# Regulation of cell dynamics by rapid integrin transport through the biosynthetic pathway

Martina Lerche, Mathilde Mathieu, Hellyeh Hamidi, Megan Chastney, Guillaume Jacquemet, Bart Marlon Herwig Bruininks, Shreyas Kaptan, Lene Malerød, Nina Marie Pedersen, Andreas Brech, et al.

► To cite this version:

Martina Lerche, Mathilde Mathieu, Hellyeh Hamidi, Megan Chastney, Guillaume Jacquemet, et al. Regulation of cell dynamics by rapid integrin transport through the biosynthetic pathway. Journal of Cell Biology, 2026, 225 (2), pp.e202508155. 10.1083/jcb.202508155. hal-05491533

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Regulation of cell dynamics by rapid integrin transport through the biosynthetic pathway

Martina Lerche<fnref n="1" />*, Mathilde Mathieu<fnref n="1" />*, Hellyeh Hamidi<fnref n="1" />, Megan Chastney<fnref n="1" />, Guillaume Jacquemet<fnref n="1" />,<fnref n="2" />,<fnref n="3" />, Bart Marlon Herwig Bruininks<fnref n="4" />, Shreyas Kaptan<fnref n="4" />, Lene Malerød<fnref n="5" />,<fnref n="6" />, Nina Marie Pedersen<fnref n="5" />,<fnref n="6" />, Andreas Brech<fnref n="5" />,<fnref n="6" />, Nobuyuki Matoba<fnref n="7" />, Yuichiro Sato<fnref n="8" />, Ilpo Vattulainen<fnref n="4" />, Pere Roca-Cusachs<fnref n="9" />,<fnref n="10" />, Franck Perez<fnref n="11" />, Gaelle Boncompain<fnref n="11" />,%, Stéphanie Miserey<fnref n="11" /> and Johanna Ivaska<fnref n="1" />, <fnref n="3" />, <fnref n="12" />-<fnref n="14" /> #

Affiliations:

<footnote n="1">¹ Turku Bioscience Centre, University of Turku and Åbo Akademi University, FI-20520 Turku, Finland</footnote>

<footnote n="2">² Faculty of Science and Engineering, Cell Biology, Åbo Akademi University, Turku, Finland</footnote>

<footnote n="3">³ InFLAMES Research Flagship Center, University of Turku and Åbo Akademi University, Turku, Finland</footnote>

<footnote n="4">⁴ Department of Physics, University of Helsinki, P. O. Box 64, FI-00014 Helsinki, Finland</footnote>

<footnote n="5">⁵ Department of Molecular Cell Biology, Institute for Research, Oslo University Hospital, Montebello, Oslo, NorwayCancer</footnote>

<footnote n="6">⁶ Centre for Cancer Cell Reprogramming, Institute of Clinical Medicine, Faculty of Medicine, University of Oslo, Montebello, Oslo, Norway</footnote>

<footnote n="7">⁷ Department of Pharmacology and Toxicology, Brown Cancer Center, and Center for Predictive Medicine, University of Louisville School of Medicine, Louisville, Kentucky 40202, U.S.A.</footnote>

<footnote n="8">⁸ Department of Medical Pharmacy, Faculty of Pharmacy, Yasuda Women’s University, Hiroshima, Japan</footnote>

<footnote n="9">⁹ Institute for Bioengineering of Catalonia (IBEC), the Barcelona Institute of Technology (BIST), 08028 Barcelona, Spain</footnote>

<footnote n="10">¹⁰ Universitat de Barcelona, 08036 Barcelona, Spain</footnote>

<footnote n="11">¹¹ Intracellular Transport : Engineering and Mechanisms Laboratory, Institut Curie, PSL Research University, Sorbonne Université, Centre National de la Recherche Scientifique, UMR 144, Paris</footnote>

<footnote n="12">¹² Department of Life Technologies, University of Turku, Turku, Finland</footnote>

<footnote n="13">¹³ Western Finnish Cancer Center (FICAN West), University of Turku, FI-20520 Turku, Finland</footnote>

<footnote n="14">¹⁴ Foundation for the Finnish Cancer Institute, Tukholmankatu 8, FI-00014 Helsinki, Finland</footnote>

<footnote># Correspondence to joivaska@utu.fi</footnote>

<footnote>* Equal contribution</footnote>

<footnote>% current address: Institut NeuroMyoGene, Physiopathology and Genetics of Neuron and Muscle, CNRS UMR5261, INSERM U1315, Université Claude Bernard Lyon1, Lyon, France</footnote>
ABSTRACT

Constitutive integrin endocytosis and recycling control cell movement and morphology. In contrast, the role of newly synthesized integrins delivered via the biosynthetic pathway has been largely overlooked. We used the retention using selective hooks (RUSH) system to monitor the localization of newly synthesized integrins exiting the endoplasmic reticulum in space and time. We discovered that new integrin delivery to the plasma membrane is polarized and enhances cell protrusion and focal adhesion growth in an extracellular matrix-ligand-dependent manner. Motor-clutch modeling explained the increased adhesion as higher integrin availability driving recruitment of additional receptors. Unexpectedly, live-cell imaging revealed a subset of fast-emerging integrin vesicles rapidly transported to the cell surface to facilitate localized spreading. This unconventional secretion depended on cell adhesion and correlated with increased surface levels of immature, high-mannose glycosylated integrin, indicating bypass of the canonical Golgi-dependent secretory pathway. Thus, spatial plasma membrane-targeting of new integrins rapidly alters adhesion receptor availability, providing cells with added plasticity to respond to their environment.
INTRODUCTION

Cells sense and respond to the extracellular matrix (ECM) through integrins and integrin localization at the plasma membrane is essential for focal adhesion (FA) formation. Numerous studies have addressed the dynamics of FA assembly as a function of integrin diffusion along the plasma membrane, and integrin activation and tethering to the cytoskeleton through components of the integrin adhesion complexes (IAC), such as talin and vinculin ¹⁻³. On the other hand, FA disassembly has been investigated from the point of view of proteolytic cleavage of adhesion components, microtubule-dependent adhesion turnover and integrin endocytosis from the plasma membrane ⁴⁻⁶. In addition, the recycling of plasma membrane endocytosed integrin back to the cell surface has been extensively studied and implicated in the generation of local cell protrusions, co-trafficking with growth-factor receptors, cell invasion and system-level metastasis ⁷⁻⁹. These studies, however, have focused on endo/exocytic traffic of mature plasma membrane integrins. In contrast, very little is known about the targeting of newly synthesized integrins to the plasma membrane and their contribution to cell adhesion, spreading and FA dynamics. The relevance of integrin biosynthetic traffic is virtually unexplored and overlooked in the regulation of cell dynamics, predominantly owing to a lack of suitable methodology.

Integrins are a family of 24 heterodimeric cell-surface receptors composed of a larger α-subunit and a smaller β-subunit. The β1-subunit constitutes 12 integrin heterodimers which mediate adhesion to a variety of ECM molecules ¹⁰. Only relatively few studies have investigated integrin maturation and delivery to the plasma membrane. Early investigations exploring the regulation of integrin expression with pulse-chase metabolic labeling have shown that the rate of integrin heterodimer assembly and maturation is determined by the availability of integrin α-subunits ¹¹. The β1-subunits are produced in an excess ratio and are retained in their immature form in the endoplasmic reticulum (ER) where they await to assemble with newly synthesized α-integrins ¹¹⁻¹³. In the ER, newly synthesized integrins bind to Ca²⁺, which maintains receptors in an inactive bent conformation until they reach the cell surface. Ca²⁺ depletion blocks the trafficking of α5β1-integrins from the ER to the Golgi ¹⁴. Talin plays an important role in trafficking of newly synthesized integrins, as talin depletion causes accumulation of α5β1-integrins in the early secretory pathway ¹⁵. How talin regulates exocytosis of newly synthesized integrins remains unknown, and other players in the regulation of integrin biosynthetic traffic are yet to be identified. To date, studies on newly synthesized integrins have been based on metabolic pulse-chase labeling and utilization of conformation specific antibodies, which have limitations for real-time studies and visualization of integrin delivery. The development of the retention using selective hooks (RUSH) assay has permitted real-time tracking of newly synthesized proteins ¹⁶. A recent study employing this method concluded that post-Golgi carriers are not transported randomly to the cell surface. Instead exocytosis of versatile cargo is focused to areas close to FAs ¹⁷. However, whether this holds true for integrins and whether their delivery is influenced by specific ECM composition or ECM rigidity remains unknown.
Here, we have generated a fully functional, extracellularly tagged α5-integrin subunit and coupled it to the RUSH system to explore context-dependent traffic of newly synthesized fibronectin-binding α5β1-integrin to the plasma membrane. We demonstrate that 1) delivery of newly synthesized integrins to the plasma membrane is faster than previously thought; 2) their polarized delivery and ECM-ligand-dependent recruitment to FAs facilitate dynamic cell responses to ECM cues providing an additional, thus far unrecognized level of integrin regulation; 3) a small subset of integrin α5 is delivered in a high-mannose glycosylation state to the plasma membrane via Golgi-bypass trafficking. These findings are the first demonstration of biosynthetic integrin traffic influencing FA maintenance and cell dynamics.

# RESULTS

## Molecular simulation-guided generation of RUSH-α5

We employed the RUSH (retention using selective hooks) system¹⁶, which has been previously used to synchronize and study post-Golgi anterograde trafficking of a variety of cargos¹⁶⁻¹⁸, to control integrin retention and release from the ER (Extended Data Fig. 1). Using this method, we explored, for the first time, the context-dependent traffic of newly synthesized integrins and its implications in cell adhesion and dynamics in real time. Integrin α5β1 is the main fibronectin receptor in many cell types and has been widely studied. Thus far, integrin α5β1 has been tagged on the C-terminal tail, potentially interfering with some established protein-protein interactions¹⁹,²⁰. To identify a suitable alternative tagging site on the receptor’s ectodomain, we examined the published crystal structure of the integrin α5β1 headpiece²¹. Given that the N-terminus of the α5 polypeptide is localized between the two integrin subunits, away from the fibronectin ligand binding site, we inserted the IL-2 signal peptide, the streptavidin binding peptide (SBP) and enhanced green fluorescent protein (EGFP) in this region to generate an SBP-EGFP-integrin α5 construct (henceforth referred to as RUSH-α5) (Fig. 1a, Movie 1, Extended Data Fig. 1). Computational modeling suggested that while the flexible EGFP C-terminal region (plus linker) is just long enough to allow direct contact between EGFP and the fibronectin ligand binding site (Extended Data Fig. 2; Movie 2 and 3), EGFP is stably positioned and cannot be displaced within the range of normal physical force. Atomistic simulations were consistent with these observations (Movies 4-6; see methods for details). Thus, EGFP tagging of the integrin α5 ectodomain does not interfere with fibronectin binding or α5β1 subunit heterodimerization (Extended Data Fig. 2, and Movies 1-6). In cells, RUSH-α5 was retained in the ER when co-expressed with an ER-hook protein composed of the ER-retrieval motif KDEL fused to streptavidin (Fig. 1b) and released upon biotin addition to be transported to the plasma membrane (Fig. 1b; Movie 7; Extended Data Fig. 1 and Extended Data Fig. 3a).

## ECM control of integrin delivery to the plasma membrane

In cells, integrin β1 subunits are produced in excess and are transported to the plasma membrane only upon heterodimerization with newly synthesized α-integrins ¹¹⁻¹³. To investigate the ability of RUSH-α5 to form functional heterodimers with the β1 subunit, we
performed GFP-pulldown in cells co-expressing RUSH-α5 and the ER-hook (ER-hook always expressed with RUSH-α5 in all experiments) with and without biotin addition. The EGFP-tagged RUSH-α5 precipitated endogenous β1 integrin. RUSH-α5 interacted with the immature integrin β1 (faster migrating lower band; Extended Data Fig. 3b) before release from the ER (0 min biotin), and progressively with the mature integrin β1 after release from the ER following biotin addition (slower migrating upper band; Extended Data Fig. 3b). Importantly, RUSH-α5, when released from the ER (60 min biotin), localized to fibrillar and FA-like structures on fibronectin that were positive for active integrin α5 (detected with active integrin α5 conformation-specific SNAKA51 antibody) (Fig. 1c). In contrast, RUSH-α5 displayed a clearly reduced localization to adhesion structures along with a more diffuse localization pattern in cells plated on collagen and treated with biotin (Fig. 1c). The ECM ligand did not dramatically influence integrin heterodimer maturation (increasing ratio of mature to immature integrin β1) as it was not significantly faster on fibronectin than on collagen 20 and 40 min after biotin addition (Extended Data Fig. 3 c-d).

Our real-time measurements revealed, however, an interesting and unexpected feature of integrin maturation and delivery. Metabolic labeling studies have indicated receptor maturation kinetics exceeding 1 hour for the total integrin β1 cellular pool<sup>11,14</sup>. These pulse-labelling methods analyze synthesis, folding and secretion of integrins, and cannot be directly compared with the RUSH-method. However, our data reveals that the secretion and maturation of the α5β1 integrin maturation can be detected here already after 20 min of integrin release from the ER (Extended Data Fig. 3c-d). Taken together, these data indicate that RUSH-α5 forms a functional heterodimer with endogenous integrin β1, undergoes ligand-specific activation on fibronectin, and the dynamics and kinetics of the newly synthesized integrin can be analyzed using this method.

To explore the kinetics of the biosynthetic delivery of integrins in more detail, we performed time-lapse imaging of RUSH-α5 release in cells plated on collagen and fibronectin, comparing it with the dynamics of a co-expressed SBP- and mCherry-tagged control cargo protein<sup>16</sup> (glycosylphosphatidylinositol-anchored proteins (GPI-APs) cluster of differentiation 59 (CD59)) (henceforth called RUSH-CD59). On both ECMs, RUSH-α5 and RUSH-CD59 were localized to the ER in the absence of biotin, and following biotin addition were released and transported to the Golgi, residing there for approximately 15 min (Extended Data Fig. 4), in line with previous reports for CD59<sup>16,17</sup>. After 20 min, RUSH-α5 was predominantly localized on adhesion-like structures on fibronectin whereas on collagen it was diffusely distributed on the plasma membrane (Extended Data Fig. 4 and Movie 8). In contrast, the RUSH-CD59 construct behaved similarly on both fibronectin and collagen (Extended Data Fig. 4 and Movie 8), indicating that the observed differences in RUSH-α5 localization in cells on fibronectin and collagen were ligand-receptor specific. This was further validated by plating RUSH-α5 and RUSH-CD59 co-transfected cells on dual-coated micropatterns<sup>22</sup> with alternating lines of fibronectin and GFOGER (a synthetic collagen peptide with high affinity for collagen-binding integrins<sup>23</sup>). Following release (20 min biotin), RUSH-α5 localized predominantly to the cell edges, showing enriched clustering on fibronectin-coated lines, compared to GFOGER-coated lines (Fig. 1d). In contrast, RUSH-CD59 localization was
independent of ligand coating. This implies, that the integrin is largely delivered directly to the FN areas. However, as we are focusing only the ventral surface of cells, we cannot exclude the possibility that integrin α5 could be delivered first on the dorsal surface of the cells and then diffuse to the ventral surface into FAs. Next, we examined RUSH-α5 targeting to adhesions by co-expressing pmKate2-paxillin as a FA marker. RUSH-α5 was localized to pmKate2-paxillin-positive adhesions as early as 20 min following release with biotin, and this localization increased over time (Fig. 2a, b). On collagen, RUSH-α5 localization to adhesions was significantly lower compared to fibronectin (Fig. 2a, b), with the increase in intensity at the later time point most likely reflecting the increased presence of the diffused receptor on the membrane. We then replaced the EGFP in RUSH-α5 with pHluorin, a pH sensitive GFP <sup>24</sup>, to image RUSH-α5 cell surface delivery. The pHluorin GFP is non-fluorescent in acidic secretory vesicles and becomes fluorescent upon cell surface exposure. To detect the localization of RUSH-α5 secretion, we prevented receptor diffusion after ventral plasma membrane delivery by employing the selective protein immobilization (SPI) method <sup>17</sup>. The ECM proteins were coated alongside anti-GFP antibodies, which bind to the GFP moiety in RUSH-α5 and trap the receptor when it is delivered to the cell surface. In cells co-transfected with RUSH-α5-pHluorin and paxillin-mScarlet, we observed a specific increase in pHluorin signal in FAs over time (Extended Data Fig. 5). Moreover, TIRF live imaging of the RUSH-α5-pHluorin showed fluorescent flashes corresponding to RUSH-α5 surface delivery. These exocytosis events were significantly closer to FAs compared to those of randomly generated points (Fig. 2c and movie 9). These results indicate that RUSH-α5 is predominantly delivered to FA sites on fibronectin. In addition, it is possible that some of the integrin is transported elsewhere on the plasma membrane and rapidly diffuses to the fibronectin-specific adhesions further contributing to FA formation by the newly synthesized integrin.

## Polarized delivery of new integrin to the cell protruding edge

We then investigated if the localization of newly synthesized integrins is polarized. First, we plated RUSH-α5 transfected cells on 9 µm-wide collagen or fibronectin micropatterned lines (in combination with the GFP-SPI method to prevent RUSH-α5 diffusion after delivery), shown previously to support front-rear cell polarity of integrins<sup>25</sup>. Time-lapse imaging revealed a significant increase in RUSH-α5 intensity on fibronectin over time preferentially at the protruding edge of the cell (region of interest 1; ROI1) (Fig. 3a,b, Movie 10) whereas, on collagen lines, the difference in RUSH-α5 localization between the two cell edges was modest and apparent only at later time points (Fig. 3a, c, Movie 11). In line with these data, RUSH-α5 delivery was also polarized on fibronectin in unconstrained cells, occurring significantly more in protruding regions, whereas on collagen RUSH-α5 intensity increased both in retracting and protruding regions (protruding or retracting areas of the cells defined based on spatiotemporal track maps<sup>26</sup>, generated from paxillin images) (Fig. 3d, e). These data indicate that the plasma membrane delivery of newly synthesized integrins is sensitive to ECM ligand engagement and the cell front-rear polarity.

## Rapid, adhesion-dependent delivery of RUSH-α5
While RUSH-α5 was predominantly trafficked to the plasma membrane via the Golgi complex conventional secretion pathway, a process that takes more than 20 min, surprisingly, some RUSH-α5-positive vesicles were evident earlier (around 10 min) within the vicinity of the plasma membrane (Fig. 4a, Movie 12). This was also apparent using TIRF live imaging where the RUSH-α5 signal was detected at the cell-ECM interface already at 13 min post release. After 15 mins surface delivery was clearly polarized to the cell leading-edge protruding area (Fig. 4b, Movie 13). Moreover, using pHluorin-RUSH-α5, we were able to observe exocytosis events from 10 min after release (Fig. 2c, Movie 9). This unexpected early plasma membrane delivered integrin localized to fibronectin line micropatterns unlike the RUSH-CD59 construct (Fig. 4c). Cell surface delivery of RUSH-α5 was also detected with flow cytometry 15 min after biotin addition with a steady increase up to 1h (Fig. 4d). Live imaging of RUSH-α5 together with an ER marker (ERoxBFP) revealed RUSH-α5 puncta outside of the ER and being trafficked in close proximity to FAs at very early time points (Movie 12). High resolution confocal imaging of cells transfected with RUSH-α5 after 10 min of release showed no obvious overlap between RUSH-α5 vesicles and Golgi or ER makers (Extended Data Fig. 6). These observations suggest that at least some of the RUSH-α5 is delivered to the plasma membrane without going through the Golgi apparatus.

Integrin rapid delivery bypasses conventional Golgi secretion

These data prompted us to hypothesize that a small proportion of integrins could undergo unconventional protein secretion (UPS), a process where secretory proteins are transported from the ER to the plasma membrane without entering the Golgi complex <sup>27</sup>. Conventional Golgi secretion is inhibited by Golgicide A <sup>28</sup>. Therefore, we treated cells with Golgicide A to investigate the relative contribution of the Golgi complex to RUSH-α5 trafficking. As expected, the delivery of the majority of RUSH-α5, 25 min post-biotin addition, was significantly inhibited by Golgicide A (Fig. 5a, b). However, both control and Golgicide A-treated cells showed a small initial increase in RUSH-α5 recruitment to protruding areas of the cell and to adhesions 15 min after release (Fig. 5a, b), suggesting that a small fraction of newly synthesized integrins may be secreted to FAs via a mechanism that bypasses conventional Golgi secretion.

In normal cells, high-mannose-type glycans are predominantly localized to the ER, and undergo enzymatic processing and maturation into complex glycoforms in the Golgi during conventional secretion <sup>29</sup>. However, several cancer types and cancer patient samples show significantly increased proportions of high-mannose glycans on the cell surface <sup>30,31</sup> and unbiased probing with high-mannose binding lectins have shown that integrins are abundant in this receptor pool <sup>32</sup>. This prompted us to investigate the presence of high-mannose glycans at the cell surface. We used the fluorescently labeled *Pseudomonas fluorescens* Pf0-1lectin (PFL) which specifically binds to high-mannose glycans <sup>33</sup> to stain the surface of RUSH-α5 transfected cells before and after release (1h). We observed increased PFL staining after release, indicating delivery of high-mannose proteins to the cell surface (Extended Data Fig. 7a). To explore this further, we took an orthogonal approach and labelled the surface of RUSH-α5 cells after biotin addition (1h release) with a high-mannose specific antibody-like lectibody
molecule<sup>32,34</sup> and subsequently performed lectibody pulldowns. We detected RUSH-α5 in these pulldowns, indicative of high-mannose α5-integrin expression on the cell surface (Extended Data Fig. 7b).

While cell surface high-mannose bearing integrins have been reported in cancer cells<sup>30,31</sup>, insights into the relevance of Golgi bypass has been limited to the integrin αPS1 subunit during Drosophila follicle epithelium development, stimulated by mechanical stress<sup>35</sup>. This prompted us to investigate whether early secretion of newly synthetized integrins in mammalian cells is linked to cell adhesion and adhesion-induced mechanics. First, we explored whether pre-existing endogenous integrin α5 adhesions are involved. We knocked-out endogenous integrin α5 (ITGA5) (Extended Data Fig. 8a-d) and compared RUSH-α5 release in wild-type (WT) and knockout (KO) cells after 15 and 60 min biotin addition (note that U2OS cells have other fibronectin-binding integrins in addition to integrin α5 and thus the KO cells adhere to fibronectin similarly to control cells). RUSH-α5 delivery to the cell surface was comparable between WT and ITGA5 KO cells (Extended Data Fig. 9a). Thus, the early secretion of newly synthesized integrin α5 is not dependent on the localization of the endogenous protein already at the cell surface. However, the early delivery of RUSH-α5 to the plasma membrane was dependent on cell-ECM adhesion, as we did not detect RUSH-α5 at the cell surface after 15 min biotin addition in suspension cells (Extended Data Fig. 9b). Adhesion was not required for slower integrin secretion (60 min biotin) via the conventional pathway (Extended Data Fig. 9b). These data are consistent with cell adhesion and perhaps active spreading/protrusions acting as necessary triggers for early secretion of integrin α5 to the cell surface. Even though the early delivery of RUSH-α5 was not, as such, dependent on endogenous integrin α5, polarized RUSH-α5 localization to cell protruding areas was significantly higher in WT compared to ITGA5 KO cells (Extended Data Fig. 9c), suggesting that polarized localization of the newly synthesized integrin is orchestrated by existing adhesions and contributes to rapid alterations in cell shape.

Unconventional RUSH-α5 protein secretion is dependent on the integrin-tail PDZ motif

The two mammalian homologues GRASP55 and GRASP65 mediate UPS of transmembrane proteins via PDZ domain-mediated interactions with cargo proteins<sup>36,37</sup>. The integrin α5 cytoplasmic domain harbors two distinct PDZ-binding motifs: a classical C-terminal Ser-Asp-Ala (SDA) sequence<sup>38</sup> and a non-canonical motif generated by two prolines (PP), which induce an internal β-hairpin that functions as a PDZ-recognition motif<sup>19</sup>. To test whether integrin α5 associates with GRASPs, we performed GFP pulldowns in cells expressing RUSH-α5. In addition to integrin β1, we detected endogenous GRASP65 co-precipitating with RUSH-α5 (Fig. 5c). Further pulldown experiments with biotinylated peptides corresponding to the C-terminal part of the integrin α5 WT tail or integrin α5 tails with mutations in the non-canonical PDZ-binding motif (PPAA peptide) or deletion of the canonical PDZ-binding motif (ΔSDA) (Fig. 5d,e) indicated that GRASP65 associates with integrin α5 via the SDA sequence (Fig. 5e), in accordance with the ability of GRASP65 to facilitate UPS of ER-resident cargo containing PDZ-binding motifs and regulate N-linked glycosylation in the ER<sup>39</sup>. However, no direct interaction between recombinant GRASP65 and integrin α5 was observed, by GST pulldown (Extended Data Fig. 10a) or ELISA (Extended Data Fig. 10b). This indicates that the
association between integrin α5 and GRASP65 is indirect, requires additional co-factors or is
sensitive to post-translational modifications.

We then explored the involvement of GRASP and the integrin tail in the early delivery
of RUSH-α5. siRNA-mediated silencing of GRASP65 and GRASP55 (Extended Data Fig.
11a) inhibited RUSH-α5 delivery to fibronectin spots on dual-coated (FN and GFOGER)
micropatterns at 10-15 min post biotin addition (Extended Data Fig. 11b-c). However, the
recruitment of RUSH-α5 to fibronectin dots remained low 25 min after release, possibly due to
GRASP silencing interfering with Golgi function ⁴⁰. Furthermore, GRASP depletion has been
shown to downregulate integrin α5β1 protein levels and could also affect the lifetime of our
exogenous construct ⁴¹. To overcome these complications, we generated a RUSH-α5 construct
lacking the integrin α5 GRASP65-binding SDA sequence (RUSH-α5-ΔSDA) and performed
time-lapse imaging. RUSH-α5-ΔSDA recruitment to adhesions was delayed compared to cells
expressing full-length RUSH-α5 (25 min versus 5 min) (Fig. 5f). Even after 45 minutes,
RUSH-α5-ΔSDA accumulation in adhesions was diminished, consistent with unconventional
secretion accounting for a small part of the overall integrin biosynthetic delivery in cells.

New integrins are delivered to focal adhesions to drive adhesion growth

The endo/exocytic traffic of cell surface integrins controls FA dynamics, size and distribution
in cells ⁵,⁴²⁻⁴⁴, however the role of integrin secretion remains to be explored. CD59, along with
several other cargo proteins, undergo anterograde post-Golgi traffic to secretion hotspots
adjacent to but discrete from FAs ¹⁷. We employed dual color TIRF imaging of RUSH-α5 and
pmKate2-Paxillin in cells plated on fibronectin to determine whether this is also the case for
integrins. SPI revealed that RUSH-α5 is recruited directly to FAs rather than to adjacent
hotspots. When initiating a new adhesion, RUSH-α5 was initially delivered to the most distal
area (Area 1) of FAs (closest to the cell periphery) after which it gradually accumulated along
the growing adhesion towards the cell center (Fig. 6a-b and Extended Data Fig. 12a). However,
when delivered to an existing and elongated FA, RUSH-α5 was equally localized all along the
FA (Fig. 6c and Extended Data Fig. 12b).

To consider the effects of increased integrin delivery on adhesions, we employed a
molecular clutch model previously developed to simulate mechanosensitive growth of
adhesions <sup>45</sup>. In this model, talin-mediated mechanosensing (through force-induced unfolding)
leads to adhesion growth, modeled as an increase in integrin density (Fig. 6d). To understand
the effect of integrin delivery, we reasoned that an increased delivery would result in a higher
availability of integrins to be incorporated into adhesions. Thus, we modeled integrin delivery
by tuning the parameter that sets the increase in integrin density that occurs upon talin
unfolding (d<sub>add</sub>). Running the model with a base set of parameters taken from previous work
(Extended Data Table 1), modifying only the d<sub>add</sub> parameter, and running the simulation as a
function of time, indicated that delivery of new receptors is predicted to increase adhesion
growth (Fig. 6e). In concordance with the model, release of RUSH-α5 significantly increased
adhesion area on fibronectin, with no significant effect on collagen (Fig. 6f). Adhesion growth
supported by RUSH-α5 release was also apparent in the ITGA5 KO cells (Fig. 6g), indicating
that an increased number of clutches translates to larger adhesions also when a new type of
integrin heterodimer is introduced to the cell surface. The increase in adhesion size correlated
with enhanced cell spreading in WT and ITGA5 KO cells transfected with RUSH-α5 (Fig. 6h)
and to increased cell dynamics with longer cell protrusions extended in cells plated on
fibronectin compared to collagen following biotin addition (Fig. 6i, Extended Data Fig. 12c).
Taken together, these data indicate that newly synthesized integrin α5 is rapidly localized to
FAs, contributing to adhesion growth towards the proximal end and facilitating cell protrusion
in a spatially defined and ligand-dependent manner.
DISCUSSION

Our findings demonstrate that cell adhesion and polarized cell dynamics can be steered by
targeted delivery of newly synthesized integrins to the plasma membrane, and that this can
occur rapidly, in a localized manner, bypassing the Golgi complex (Fig. 6j). The biosynthetic
trafficking route of integrins has not been previously investigated spatiotemporally in cells or
associated with dynamic regulation of cell morphology and integrin adhesions. Earlier
biochemical labelling studies analyzed synthesis, folding and secretion of integrins but were
not amendable for imaging of their delivery to the plasma membrane. Therefore, the
contribution of newly synthesized to cell dynamics has remained uninvestigated. Our findings
place integrin delivery via the biosynthetic pathway on par with well-established pathways that
control integrin dynamics on the plasma membrane, namely endocytosis and recycling.

Motor-clutch-based modelling has been employed to understand many fundamental aspects of cell dynamics in response to ECM ligands and matrix rigidity <sup>45-47</sup>. Here, the model predicted time-dependent growth of adhesions in response to delivery of an increased number of integrins. These matched with our experimental data revealing increased adhesion growth and generation of cell protrusions in response to RUSH-α5 release from the ER. These data further underline that biosynthetic delivery of integrins is an important and thus far underestimated contributor to cell responses to ECM mechanics and composition.

Many open questions remain regarding the biological significance of integrin delivery via the biosynthetic route and the mechanisms governing this process. We find that GRASP65 is required for the rapid unconventional secretion of RUSH-α5. The GRASP proteins are localized to the Golgi and are classically associated with Golgi stacking. However, they are also involved in stress-induced UPS <sup>48</sup>. Our data imply that integrin UPS is adhesion-dependent and seems to be always linked with cell spreading. Therefore, it is tempting to speculate that local mechanical stress could be a signal for one pool of integrins to rapidly exit the ER and be delivered to the proximal plasma membrane. However, this remains to be investigated. While proteins can undergo N-glycosylation in the ER <sup>39</sup>, it is plausible that the glycans of the early secreted pool of integrins are not fully processed but rather remain in an immature, high-mannose state. N-glycan chains support integrin headpiece opening (activation) and increase integrin-ligand binding affinity <sup>49</sup>. However, high-mannose integrins are also functional in ECM engagement <sup>49</sup>, consistent with the early delivery of RUSH-α5 supporting cell protrusions and adhesion growth. The presence of high-mannose glycosylated integrins at the cell surface has been so far only demonstrated in cancer cells <sup>32,50</sup>. Moreover, we showed integrin delivery via unconventional secretion only in a cancer cell line model. Whether this is also relevant to normal cells or is specific to cancer cells remains to be determined.

Our data indicate that cells sense their underlying ECM and have the ability to deliver new
integrins in a polarized manner in response to receptor-specific ECM ligands. This is an
exciting observation that might be particularly relevant in the context of development, tissue
patterning and in wound healing where directed cell migration is paramount <sup>51</sup>. Thus far, we
have explored this process and its regulation in the context of the fibronectin-binding integrin
α5β1. Existing data suggest that for all β1-integrins the β1-subunit is produced in excess and
localizes to the ER where it homodimerizes with an α-subunit to become transported to the plasma membrane <sup>11</sup>. Integrin α-subunits harbor distinct cytoplasmic domains, and only a subset has a putative GRASP-binding PDZ motif <sup>52</sup>. In the future it will be interesting to explore whether integrin UPS is applicable only to a subset of integrins and whether these are perhaps biologically differentially employed in processes requiring a rapid and dynamic cell response.

The field of cell-ECM adhesion is well-established and there is a wide consensus that endocytosis and recycling of integrins from and to the plasma membrane are essential regulators of cell dynamics, migration and invasion <sup>53-57</sup>. Dysregulation of many of these integrin trafficking regulators is also linked to cancer progression <sup>55</sup>. We find here that delivery of fresh integrins, along the biosynthetic pathway, also operates to determine cell dynamics. Thus, mechanisms regulating integrin secretion are likely to be intertwined with established integrin trafficking pathways in previously unappreciated ways and most likely these mechanisms are relevant to human disease and operate alternately or even simultaneously endowing cells with greater plasticity to adapt to dynamic alterations in their extracellular environments.
# Materials and Methods

## Cell culture

CHO cells were grown in Ham's F12 supplemented with 10% fetal calf serum (FCS) and 1% L-glutamine. U2OS cells were grown in Dulbecco's Modified Eagle's medium (DMEM, Sigma D5796) supplemented with 10% fetal calf serum (FCS, Sigma F7524), 1% L-glutamine (Sigma G7513) and 1% penicillin/streptomycin (Sigma P0781). Cells were routinely tested for mycoplasma contamination.

## CRIPSR/Cas9 ITGA5 KO clone generation

The U2OS ITGA5 KO clone was generated by CRISPR-Cas9 in Finnish Genome Editing Unit (Sanger's gRNA Library; Finnish Genome Editing Unit supported by HiLIFE and the Faculty of Medicine, University of Helsinki, and Biocenter Finland). U2OS cells were transfected with one gRNA and further grown as single cell clones. The KO efficiency was assessed by Western Blot prior to TOPO cloning and Sanger sequencing to validate the KO in both alleles of the clone using the following primers: forward 5'-GGATTTGGCTTGGGAGGAGAGTATATAG-3' and reverse 5'- CCAGTCCCTCCCTGAATTTCAC-3'. One clone from the procedure in which the KO was not efficient was used as a control WT clone.

## Plasmids

ERoxBFP was purchased from Addgene (68126) and pmKate2-paxillin from Evrogen (FP323). pEGFP-GRASP65 was a kind gift from Martin Lowe (University of Manchester, UK) ⁵⁸. Streptavidin-KDEL_SBP-mCherry-CD59 was generated as previously described ¹⁷. Streptavidin-KDEL_SBP-EGFP-ITGA5 was generated by PCR amplification of human ITGA5 without its signal peptide using Integrin-α5-EGFP template from Patrick Caswell (University of Manchester, UK) and the following PCR primers: forward 5'-AATTggccggccgTTCAACTTAGACGCGGAGGC-3' and reverse 5'-AACCttaattaatcaGGCATCAGAGGTGGCTGG-3'. The PCR fragment was then subcloned in the RUSH plasmid Streptavidin-KDEL_ss-SBP-EGFP ¹⁶ using FseI and PacI restriction enzymes. The EGFP was replaced by pHluorin to generate Streptavidin-KDEL_SBP-pHluorin-ITGA5 using FseI and SbfI restriction sites. The hook (streptavidin-KDEL) allows anchoring of the SBP-tagged reporter (integrin α5 and CD59) in the ER in the absence of biotin due to streptavidin-SBP interaction.

## Transfection

Plasmids of interest were transfected using Lipofectamine 3000, Lipofectamine 2000 (Thermo Fisher Scientific, 11668019 and L3000001, respectively) or jetPRIME® (Polyplus transfection, 101000027) according to the manufacturer's instructions. Protein downregulation was carried out with Lipofectamine siRNA Max or Lipofectamine 3000 (ThermoFisher Scientific, 13778075 and L3000001, respectively) according to manufacturer's instructions. The siRNA used as control (siCTRL) was Allstars negative control siRNA (Qiagen, 1027281). GRASP65 and GRASP55 were downregulated with Flexitube siRNAs (Qiagen, GS64689 and GS26003 respectively) or custom ordered siRNA oligonucleotides (GRASP65 target sequence: AAG-
GCA-CUA-CUG-AAA-GCC-AAU and GRASP55 target sequence: AAC-UGU-CGA-GAA-GUG-AUU-AUU, Qiagen).

## RUSH-α5 transfection and release

Cells grown to 25 % confluence were used for transfection. For a 6 cm dish $1 \times 10^5$ cells were transfected with 10 µg RUSH-α5 using Lipofectamine 3000. The cells were from this point grown in medium containing 1-2.5 µg/ml streptavidin (Sigma-Aldrich, S4762) to block biotin in the media and transfection reagents. The day after transfection, cells were used for experiments, either directly by releasing the RUSH or detaching the cells with trypsin and seeding the cells beforehand on appropriate surfaces for imaging experiments as described below. The release of the RUSH-α5 or RUSH-CD59 from the ER-hook was induced by replacing the medium with biotin-supplemented medium (3 mM of D-biotin, Sigma-Aldrich, B4501) for the indicated times.

## Immunoprecipitations and immunoblotting

### GFP pulldown:

CHO or U2OS cells expressing GFP-tagged (RUSH constructs) proteins (three 10 cm dishes per condition or one 10 cm dish, 80 % confluence, for GFP-control construct, due to differences in expression efficiency) were washed with cold phosphate buffered saline (PBS Sigma D8537-500ML), harvested in PBS and pelleted. The cell pellet was resuspended in 200 µl of IP-lysis buffer (40 mM Hepes-NaOH, 75 mM NaCl, 2 mM EDTA, 1 % NP40, protease and phosphatase inhibitors) and incubated at 4 °C for 30 min, followed by centrifugation (10,000 xg for 10 min, 4 °C). 20 µl of the supernatant was kept aside as the lysate control. The remainder of the supernatant was incubated with GFP-Trap beads (ChromoTek; gtak-20), for 55 min at 4 °C.

### AvFc lectibody pulldown:

Two 10 cm dishes containing 80% confluent U2OS cells expressing RUSH-α5 were incubated with 3 mM biotin at 37 °C for 1 h and then with 5 µg AvFc lectibody ³² in PBS on ice for 1h. The cells were washed three times with PBS before addition of lysis buffer (1.5% Octylglycoside, 1% NP-40, 0.5% BSA, 1 mM EDTA, protease and phosphatase inhibitors). The lysates were spined down and the supernatant was collected. 20 µL of the supernatant was kept aside as the lysate control. 25 µL of protein G beads (SureBeads™ Protein G Magnetic Beads, BioRad) were added to the remaining lysates and incubated for 2 h at 4 °C.

### GST pulldown:

Pulldown with N-terminally biotinylated integrin α5 tail peptides (GenScript) was performed as follows: biotinylated peptides were incubated with streptavidin-conjugated Dynabeads (65001, ThermoFisher) for 30 min at room temperature followed by a 2 h incubation with supernatant (prepared in the same way as described above for GFP-immunoprecipitated samples) from EGFP-GRASP65-overexpressing CHO cells or with recombinant GST-GRASP65 or GST alone (GenScript), at 4°C.
## Immunoblotting:

Immunoprecipitated complexes were washed three times with wash-buffer (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% NP-40) or RIPA buffer (lectibody IP) (10 mM Tris-HCl pH8, 1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1% SodiumDeoxcholate, 0.1% SDS, 140 mM NaCl, protease and phosphatase inhibitors) and denatured for 10 min at 95 °C in reducing Laemmli buffer before SDS-PAGE analysis under denaturing conditions (4-20 % Mini-PROTEAN TGX Gels, Bio-Rad Laboratories, #561096). The proteins were then transferred to nitrocellulose membranes (Bio-Rad Laboratories, #1704158) before blocking with blocking buffer (Thermo, StartingBlock blocking, #37538) and PBS (1:1 ratio). The membranes were incubated with primary antibodies diluted in blocking buffer overnight at 4 °C. Following this step, membranes were washed three times with TBST and incubated with fluorophore-conjugated secondary antibodies (LI-COR) diluted (1:10,000) in blocking buffer at room temperature for 1 h. Membranes were scanned using BioRad ChemiDoc MP Gel Analyzer, an infrared imaging system (Odyssey; LI-COR Biosciences) or Azure Sapphire RGBNIR Biomolecular Imager. Primary antibodies used: Mouse anti-CD29 (integrin β1) (BD Biosciences, 610468), rabbit anti-GRASP55 (Sigma, HPA035274), rabbit anti-GRASP65 (Sigma, HPA056283), rabbit anti-GFP (abcam, ab290), mouse anti-GAPDH (Bioz, 5G4MaB6C5), rabbit anti-GST tag (Invitrogen, A-5800). Secondary antibodies used: IRDye® 800CW Donkey anti-Mouse IgG (LICOR, 926-32212), IRDye® 800CW Donkey anti-Rabbit IgG (LICOR, 926-32213), IRDye® 680LT Donkey anti-Mouse IgG (LICOR, 926-68022) and IRDye® 680LT Donkey anti-Rabbit IgG (LICOR, 926-68023), diluted 1:10,000 in odyssey blocking buffer (LI-COR, 927-40000).

## ELISA

Wells of a Nunc Maxisorp 96-well plate (Sigma-Aldrich) were first incubated with GST or GST-GRASP65 (0-1-2-5 and 10 µg in 50 µL) overnight at 4 °C. The wells were then washed with HMN-Tween 0.05% (HMN-T, HMN buffer: 20 mM Hepes, 100 mM NaCl, 5 mM MgCl2), blocked with 1% BSA in HMN-T 2 h on ice and washed with HMN-T. 0.2 µg of integrin β1 or integrin α5 tail peptides were incubated in the wells at 37°C for 2 h at room temperature. After two washes with HMN-T, HRP-streptavidin was added for 1 h at room temperature and the wells were washed again twice. 100 µL of TMB ELISA reagent was added and incubated for 10 min. The reaction was stopped with H2SO4 2M and the absorption at 450 nm was measured with a plate reader.

## Flow Cytometry

Cells were detached on ice, before or after biotin addition in the case of RUSH-α5 transfected cells, with enzyme-free cell dissociation buffer (Gibco 13150016). Pelleted cells were incubated with anti-GFP-AF647 antibody (1:150, Alexa Fluor 647 Mouse Anti-GFP Clone 1A12-6-18, BD Biosciences 565197), rabbit anti-ITGA5 (1:100, clone EPR7854, Abcam ab150361) or HiLyte Fluor 647-labeled PFL (0.5 µM) ³³), in Tyrodes Buffer (10 mM HEPES-NaOH at pH 7.5, 137 mM NaCl, 2.68 mM KCl, 1.7 mM MgCl2, 11.9 mM NaHCO3, 5 mM
glucose, 0.1 % BSA) for 40 min on ice. For PFL staining, cells were blocked with 5 µM unlabeled PFL before biotin addition. Cells were washed twice with Tyrodes Buffer. In the case of anti-ITGA5 staining, cells were then incubated with a donkey anti-Rabbit-AF647 (1:400, Invitrogen A31573) for 30 min on ice and washed twice. Cells were fixed for 10 min with 2 % PFA, resuspended in PBS and analyzed with LSRFortessa (BD Biosciences). Data analysis was performed with FlowJo software version 5. To quantify cell surface RUSH-α5 levels, the geometric mean of the anti-GFP AF647 antibody or PFL647 signal (surface labeling) was divided by the total GFP signal for each time point, and the value at T0 was subtracted from all time points.

## Live cell imaging

Cells were plated in imaging media (1:1 ratio of DMEM (Sigma, D5796) and FluoroBrite DMEM Media (ThermoFisher, A189670), supplemented with 20 mM HEPES (ThermoFisher, 15630080), 1 µg/ml streptavidin (Sigma-Aldrich, S4762) 10 % fetal calf serum (FCS, Sigma, F7524), 1 % L-glutamine (Sigma, G7513) and 1 % penicillin/streptomycin (Sigma, P0781) and allowed to spread for 2-4 h before imaging on fibronectin (Merck-Millipore, 341631) or collagen (Merck-Millipore, catalog number 08-115) (10 µg/ml) coated coverslips. Additional coating of 2.5 µg/ml Alpaca anti-GFP VHH nanobody (gt-250, Chromotek) was used in the indicated experiments. Time-lapse imaging was performed at 37 °C using Spinning-disk confocal 3i (Intelligent Imaging Innovations, 3i Inc) Marianas Spinning disk confocal microscope with a Yokogawa CSU-W1 scanner and back illuminated 10 MHz EMCCD camera (Photometrics Evolve) using 63x/1.4 oil objective. TIRF imaging was carried out using DeltaVision OMX with 60x/1.49 Olympus APO N TIRF objective. Conventional protein secretion was blocked (in indicated experiments) by incubating the cells with 10 µM Golgicide A (G0923, Sigma-Aldrich) 30 min prior to imaging. The release of the RUSH cargos was induced by removing the streptavidin supplemented media and addition of biotin supplemented imaging media (3mM of D-biotin, B4501 Sigma-Aldrich), by using a magnetic imaging chamber with L-shape tubing (CM-B25-1PB, Live Cell Instrument CO., LTD) during the live cell imaging experiments.

## Image analyses

As the intensity of RUSH-α5 varied from cell to cell based on transfection efficiency, relative RUSH-α5 recruitment was measured by normalizing the intensity at the indicated time point to the intensity before release (0 min) in the same measured region. Due to the low exposure time used for image acquisition of pmKate2-Paxillin (to reduce phototoxicity), de-noising of paxillin adhesions was carried out using the deep learning CARE2D network⁵⁹ in the ZeroCostDL4Mic platform⁶⁰, where 200 images were used to train the model prior to analysis. Paxillin adhesions were then segmented when needed. Images of paxillin adhesions were made binary and adhesions larger than 0.6 µm² were segmented and analyzed with the Analyze Particles tool in ImageJ⁶¹. The segmented adhesions were saved as regions of interest (ROIs) in the ROI manager and used to measure the intensity of RUSH-α5 within the paxillin adhesions. Spatiotemporal track maps of cells were generated based on the RUSH-α5 signal using the QuimP plugin²⁶ in ImageJ. Localization of RUSH-α5 to adhesions was studied by
drawing a ROI around the whole adhesion area based on the paxillin signal and then dividing
the ROI into 4 areas of equal size. RUSH-α5 intensity in the four areas relative to signal
intensity in the respective area 2.5 min prior to RUSH-α5 appearance in adhesions (determined
from time-lapse imaging) was measured. Adhesions close to the cell edge and with a minimum
lifetime of 15 min were analyzed and changes of RUSH-α5 intensity were plotted over time in
the indicated areas ranging from distal to proximal to the cell body. Detection of exocytosis
events from the TIRF RUSH-α5-pHluorin imaging was performed by dividing each frame by
the previous with the ImageJ image calculator function. Exocytosis spots were then segmented
by manual thresholding. The last frame was used to segment focal adhesions using trainable
WEKA segmentation⁶². The nearest distance between exocytosis events or randomly generated
spots and focal adhesions was measured with the Distance Analysis (DiAna) Fiji plugin ⁶³.

Immunofluorescence and image acquiring of fixed samples

Cells were plated on Ibidi 35 mm µ-dishes (80136) coated with 10 µg/ml collagen I (Merck-Millipore, catalog number 08-115) or fibronectin (Merck-Millipore, 341631). Samples were fixed for 10 min with 4 % PFA followed by permeabilization for 10 min with 0.1 % Triton X-100 in PBS. For ER staining with anti-PDI, samples were fixed in warm PEM buffer (8mM PIPES, 5mM EGTA and 2mM MgCl2) containing 0.5% glutaraldehyde and 0.25% Triton X-100 for 10 min at 37°C. Glutaraldehyde was then quenched with 0.1% NaBH4 for 7min. To block nonspecific binding of antibodies, cells were incubated in 10 % horse serum (HRS) for 1 h or in 1 M Glycine for 20 min at room temperature. Primary and secondary antibodies were diluted in 10 % HRS and incubated for 1 h at room temperature. Primary antibodies used: mouse anti-integrin α5 (clone SNAKA51) (MABT201, Millipore 1:500), mouse anti-GM130 (BD Biosciences, clone 35/GM130 1:1000), and mouse anti-PDI (Enzo, clone 1D3, 1:100). Secondary antibodies used: Alexa Fluor 555 anti-mouse (A32727, Thermo Fisher 1:400), Alexa Fluor 568 anti-mouse (A10037 Thermo Fisher, 1:400). F-actin was stained with Phalloidin-Atto 647N (65906, Sigma, 1:400), incubated together with secondary antibodies. Nuclei were stained with DAPI (D1306, Life Technologies 1:3000) for 10 min at room temperature after secondary antibody incubation. Samples were imaged using either A) 3i (Intelligent Imaging Innovations, 3i Inc) Marianas Spinning disk confocal microscope with a Yokogawa CSU-W1 scanner and Hamamatsu sCMOS Orca Flash 4.0 camera (Hamamatsu Photonics K.K.) or back illuminated 10 MHz EMCCD camera (Photometrics Evolve) using 63x/1.4 oil objective or 40x/1.1 water objective, B) Zeiss LSM780 laser scanning confocal microscope using 40x/1.2 water Zeiss C-Apochromat objective, or C) LSM880 laser scanning confocal microscope with AiryScan using 63x/1.4 oil Zeiss C Plan-Apochromat objective for high resolution imaging.

Micropatterns

Micropatterns were produced on glass coverslips as described in ⁶⁴. Cells were seeded in culture medium with fibronectin-depleted serum and allowed to spread on micropatterns for a minimum 2 h. For experiments with double coated micropatterns PLL(20)-g[3.5]-PEG(2)/PEG(3.4)-biotin (50 %) (SuSoS) was used to a recently developed method ²² allow coating of the non-micropatterned areas: the micropatterned areas were first coated with 50
µg/ml fibronectin and 5 µg/ml Bovine Serum (BSA) Alexa Fluor™ 647 conjugate followed by 30 min blocking with 3 % BSA, then the non-micropatterned areas were coated with 10 µg/ml GFOGER (Auspep) conjugated to streptavidin using the FastLink Streptavidin Labeling Kit (KA1556, Abnova) according to manufacturer's instruction.

## Constructing complete atomistic and coarse-grained models

We constructed simulation models that match the protein complexes studied in experiments as accurately as possible. To this end, we built both an atomistic and a coarse-grained Martini 3 ⁶⁵ model of the bound integrin construct as described here and in the next paragraph. For the fibronectin and antibody-bound integrin structure, we used the PDB id 7NWL, with the bound antibody removed. In this structure the transmembrane and intracellular domains of the integrin alpha and beta are not included. As the starting structure for EGFP, we used the PDB id 2Y0G, where we did not include the three chromatic residues (TYG) in 2Y0G as they are non-standard. To speed up the construction of the complete protein complex, these atomistic structures were individually coarse-grained to the Martini 3 representation with elastic networks and then put together in a single box (⁶⁶; https://github.com/marrink-lab/vermouth-martinize). The box was solvated with default Martini 3 water at 150 mM NaCl. In this coarse-grained representation, the EGFP C-terminus was pulled toward the integrin alpha N-terminus (constant rate at 1 nm ns⁻¹ with a harmonic force constant of 1000 kJ mol⁻¹ nm⁻²) to reach a structure where the termini were less than 1 nm apart. When this state was achieved, the coarse-grained structure (the fibronectin-integrin-EGFP complex) was backmapped to an atomistic representation by aligning the atomistic structures to the pulled coarse-grained system using the C-alpha backbone beads for reference. Finally, as the last few residues at the C-terminus of the EGFP (2Y0G) structure were missing (LGMDELYK), they were added together with the linker using the Modeller loop protocol ⁶⁷, i.e., the extended EGFP C-terminus was connected to the N-terminus of the integrin alpha subunit of 7NWL. This final atomistic model contained all sugars and bound ions from the crystal structures and was used as the starting structure in atomistic molecular dynamics simulations. Further, this final atomistic structure of the complete complex was coarse-grained to the Martini 3 representation, which was used as the starting structure in the construction of the complete coarse-grained simulation model (see the next paragraph).

## Constructing the production coarse-grained model

For the Martini 3 coarse-grained production simulations the sugars and manganese cations in the 7NWL structure were not taken into consideration. To maintain the folded structure, an elastic network was added using Martinize2 based on the ElNeDyn protocol ⁶⁶. Both intra- and intermolecular contacts were stabilized by the elastic network, but no harmonic bonds were added between EGFP and integrins/fibronectin. The elastic bonds in the linker residues were removed completely. The complex was solvated using insane (INSert membrANE) in default Martini 3 water with 150 mM NaCl ⁶⁸ with a minimum periodic distance of 4 nm.

## Simulations of the production coarse-grained models
To run the coarse-grained simulations, we used the GROMACS 2021.2 package⁶⁹. For energy minimization the steepest descent algorithm was used, and during equilibration the default Martini settings were employed, making use of a 1 fs time step up to the point that numerical stability was achieved⁷⁰. The Verlet cut-off scheme was used with a 1.1 nm cut-off for both the Coulombic (reaction-field) and van der Waals interactions. We used v-rescale for the thermostat at 300 K, coupling the protein and solvent in separate groups. Pressure coupling was initially performed using the Berendsen barostat⁷¹ for isotropic systems at 1 atm. The production runs made use of a 20 fs time step, where the pressure coupling was switched to Parrinello-Rahman⁷². For the pulling simulations, the pull code as implemented in GROMACS 2021.2 was used. The umbrella pulling method was employed to pull EGFP along a vector joining the center of mass of EGFP towards the fibronectin binding site. For each pulling simulation, a rate of 0.1 nm ns⁻¹ was used with a harmonic force constant of 1000 kJ mol⁻¹ nm⁻². The videos were made with the VMD movie maker plugin⁷³. The production runs spanned 80 ns for the binding site pulling and 3200 ns for the subsequent (non-biased) relaxation.

## Simulations of the atomistic models

To run the atomistic simulations, we used the GROMACS 2021.2 package⁶⁹. The protein was solvated in the presence of 150 mM of sodium chloride at 310 K temperature with a pressure of 1 atmosphere. The LINCS algorithm⁷⁴ was used to constrain the bond lengths in the system during simulations. The CHARMM36m force field⁷⁵ was used to derive the parameters for the protein and the ions. The CHARMM water model⁷⁶ was used to obtain parameters for the water molecules used to solvate the protein. The particle mesh Ewald technique⁷⁷ was used to calculate electrostatic interactions within the simulation system with a real-space cut-off of 1.2 nm. The protein structure was first energy-minimized and then subjected to a 100 ns equilibration. Ten independent simulations were then performed for generating the production runs. A time step of 4 fs was used for the simulations using the hydrogen mass partitioning method⁷⁸. For the pulling simulations, the pull code as implemented in GROMACS 2021.2 was used. The umbrella pulling method was employed to pull the EGFP along a vector joining the center of mass (COM) of EGFP towards the fibronectin binding site. For each pulling simulation, a rate of 0.1 nm/ns was used. The videos were made with the VMD movie maker plugin⁷³.

## Computational clutch model

The clutch model used considers how force transmitted from myosin motors to the substrate is applied to talin molecules and integrin-substrate bonds. Integrins bind and unbind from the substrate through binding rate $k_{on}$ and unbinding rate $k_{off}$, and talin folds and unfolds with folding and unfolding rates $k_{fold}$ and $k_{unfold}$. $K_{off}$, $k_{fold}$, and $k_{unfold}$ depend on force as previously described experimentally. Binding sites on the substrate are modeled explicitly, whereas integrins are modeled implicitly via a given integrin density $d_{int}$. Each time that talin unfolds an adhesion reinforcement event is assumed to happen, which is modeled as an increase in integrin density $d_{add}$. Model code and all parameters were taken from previous work⁴⁵. The only differences were the following:
* Our previous work considered that integrin density could both increase (when talin unfolds) and decrease (when integrins unbind from the substrate without talin unfolding). Here, we are only modeling adhesion growth, so we only consider growth.

* We set an upper limit to integrin density (3 times the initial value), to consider that adhesions only grow to a maximum size.

* We decreased the parameter $d_{add}$ to match the timescale of adhesion growth (to 0.01 or 0.005 integrins/µm²).

## Statistical analysis

GraphPad Prism software was used and the names and/or numbers of individual statistical tests, samples and data points are indicated in figure legends. Unless otherwise noted, all results are representative of three independent experiments and P values <0.05 are shown in graphs.
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**Acknowledgements:** We thank P. Laasola, J. Siivonen and M. Miihkinen for technical assistance and the Ivaska lab for scientific discussions. Prof. J. Norman, Prof. D. Odde and Dr S. Goodman are acknowledged for critical reading of the manuscript and Prof. B. Goud for hosting M.L.'s research visit at Institut Curie. The Cell Imaging and Cytometry Core (Turku Bioscience Centre, University of Turku) and the University of Helsinki Genome Biology Unit, both supported by Biocenter Finland, and the Euro-BioImaging Finnish Node (Turku Finland) are acknowledged for services, instrumentation, and expertise. We also acknowledge CSC – IT Center for Science for providing computing resources.

This study has been supported by the Finnish Cancer Institute (K. Albin Johansson Professorship J.I.), the Research Council of Finland project (325464 J.I., 331349, 335527 I.V., 338537 G.J.), and CoE in Biological Barrier Mechanics and Disease (346131 J.I., and 346135 I.V) funding, the Sigrid Juselius Foundation (J.I., I.V., G.J.), the Finnish Cancer Organization (J.I., G.J.), the Turku Doctoral Programme of Molecular Medicine (TuDMM; M.L.), Svenska Kulturfonden (M.L.), Victoriastiftelsen (M.L.), K. Albin Johanssons Stiftelse (M.L.), Marie Curie Skłodowska fellowship scheme (101033606 S.K.), Helsinki Institute of Life Science (HiLIFE) Fellow Program (I.V.), Human Frontier Science Program (RGP0059/2019, I.V.), the Åbo Akademi University Research Foundation (G.J., CoE CellMech), the Drug Discovery and Diagnostics strategic funding to Åbo Akademi University (G.J.), EMBO (ALTF 798-2021 M.M.), the Foundation for the Finnish Cancer Institute (M.M.), a Research Council of Finland postdoctoral research grant (343239, M.R.C), the Norwegian Cancer Society (N.M.P.), the Spanish Ministry of Science and Innovation (PID2022-142672NB-I00 to P.R.-C), the European Research Council (grant 101097753 MechanoSynth to P.R-C), the prize "ICREA Academia" for excellence in research (P.R.-C.). IBEC is a recipient of a Severo Ochoa Award of Excellence from the Spanish Ministry of Science and Innovation.

**Author contributions:** Conceptualization: J.I., M.L, S.M. Methodology: A.B., G.B., J.I., M.L, M.M, S.M., F.P., P.R-C., I.V. Formal Analysis: A.B., B.M.H.B., M.R.C., J.I., G.J., S.K., M.L, M.M. Investigation, B.M.H.B., M.R.C., J.I., S.K., M.L. N.M.P, L.M, M.M., P.R-C. Resources: A.B., G.B., G.J., F.P., I.V., N.M., Y.S. Visualization: A.B., H.H., M.L, M.M., Writing – Original Draft: J.I., M.L. Writing – review & editing: H.H., J.I., M.L., M.M., Supervision: A.B., J.I., S.M., I.V. Funding Acquisition: J.I., M.L., M.M.

**Competing interests:** The authors declare no competing interests.

**Data and materials availability:** The data supporting the findings of this study are available within the article and from the authors on reasonable request. Correspondence and material requests should be addressed to Johanna Ivaska (joivaska@utu.fi).

**Supplementary Materials:**

Extended Data Figs. S1 to S13

Extended Data Table 1

Movies S1 to S13
<figure><img src="image_2.png" /><figcaption>Figure 1. RUSH-α5 delivery to the plasma membrane is spatially regulated by the ECM.</figcaption></figure>

a) Model of RUSH-α5 (EGFP-integrin α5) - integrin-β1 heterodimer based on the structure of human α5β1 integrin bound to fibronectin (PDB: 7NWL) (see also Movie 1).
b) Representative immunofluorescence timelapse images of a U2OS cell expressing RUSH-α5 (SBP-EGFP-integrin α5) and plated on fibronectin ± biotin treatment (single greyscale images for the indicated time points are shown; see Movie 7).
c) Representative immunofluorescence images of RUSH-α5 and active integrin α5β1 (SNAKA51 antibody) in RUSH-α5-expressing U2OS cells plated on fibronectin or collagen + biotin (60 min). Greyscale single-channel images and merged images (white, colocalization; blue, nuclei (DAPI)) are shown.
d) Representative images of RUSH-α5 and RUSH-CD59 release in U2OS cells co-expressing both constructs and plated on dual-coated micropatterns (alternating fibronectin coating (cyan) and collagen-peptide (GFOGER) (non-fluorescent) lines). Nuclei (blue) are co-labeled. Intensity line profiles generated across the yellow line are displayed relative to the position of the fibronectin-coated micropattern lines. White insets represent regions of interest (ROIs) that are magnified for each channel. FN: fibronectin.
<figure><img src="image_3.png" /><figcaption>Figure 2. RUSH-α5 delivery to focal adhesions.</figcaption></figure>

a) Representative immunofluorescence images of U2OS cells co-expressing RUSH-α5 and pmKate2-Paxillin plated on fibronectin or collagen ± biotin treatment for the indicated times. Insets represent ROIs that are magnified and show paxillin-segmented adhesions (red outlines).

b) Quantification of the relative mean intensity of RUSH-α5 in segmented adhesions/cell ± biotin treatment for the indicated times. Data are mean ± SD; n = 64 cells on collagen, 50 cells on fibronectin, pooled from three independent experiments; One-way ANOVA, Holm-Sidak's multiple comparison test.

c) TIRF imaging of U2OS expressing RUSH-α5-pHluorin on fibronectin coated surface after biotin release at T=0. The arrows indicate exocytosis events. Exocytosis events were detected by performing a ratiometric analysis which consisted in
dividing each frame by the previous. All detected events before 19 min of release are indicated
in red. The graph indicates the distance between the exocytosis events and the nearest focal
adhesion segmented on the last frame of the movie (37 min after release), compared to the
distance of random dots with focal adhesions, showing that the localization to FAs is not
random. Individual measurements and the mean are represented. Unpaired T-test.
<figure><img src="image_4.png" /><figcaption>Figure 3. Polarized delivery of newly synthesized integrin to the cell protruding edge.</figcaption></figure>

a-c) RUSH-α5 intensity in U2OS cells plated on 9 µm-wide micropatterns coated with fibronectin and anti-GFP or collagen and anti-GFP ± biotin treatment for the indicated times was analyzed at both cell edges (the predominantly protruding edge was denoted ROI1 and the other edge ROI2). Representative intensity coded images (a) and quantification of RUSH-α5 release on fibronectin (b) and collagen (c) (normalized first to the total intensity of the cell and then to 0 min biotin) are shown. Data are mean ± SEM. d) Representative images and
spatiotemporal track maps of cell edge contours over time in U2OS cells expressing RUSH-α5 ± biotin treatment for the indicated times. Red insets represent protruding ROIs that are magnified. Blue insets represent retracting ROIs that are magnified. Spatiotemporal track maps: blue colors represent early time points and magenta colors represent late time points in the time-lapse series. e) Quantifications of RUSH-α5 intensity in ROIs (retracting or protruding areas determined from spatiotemporal track maps). Data are mean +/- SD. Scale bars: 20 µm. b, c) N = 33 cells on fibronectin and 38 cells on collagen, pooled from three independent experiments, Two-Way ANOVA, Sidak's multiple comparison test. e) N = 53 cells on collagen, 49 cells on fibronectin, pooled from three independent experiments, One-way ANOVA, Holm-Sidak's multiple comparisons test.
<figure><img src="image_5.png" /><figcaption>Figure 4. Early delivery of RUSH-α5 to the cell surface.</figcaption></figure>

a) Representative immunofluorescence images of U2OS cells co-expressing RUSH-α5 (green) and the ER marker ERoxBFP plated on fibronectin (10 µg/ml) ± biotin treatment for the indicated times. Arrows indicate rapidly budding RUSH-α5-positive vesicles adjacent to cell protrusions (≤ 15 min after release). (see also Movie 9).
b) TIRF imaging of RUSH-α5 after release (0 min). The polarized delivery to the cell surface at the protruding area can be observed from 15 min after release.
c) Representative images of RUSH-α5 and RUSH-CD59 release in U2OS cells co-expressing both constructs and plated on dual-coated micropatterns (alternating fibronectin coating (cyan) and collagen-peptide (GFOGER) (non-fluorescent) lines). Nuclei (blue) are co-labeled. Intensity line profiles generated across the yellow line are displayed relative to the position of the fibronectin-coated micropattern lines. White insets represent ROIs that are magnified for each channel. FN: fibronectin.
d) Flow cytometry analysis of cell surface RUSH-α5 levels (detected with the anti-GFP-AF647 antibody) in RUSH-α5-expressing U2OS cells ± biotin. Representative histograms and quantification of cell surface GFP (ratio of the geometric means of the surface signal divided by the total GFP signal, normalized by subtracting the 0 min value) are shown. Scale bars, 20 µm.
<figure><img src="image_6.png" /><figcaption>**Figure 5. Golgi bypass early delivery of RUSH-α5 is PDZ-motif dependent.**</figcaption></figure>

a,b) Quantification of relative RUSH-α5 recruitment to protruding areas (a) or adhesions (b) in U2OS cells expressing RUSH-α5 ± biotin treatment for the indicated times with or without Golgicide A (10 µM). Data are mean ± SD. c) Representative immunoblot of GFP-pulldowns from RUSH-α5 or GFP control transfected cells plated on fibronectin and probed for GFP and endogenous GRASP65. N= 3 independent experiments. d) Amino acid sequence of the integrin α5 tail highlighting the canonical PDZ-binding motif (SDA) and the two proline residues critical for the formation of the non-canonical PDZ-binding motif. The mutations of these sites used in our experiments are indicated below. e) Representative streptavidin pulldowns of the indicated biotinylated recombinant integrin peptides incubated with cell lysates collected from CHO cells overexpressing GFP-GRASP65. Representative immunoblots probed for GRASP65 (upper band, GFP-GRASP65; lower band, endogenous GRASP65) and GAPDH are shown. N = 3 independent experiments. f) Quantification of RUSH-α5 or RUSH-α5 ΔSDA recruitment to adhesions in U2OS cells ± biotin treatment for the indicated times.

All data are mean +/- SD. a,b) One-way Anova with Tukey's multiple comparison test for comparing time points, one way Anova with Šídák's multiple comparisons test for comparing untreated and Golgicide A. a) N = 26 cells RUSH-α5, N = 22 cell RUSH-α5 Golgicide A, pooled from three independent experiments. b) N = 24 cells RUSH-α5, N = 27 cell RUSH-α5 Golgicide A, pooled from three independent experiments. N=2 independent experiments. f) One sample t-test to compare time points with T=0, ordinary one-way Anova with Holm-Šídák's multiple comparisons test to compare RUSH-α5 and RUSH-α5 ΔSDA. N = 23 cells RUSH-α5, N = 23 cells RUSH-α5 ΔSDA, pooled from two independent experiments.
<figure><img src="image_7.png" /><figcaption>Figure 6. RUSH-α5 is delivered to the tip of adhesions and mediates adhesion growth.</figcaption></figure>

a-b) Representative immunofluorescence image of U2OS cells expressing RUSH-α5 and pmKate2-Paxillin (white, colocalization) plated on fibronectin (10 µg/ml) and anti-GFP (2.5 µg/ml; to trap cell surface RUSH-α5 at the point of delivery). Insets represent ROIs that are
magnified. ROI2 is a FA demarcated into four equal areas for analysis and is further magnified
in b. Scale bars 20 µm (whole cell image) and 0.5 µm (insets). c) Representative image of an
already established focal adhesion where RUSH-α5 delivery was quantified. Scale bar 0.5 µm.
d) Cartoon showing clutch model elements. Myosin motors pull on actin filaments with a speed
v. This applies force to a substrate via integrins and adaptor proteins (talin). The effect of force
regulates the unbinding rates from integrins to the substrate (koff) and the folding/unfolding
rates of talin (kfold/kunfold). When talin unfolds, adhesion reinforcement is assumed to happen,
which is modeled by an increase in integrin density with value dadd. Changes in integrin
availability are modeled by changing the parameter dadd. e) Model prediction of adhesion
growth with time for conditions in which integrin availability is low (dadd = 0.005
integrins/µm²) or high (dadd = 0.01 integrins/µm²). Adhesion growth (y-axis) is modeled
through integrin density, which is plotted normalized to the starting value. f) Quantification of
adhesion growth in U2OS cells expressing RUSH-α5 and plated on fibronectin or collagen ±
biotin treatment for the indicated times. Shown are the relative sums of segmented adhesion
area/cell at indicated time points. Data are mean ± SD. g) Quantification of adhesion growth in
WT and ITGA5 KO U2OS cells expressing RUSH-α5 ± biotin treatment for the indicated
times. Shown are the relative sums of segmented adhesion area/cell at indicated time points.
Data are mean ± SD. h) Quantification of cell spreading in WT and ITGA5 KO cells
expressing RUSH-α5 ± biotin treatment for the indicated times. Data are Mean ± SEM. i)
Quantification of the length of the longest protrusion (extending furthest from the initial plasma
membrane localization during imaging) formed per cell after 45 min of biotin. Data are mean ±
SEM. j) Schematic depiction of the regulation of cell dynamics by transport of integrins
through the biosynthetic pathway. Adhesion and cell spreading-dependent delivery of integrin
from the ER is detected rapidly after release in cell protrusions. Canonical Golgi-dependent
delivery is also polarized to cell protruding areas in an ECM-specific manner and contributes to
FA growth and cell protrusion.

(f, g and h) One-way ANOVA, Holm-Sidak's multiple comparison test. (f) N = 64 cells on collagen, 50 cells on fibronectin, pooled from three independent experiments. (g) 57 WT cells and 52 ITGA5 KO cells, (h) 59 WT cells and 55 ITGA5 KO cells, pooled from three independent experiments. (i) Mann-Whitney test, N = 55 cells on fibronectin, 66 cells on collagen, pooled from three independent experiments.