium was supplemented with CHX 100µM and with D/D solubilizer 1µM to allow cargoes to re-enter the secretory pathway. Cells were then fixed, mounted, and visualized to determine the effects of spreading time and ligand on secretion. For secretion arrest experiments, 30 min pre-seeded cells were treated with Golgicide A 10 µM for 30 or 210min. The FM4-mKate2-PAUF construct contains the FM4 aggregation domain, retaining the cargoes in the ER after synthesis. By adding to the cell medium D/D solubilizer, a small molecule responsible for the solubilization of the FM4 aggregates, it is possible to synchronize a wave of cargo release from the ER and follow their intracellular localization in time, all the way from the ER, to the Golgi apparatus and to CARTS for secretion. For those cells transiently transfected with mKate2-FM4-PAUF plasmid, D/D solubilizer was added at 1µM final concentration to allow the cargoes to be exported out of the ER. Immunofluorescence and fluorescence microscopy imaging Cells were fixed with 4% (v/v) paraformaldehyde (PFA) in PBS for 15 minutes at room temperature (RT). After three washes with PBS, cells were permeabilized with 0.2% TX-100 in PBS for 30 minutes at RT. After washing with PBS, blocking was performed with 4% bovine serum albumin (BSA) for 30 minutes at RT or o/n at 4°C. After washing the samples, the BSA solution at 2% was used to dilute the primary and secondary antibodies for incubation 1h at RT. Samples were then mounted on glass slides using either ProLong Gold Antifade Reagent (Thermo Fisher Scientific) or Mowiol. Widefield microscopy images were acquired on an upright Nikon Eclipse Ni-U microscope equipped with a 60× water-dipping objective (NIR Apo 60×/WD 2.8, Nikon), an ORCA-Flash 4.0 camera (Hamamatsu), and controlled by Metamorph software. Confocal microscopy images were acquired on either (i) a TCS SP8 microscope (Leica Microsystems GmbH, Germany) equipped with an HC PL APO CS2 100x/1.40 oil objective, a pulsed white light laser (WLL) operating at 20 MHz repetition rate, 70 % master power and a hybrid detector (HyD) in photon-counting mode and 8-bit depth; or (ii) a spinning disk confocal microscopy images were acquired on a Nikon Inverted Eclipse Ti-E (Nikon) microscope equipped with a Spinning disk CSU-X1 (Yokogawa), an iXon EMCCD camera (Andor) or sCMOS Kinetx 22 camera (Photometrics) integrated in Metamorph software by Gataca Systems, using 60x or 100x CFI plan apochromat VC 1.4 NA oil immersion objectives (Nikon). The spinning disk microscope was in a cage incubator from Life Imaging Services for temperature control. GFP, mKate2, RFP, Alexa488, Alexa647 were excited at 489, 589, 554, 499, 653 nm, respectively, and the emission was detected between 499-580, 599-770, 564-750, 509-600, 663-781nm, respectively. For dual and triple-color imaging, sequential scan mode between lines was used. For z-stack acquisitions, the distance between confocal planes was set as system optimized by the microscope acquisition software, the planes were reaching throughout the entire cell volume, and maximum intensity (for visual representation) or sum slices projections (for total intensity measurements) were computed. Typically, images were acquired using 2x line accumulations. Laser power was set to prevent fluorophore saturation and maintained throughout the whole experiment. Images were analyzed using ImageJ software. Selective protein immobilization (SPI) assay The assay was firstly described in (Fourriere et al., 2019). Briefly, coverslips were incubated with bicarbonate buffer 100mM for 1h at 37°C. Then, coverslips were washed three times in PBS and dried before incubation with the anti-tRFP antibody for 3h at 37°C or o/n at 4°C, using a 1:250 dilution. For the detection of the coated antibody, anti-rabbit Star Orange (Abberior) was used. Live cell imaging Images were acquired with a commercial Nikon Eclipse Ti System, equipped with a 100x oil objective with NA 1.49 using TIRF illumination. The detection was carried out using an ANDOR technology EMCCD iXon 897 camera. The equipment presents an Agilent technologies laser box with wavelengths of 488 and 560 nm. Laser power was adjusted for each channel to prevent saturation. For the experiments tracking post-Golgi carriers transport to FAs, cells were seeded on FN-coated 35 mm glass bottom dishes (MatTek), and cells were subjected to a cargo release synchronization before imaging. Before cargo channel acquisition, an image of the FAs channel was taken as a reference. Then, cargo channel was acquired 1fps for 45 minutes. For SPI live cell imaging, both channels were acquired quasi simultaneously every 30s for 1h. Retention using selective hooks (RUSH) assay The RUSH assay was performed as described in ( Boncompain et al., 2012 ). Fluorescence lifetime imaging microscopy (FLIM) A FLIM module from PicoQuant (Kit LSM Upgrade pour Nikon A1 PicoQuant, Opton Laser International) was added to the A1R Nikon confocal microscope with both resonant and galvano mode scanners. Images were acquired by using 485 nm laser excitation and a 27.5ns dwell time per pixel with5-frame acquisition. The emission bandpass filter wavelength was 506-619nm. Single-stack images were captured and used for FLIM using TCSPC (Time-Correlated Single Photon Counting principle). Images were analyzed using the SymPhoTime 64 software (PicoQuant, Opton Laser International). The image acquisition time was typically 48 seconds for a 512×512 image. A region of interest was drawn manually based on a Golgi mask (Janelia Fluor® 646 HaloTag channel, see below for details) using the Screen Dragon software. The photon arrival time histograms were fitted with a double exponential model, after deconvolution with the calculated impulse response function in SymPhoTime. Upon fitting, the average fluorescence lifetime was recorded for analysis. Halo-Flipper ManII-Halo and ST-Halo RPE1 cells were labeled with Halo-Flipper in all the experiments to report lipid packing defects in Golgi membranes due changes in membrane tension or lipid composition. Cells were plated on Fluorodishes with glass or PAA gel substrates from 30 minutes to 4 hours (depending on the experiment). Halo-Flipper (kindly provided by S. Matile Lab) of 100nM concentration was prepared in 200uL serum free media and added to the cells by replacing the old media. After a 15 min incubation, Halo-Flipper was replaced with 160 µL media (+ 10% FBS) + 40µL of 1µM Janelia Fluor® 646 HaloTag (JF-646, 200 nM final working solution) and further incubated for 15 minutes. before FLIM and fluorescent images of the Golgi membrane were captured. Halo-Flippers were gently imaged avoiding as much as possible light exposure of the samples before FLIM recording. To minimize possible Flipper-induced phototoxicity and singlet oxygen photosensitization ( Torra et al., 2024 ), individual cells were measured only once. Halo-Flippers were made commercially available in 2025 and are now supplied by Spirochrome (Halo-Flipper, Spirochrome). Janelia Fluor HaloTag ligands Janelia Fluor® 646 HaloTag (JF-646) was used at 1:2000 dilution in culture media. Cells were labelled by incubation for 15 mins in a 37 °C with 5% CO 2 cell culture incubator. Preparation of polyacrylamide (PAA) gels PAA gels were prepared as described in (Pérez-Gonzales, et al. 2021). Preparation of PDMS stretchable membranes and ring mounting The whole protocol is described in ( Le Roux et al., 2025 ) . Briefly, the PDMS mixture was prepared in a 1:10 ratio and degassed in a vacuum chamber for 1h. PDMS was spread over PMMA plates using a spin coater in 2 steps: a first step during 5 s at 500 rpm with a 100 rpm/s acceleration, and a second step during 1min at 500 rpm with a 300 rpm/s acceleration. PDMS was polymerized o/n at 65°C in an oven. PDMS membranes were peeled off from the PMMA plates and mounted on the stretch-rings. PDMS rings were sterilized by exposing them to UV light for 15 min and then were coated with FN o/n. The following day, FN excess was rinsed off and cells were seeded on top of the FN-coated PDMS membranes for 1h prior the experiment. For experiments carried out with non-transiently transfected cells, cells were subjected to 15% stretch for 30 min at 37°C before fixation. In experiments characterizing the formation of CARTS upon mechanical strain stimulation, an intermediate step at 20°C for 2h to synchronize cargo release was set before stretching. Control experiments were performed under the same conditions, but in the absence of mechanical strain. Cells subjected to mechanical forces were fixed and imaged under stretch to prevent visual aberrations. Quantification of images and movies Analysis of the cell area was performed using a custom-designed macro in ImageJ. In brief, for each image cell area was determined by thresholding the GFP-mem channel, converting it to a mask and creating the selection. To determine number of CARTS, the ImageJ “Detect particles (ComDet)” plugin developed by Eugene A. Katrukha (Utrecht University, the Netherlands) was used, avoiding the perinuclear Golgi-positive area, using a maximum intensity z-projection of the entire confocal fluorescence microscopy z-stack of the cell or the epifluorescence image of the cell (when imaging the substrate stretching experiments). The normalized total intensity levels of acetylated tubulin were calculated according to the equation: Ratio of acetylated/total tubulin level = Acetylated tubulin intensity/Total tubulin intensity. DAG production (area and intensity) and Golgi area were calculated manually drawing an ROI around the fluorescent-positive mass present in the perinuclear area of the cells. In the bulk SPI analysis, FAs and secreted cargo objects were segmented and the localization and fluorescence intensity of each pixel was calculated. The nearest neighbor distance (nnd) between each pixel of secreted cargo to the closest pixel of FAs was measured using MatLab. The intensity of each pixel of secreted cargo was also included to obtain the 2D plots. CD59-positive vesicle count analysis was carried out using the cells expressing RUSH cargo of interest, which were fixed at 10 min, 35 minutes, and 50 minutes post-biotin addition to record ER to Golgi, early post-Golgi, and late post-Golgi carriers, respectively. The vesicles/carriers were manually counted from maximum intensity z-projection of the entire z-stack confocal fluorescence microscopy image of the cell using ImageJ and used for further analysis. Statistical analysis and data representation Data is represented using SuperPlots ( Lord et al., 2020 ), as indicated in the corresponding figure panels, individual cell measurements are shown as small, light-colored symbols, and the mean values for each independent biological replicate are shown as larger, black-outlined circles. Each color represents a different experimental replicate. Horizontal black lines and error bars indicate the mean and standard error of the mean (SEM) of the biological replicates. Statistical analysis was obtained using specific tests as indicated in the figure legends. Analyses were performed with GraphPad Prism 6.0 or 10.0. P-values are reported as numerical values, and specific statistical tests used are stated in each figure legend, using the number of independent experiments (and not the number of individual cells) as n values. SUPPLEMENTARY FIGURE LEGENDS Download figure Open in new tab Figure S1. CARTS are delivered close to FAs in a polarized manner. (A) TIRF image of a live HeLa cell expressing Paxillin-eGFP (green) and mKate2-FM4-PAUF (magenta). White squares highlight (i) FA-enriched (FA+) and (ii) FA-non-enriched (FA-) regions within the plasma membrane of the cell. Scale bar, 10µm. Zoom-ins of the highlighted regions show a frame time sequence of the two merged channels. White circles highlight the vesicle disappearance along the frame sequence (i), and a random position of equal size in the FA-region of the cell. Plots on the right correspond to the fluorescence plot profiles of mKate2-FM4-PAUF measured from the highlighted regions in the zoom-ins. Scale bar, 1µm. (B) HeLa cell expressing Paxillin-GFP (green), and mKate2-FM4-PAUF (magenta), subjected to an SPI assay (see Methods), fixed at the indicated times after cargo release from the ER, and imaged by confocal fluorescence microscopy. Schematic of the experiment is shown on the right. D/D is D/D solubilizer, CHX is cycloheximide. Scale bar, 10µm. ( C ) Quantification of ( B ), showing a 2D histogram (and corresponding 1D projections) as a function of mKate2-FM4-PAUF fluorescence intensity per pixel and distance from that pixel to the closest FA area (see Methods for details) at the indicated time points. ( D ) Confocal fluorescence microscopy images of fixed HeLa cells transiently transfected with Paxillin-GFP and seeded on PLL for 4h. Scale bar, 10µm. ( E ) SuperPlot showing the number of CARTS per cell (small, light-colored symbols; N∼10 per biological replicate) on cells lifted using Trypsin or EDTA (see Methods). The mean value for each independent biological replicate (larger, black-outlined circles; n=3). Repeated-measures 2-way ANOVA tests were performed, and P values were obtained using Tukey’s post-hoc multiple comparison test, with only the values for the comparisons between Trypsin and EDTA being shown. ( F ) Spinning disk confocal fluorescence microscopy images of fixed HeLa cells that were plated on stiff and soft PAA gels, let spread for 4h, fixed, and stained for vinculin, a FA marker. Scale bar, 10 µm. Download figure Open in new tab Figure S2: Golgi membrane tension responds to mechanical forces. ( A ) Images of live RPE1-ManII-Halo stably expressing cells before and after treatment with 200 nM Latrunculin A. The FLIM signal is displayed. The Golgi apparatus was post-labelled using JF-646. Higher magnification is shown on the right. Scale bar, 10 µm ( B ) SuperPlot of FLIM value expressed as fluorescence lifetime in ns was quantified for each experimental condition. Download figure Open in new tab Figure S3. Microtubule hyperacetylation upon Tubacin treatment. ( A ) Representative confocal microscopy images of fixed HeLa cells stained for acetylated or total microtubules subjected or not to a treatment with Tubacin. ( B ) SuperPlots showing the ratio between fluorescence intensity of acetylated vs. total tubulin following the different treatments. (C) Schematics of the pipeline followed in the Tubacin-induced stimulation of the secretory pathway. Download figure Open in new tab Figure S4: Golgi-derived export is necessary for cell spreading and mechanoadaptation. ( A ) Representative confocal microscopy images of fixed HeLa cells transfected with the GFP-mem plasmid, subjected or not to a treatment with Golgicide A. Scale bars, 10 µm; representative cell length scales shown. ( B ) SuperPlots showing individual cell measurements (small, light-colored symbols; N∼10 per biological replicate) and the mean value for each independent biological replicate (larger, black-outlined circles; n=3). Each color represents a different experimental replicate. Horizontal black lines and error bars indicate the mean and SEM of the n=3 biological replicates. A two-sided parametric ratio paired t-test was used. The P value is indicated in the plot. ACKNOWLEDGMENTS We thank members of the Single Molecule Biophotonics lab at ICFO, the ITEM lab at Institut Curie, the Intracellular Mechanics group and the Physics of Living Systems team at MSC, as well as Neus Sanfeliu-Cerdán, Yuichi Wakana, Santosh Phuyal, Hesso Farhan, Aurélien Roux and Vivek Malhotra for valuable discussions. We thank Merche Rivas, Marina Perez, Angel Sandoval, and Maria Marsal for technical support at ICFO. We thank Rick Horwitz, Yuichi Wakana, Francesc Tebar, Vivek Malhotra, and Stefan Matile for kindly sharing reagents, and Yuichi Wakana for critical reading of the manuscript. We acknowledge support from the Government of Spain (RYC-2017-22227, PID2019-106232RB-I00/10.13039/501100011033/110198RB-I00, PID2020-113068RB-I00/10.13039/501100011033 and PID2023-147711NB-100; PID2022-138282NB-I00 project funded by the MCIN/AEI/10.13039/501100011033/FEDER, UE; PID2022-142672NB-I00; and Severo Ochoa CEX2019-000910-S to ICFO and CEX2023-001282-S to IBEC), Fundació Privada Cellex, Fundació Privada Mir-Puig, and Generalitat de Catalunya (CERCA, AGAUR), ERC Advanced Grants NANO-MEMEC (GA 788546) and MechanoSynth (GA 101097753), as well as LaserLab 4 Europe (GA 654148). N.M. acknowledges funding from the European Union H2020 under Marie Sklodowska-Curie grant 754558-PREBIST. J. A.-C. acknowledges funding from the European Union H2020 under the Marie Sklodowska-Curie grant agreement No 847517. A.W. was supported by joint funding from an ICFO Student Research Fellowship (Fall 2024) and a grant from Homerton College, Cambridge, UK. 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