Spatiotemporal control of integrin-mediated cell adhesion to the extracellular matrix (ECM) is critical for physiological and pathological events in multicellular organisms, such as embryonic development, angiogenesis, platelet aggregation, leukocytes extravasation, and cancer cell metastatic dissemination. Regulation of integrin adhesive function and signaling relies on the modulation of both conformation and traffic. Indeed, integrins exist in a dynamic equilibrium between a bent/closed (inactive) and an extended/open (active) conformation, respectively endowed with low and high affinity for ECM ligands. Increasing evidence proves that, differently to what hypothesized in the past, detachment from the ECM and conformational inactivation are not mandatory for integrin to get endocytosed and trafficked. Specific transmembrane and cytosolic proteins involved in the control of ECM proteolytic fragment-bound active integrin internalization and recycling exist. In the complex masterplan that governs cell behavior, active integrin traffic is key to the turnover of ECM polymers and adhesion sites, the polarized secretion of endogenous ECM proteins and modifying enzymes, the propagation of motility and survival endosomal signals, and the control of cell metabolism.

Introduction

Metazoan cells attach to extracellular matrix (ECM) proteins via integrin αβ heterodimers that, through a series of cytosolic adaptor proteins, physically connect to the actin cytoskeleton and modulate the enzymatic activity of kinases, phosphatases, and small GTPases [1–3]. Overall, the complex protein network associated with integrin-based adhesion sites is known as the adhesome [2,3]. Twenty-four integrin heterodimers exist which promiscuously allow the interaction with hundreds of ECM proteins in different tissues and organs [4]. On the cellular surface, integrin receptors are in an allosteric equilibrium between a bent/inactive and an extended/active conformation that respectively interact at a low and high affinity with ligands [1]. The percentage of surface integrins adopting the extended/active conformation vary significantly depending on the adhesion and spreading degree of cell types, being for example ∼0.2% in poorly adherent K562 leukemia cells [5] or 10% in widely spread endothelial cells (ECs) [6]. Active integrin conformers are stabilized by the four-point-one, ezrin, radixin, moesin (FERM) domain of talin [7,8] and kindlin [8,9] adaptor proteins with a membrane proximal and distal Asn-Pro-X-Tyr (NPXY) motif in the cytodomain of integrin β subunits, respectively. Both talins and kindlins are required for integrin conformational activation, to which they seem to contribute differently by allowing the vinculin-mediated perception of mechanical forces (talins) and triggering biochemical signaling pathways (kindlins) [8], e.g. through paxillin and focal adhesion kinase (FAK) [10–12]. It has long been thought that the appearance of integrins and associated proteins allowed the evolutionary transition from unicellular to multicellular organisms [13]. Yet, over the last decade, independent studies proved that genes encoding for integrins [14,15] and most adhesome proteins, such as talins [15], but not kindlins [16], were already present in unicellular ancestors of animals that exploited them in the aggregation phase of their life cycle [13]. Hence, it is conceivable that the emergence of kindlins had been important for the development of multicellular organisms [16].

The dynamic control of integrin conformational activation is central in different physiological and pathological settings, such as tissue and organ development, platelet aggregation, leukocyte extravasation, autoimmune diseases, fibrosis, and cancer [1,17–19]. In these contexts, the small GTPase Rap1 is a main driver of integrin conformational activation. Indeed, talin exists in an autoinhibited state that is relieved by its direct [20–24] or indirect (e.g. through Rap1-interacting adapter molecule — RIAM) [25] binding to Rap1-GTP, allowing the FERM domain of talin to bind and promote integrin activation. Both the fulfillment of complex morphogenetic programs [26,27] as well as the normal functioning of platelets and leukocytes [1,25,28] rely on a fine spatiotemporal modulation of integrin activation by Rap1 and talin. In this scenario, chemoattractant ligands, e.g. C-X-C motif chemokine 12 (CXCL12) [29] or vascular endothelial growth factor-A (VEGF-A) [30], promote Rap1 activation via G protein coupled receptors (GPCRs) or tyrosine kinase receptors (TKRs) coupled to downstream Rap1 guanine nucleotide exchange factors (GEFs) [31]. Conversely, chemorepulsive ligands such as semaphorins (SEMAs) signal through the cytosolic GTPase activating protein (GAP) of Plexin receptors to inhibit Rap1 GTP loading [32–34] and integrin activation [17,18]. The evidence that both talin autoinhibition [27] and Plexin-mediated SEMA inhibition of Rap1 [17,34,35] are required for morphogenesis, in diverse experimental systems, supports the concept that conformational integrin inactivation is as crucial as activation for the accomplishment of complex shaping programs in animal tissues, organs and systems [17,36–39].

In addition to integrin conformational activation, cell-to-ECM adhesion contact dynamics critically relies on integrin traffic [19,40–43]. While it has long been thought that detachment from ECM ligands [44,45] and conformational inactivation [46] were necessary to allow integrin internalization, in the last decade, thanks to the availability of monoclonal antibodies detecting activation dependent epitopes [47], different laboratories showed that conformationally active integrins can also be endocytosed and trafficked under the control of molecular machineries and signaling pathways distinct from those regulating inactive integrin traffic. Here, we will review our current understanding of the rationale and the mechanisms by which cells traffic conformationally active integrins in both normal and cancer cells.

Active integrin traffic controls ECM adhesion site turnover and cell metabolism

Integrin binding to soluble monomeric ECM proteins, such as fibronectin (FN), results in traction force-dependent unmasking the otherwise cryptic protein–protein interaction sites that promote ECM polymerization in insoluble protein networks [48,49]. Polymerized FN act as a scaffold on which collagen I molecules are deposited to form fibrils and fibers via repeated cycles of integrin-mediated contraction and relaxation [50]. Subsequently, matrix metalloproteinases (MMPs) cleave ECM fibrils into fragments that get endocytosed and degraded, thus requiring the replenishment with freshly synthesized ECM molecules (Figure 1). Thus, ECM meshworks are unstable objects whose dynamic remodeling is crucial for tissue morphogenesis and healing as well as cancer cell invasion and dissemination [51]. Additionally, it was found, in keratinocytes, that microtubules anchored by cytoplasmic linker associated proteins (CLASPs) in proximity of ECM adhesions allow the targeted transport of secretory vesicles, which locally deliver membrane-type 1 MMP (MT1-MMP) [52] to degrade the ECM [53]. Moreover, it has been shown, in fibroblasts, that, upon MT1-MMP dependent cleavage [54], FN fragments bound to active α5β1 get endocytosed, trafficked and degraded into lysosomes [55]. As a result, in the absence of such constant FN synthesis, secretion and polymerization, FN fibrils disappear, being degraded [56]. Integrins have been reported to mediate the internalization and turnover of other ECM proteins, e.g. type I collagen [57] in fibroblasts, vitronectin [58] and laminin 111 [59] in cancer cells. Therefore, active integrins participate to the control of ECM turnover on the cell surface by mediating the endocytosis of MMP-cleaved ECM fragments in different cell types.

Active integrin internalization and trafficking pathways.

Figure 1.
Active integrin internalization and trafficking pathways.

Upon MMP-dependent cleavage of polymerized FN fibrils, the endocytic receptor NRP1 interacts on the cell surface with FN fragment-bound ubiquitinated α5β1 active-integrin that is internalized within early endosomes (EEs) through either Rab21, or Rab5, or Afr4 small GTPase, depending on the internalization site and cell type (see text and Figure 2). In EEs, active α5β1 integrin may activate different signaling pathways such as: (i) Rac1 to promote integrin-dependent cell spreading and migration on FN; (ii) FAK to suppress anoikis in normal cells or promote cancer cell anchorage-independent growth. Moreover, in Rab25 overexpressing ovarian cancer cells Arf4-dependent active α5β1 integrin is required for correct lysosome positioning and activation of mTORC1, whose kinase activity is also regulated by glucose levels. From the EE compartment, inactive integrins can move back to the plasma membrane via sorting nexin 17 (SNX17) and the retriever complex [124]. A fraction of FN fragment-bound active α5β1 integrins is sorted into multivesicular bodies (MVBs) and degraded into lysosomes; this degradative fate depends on α5β1 integrin ubiquitination and on endosomal sorting complexes required for transport (ESCRT) protein complex. Another fraction of FN fragment-bound active α5β1 integrin instead traffics to PGCs; it is still unclear if this latter step relies on deubiquitinase (DUB) and/or SNX activities. It is posited that, in PGCs, endocytosed old FN may be separated from active α5β1 integrin that would then be free and able to bind freshly synthesized FN originating from TGN cisternae in a PI4KB and AP-1A-dependent manner. Through a RAB11B-dependent endosomal recycling pathway, vesicles containing newly synthesized FN-bound active α5β1 integrins are next directed to the basolateral side of the cell surface, where the PTPRF/PPFIA1 complex support their docking, likely via PPFIA1-mediated interaction with the β1 integrin subunit cytodomain.

Figure 1.
Active integrin internalization and trafficking pathways.

Upon MMP-dependent cleavage of polymerized FN fibrils, the endocytic receptor NRP1 interacts on the cell surface with FN fragment-bound ubiquitinated α5β1 active-integrin that is internalized within early endosomes (EEs) through either Rab21, or Rab5, or Afr4 small GTPase, depending on the internalization site and cell type (see text and Figure 2). In EEs, active α5β1 integrin may activate different signaling pathways such as: (i) Rac1 to promote integrin-dependent cell spreading and migration on FN; (ii) FAK to suppress anoikis in normal cells or promote cancer cell anchorage-independent growth. Moreover, in Rab25 overexpressing ovarian cancer cells Arf4-dependent active α5β1 integrin is required for correct lysosome positioning and activation of mTORC1, whose kinase activity is also regulated by glucose levels. From the EE compartment, inactive integrins can move back to the plasma membrane via sorting nexin 17 (SNX17) and the retriever complex [124]. A fraction of FN fragment-bound active α5β1 integrins is sorted into multivesicular bodies (MVBs) and degraded into lysosomes; this degradative fate depends on α5β1 integrin ubiquitination and on endosomal sorting complexes required for transport (ESCRT) protein complex. Another fraction of FN fragment-bound active α5β1 integrin instead traffics to PGCs; it is still unclear if this latter step relies on deubiquitinase (DUB) and/or SNX activities. It is posited that, in PGCs, endocytosed old FN may be separated from active α5β1 integrin that would then be free and able to bind freshly synthesized FN originating from TGN cisternae in a PI4KB and AP-1A-dependent manner. Through a RAB11B-dependent endosomal recycling pathway, vesicles containing newly synthesized FN-bound active α5β1 integrins are next directed to the basolateral side of the cell surface, where the PTPRF/PPFIA1 complex support their docking, likely via PPFIA1-mediated interaction with the β1 integrin subunit cytodomain.

In agreement with the fact that active integrins act as receptors for ECM internalization, several key regulators of endocytosis were found to localize at adhesion sites and regulate the rate at which their molecular components are renewed both in vitro and in vivo. The small GTPase Rab5 concentrates at myotendinous junctions (MTJs) of Drosophila embryos, where it promotes β position-specific (βPS) integrin turnover allowing MTJ remodeling in developing skeletal muscles [60]. Of note, internalization rather than lateral diffusion in the plasma membrane is the main mechanism responsible of βPS integrin dynamics in Drosophila MTJs [60]. In adhesion sites, integrins withstand retrograde actin flow-driven traction [61], exist in stationary ECM-bound subpopulations [62], and active integrins form tightly ordered nanoclusters [63]. MMP-assisted endocytosis may hence represent an efficient strategy to allow the turnover of ECM-bound active integrins at adhesion sites. The Rab5 GEF Ras and Rab interactor 2 (RIN2) [64] and the Rab5 GAP USP6NL (also known as RN-Tre) [65,66] respectively localize in nascent adhesions and focal adhesions to promote or inhibit active β1 integrin endocytosis and motility of ECs and fibroblasts (Figure 2). In migrating cells, small, round, and peripheral nascent adhesions initially form at leading edge lamellipodium and later either disassemble or, due to actomyosin contractility, mature into elongated and stable focal adhesions [67]. It is tempting to speculate that, by exerting opposite effects on Rab5 GTP-loading and active β1 integrin internalization, RIN2 [64] and RN-Tre [65,66] may co-operate in funneling the conversion of nascent adhesions into focal adhesions in migrating cells. In fibroblasts, FN-bound α5β1 integrin slides outside focal adhesions and translocates along stress fibers giving rise to elongated fibrillar adhesions and FN fibrils [67] that are not influenced by Rab5 activity [65]. At least in ECs [6], active α5β1 integrin endocytosis at fibrillar adhesions and FN fibril turnover are promoted instead by Rab21, which localizes to adhesion sites [68,69] and was previously reported to stimulate integrin internalization [68,70,71] (Figure 2).

Working model for Rab5/Rab21 small GTPase interplay in active α5β1 integrin endocytosis in non-cancer cells.

Figure 2.
Working model for Rab5/Rab21 small GTPase interplay in active α5β1 integrin endocytosis in non-cancer cells.

The Rab5 GEF RIN2 and Rab5 GAP RN-Tre/USP6NL are respectively located in nascent and focal adhesions. Due to their enzymatic antagonistic activity on Rab5 GTP-loading and active α5β1 integrin internalization, RIN2 and RN-Tre might co-operate to foster the evolution of nascent into focal adhesions. Moving along stress fibers, active α5β1 integrins generate fibrillar adhesions and FN fibrils, which are not influenced by Rab5. Indeed, in ECs active α5β1 integrin endocytosis at fibrillar adhesions and FN fibril turnover are promoted instead by Rab21.

Figure 2.
Working model for Rab5/Rab21 small GTPase interplay in active α5β1 integrin endocytosis in non-cancer cells.

The Rab5 GEF RIN2 and Rab5 GAP RN-Tre/USP6NL are respectively located in nascent and focal adhesions. Due to their enzymatic antagonistic activity on Rab5 GTP-loading and active α5β1 integrin internalization, RIN2 and RN-Tre might co-operate to foster the evolution of nascent into focal adhesions. Moving along stress fibers, active α5β1 integrins generate fibrillar adhesions and FN fibrils, which are not influenced by Rab5. Indeed, in ECs active α5β1 integrin endocytosis at fibrillar adhesions and FN fibril turnover are promoted instead by Rab21.

Differently from ECs [6], Arf4, rather than Rab21, was implicated in the endocytosis of FN-bound active α5β1 integrin from FN fibrils specifically localized in the subnuclear of area of A2780 ovarian cancer cells overexpressing Rab25 (A2780-Rab25) [72] (Figure 1), which was previously found to co-operate with chloride intracellular channel protein 3 (CLIC3) to allow the recycling from late endosomes/lysosomes of endocytosed FN-bound active α5β1 integrin in this cell line [73]. Interestingly, in A2780-Rab25 ovarian cancer cells, upon endocytosis of FN-bound active α5β1 integrin and its delivery to late endosomes/lysosomes, likely due to the increased lysosomal concentration of amino acids caused by FN degradation, the master regulator of cell metabolism and growth mechanistic target of rapamycin complex 1 (mTORC1) gets activated [72] (Figure 1). Similarly, in starving conditions, mammary epithelial cells up-regulate α6β4 integrin-mediated laminin endocytosis and lysosomal degradation thus resulting in mTORC1 activation [74]. In addition, pancreatic ductal adenocarcinoma cells internalize and degrade type I and type IV collagens as source of proline that fuels tricarboxylic acid cycle metabolism under nutrient limited conditions [75]. Finally, the metabolic sensor adenosine monophosphate activated protein kinase (AMPK), which is activated when AMP level raises during energy stresses, was discovered to inhibit the transcription of the α5β1 integrin-specific adaptors and activators tensin1 and 3, thus impairing α5β1 integrin activation and FN fibrillogenesis [76].

In sum, the turnover of the different types of cell adhesion structures relies on distinct pro-endocytic small GTPases and regulatory proteins that differentially modulate in space and time the internalization of ECM-bound active integrins. Furthermore, a direct link between ECM-bound active integrin traffic and nutrient signaling exists. To support their proliferation rate, cancer cells exploit active integrin-mediated ECM endocytosis as an effective strategy to directly acquire nutrients from the extracellular environment.

Active integrin traffic role in establishing cell polarity

Polarized epithelial cells, neurons, vascular ECs, and directional migrating cells are characterized by spatially and functionally distinct plasma membrane areas defined by PAR proteins, the CRB complex, and phosphatidylinositol-phosphates (PIPs) [77]. β1 integrin-mediated adhesion to the ECM elicits biochemical signals aimed at establishing and maintaining apico-basal polarity both in epithelial cells [78] and ECs [79]. Specifically, β1 integrin functions upstream of PAR polarity proteins in the signaling cascade that defines EC apico-basal axis, driving vascular morphogenesis and lumen formation [79] in response to FN [80,81]. On the contrary, basement membrane proteins, such as laminin, are alternative β1 integrin ligands that inhibit vascular morphogenesis, while maintaining the stability of mature blood vessels [80,81]. During blood vessel formation, once apico-basal axis is defined, FN-bound active α5β1 integrin signals to keep directing the secretion of freshly synthesized endogenous FN towards the abluminal basolateral plasma membrane of ECs [82], giving rise to a self-sustaining polarity signaling cascade.

In ECs, apart from its ability to extracellularly interact with SEMA3A or VEGF-A [83], the transmembrane glycoprotein neuropilin 1 (NRP1) localizes at adhesion sites [66,69,84–86], where it promotes FN-bound active α5β1 integrin endocytosis [84] (Figure 1). The binding of extracellular NRP1 b1 domain to the C-terminal basic motif of SEMA3A [34,83] or C-end rule (CendR) peptides [87] fosters the internalization of NRP1 [88] and associated membrane receptor cargos, such as active α5β1 integrin [34,84]. NRP1-dependent endocytosis largely relies on its short cytodomain [84,88] that, via its C-terminal Ser-Glu-Ala (SEA) motif, binds the PSD95-DLG1-ZO1 (PDZ) domain of the endocytic adaptor GAIP interacting protein C terminus, member 1 (GIPC1, also known as synectin) that associates to myosin VI (MYO6) motor to allow the transport of early endosomes through the cortical actin network [89]. Arterial branching morphogenesis is substantially impaired in knock-in mice lacking NRP1 cytodomain [90] and, albeit at lower extent, in GIPC1 [91] and MYO6 [92] knock-out animals. In contrast, knock-in mice expressing a b1 domain NRP1 mutant unable to bind VEGF-A only do not display any vascular defect [93]. Altogether, these findings support a model in which, VEGF-A-independent NRP1/GIPC1/MYO6-driven internalization of transmembrane cargos, such as integrins, promote FN-dependent branching [94] and vascular [95] morphogenesis.

Along with the fact that, upon NRP1-driven internalization, active α5β1 integrins are returned back to the EC surface [84], the observation that, via its cytodomain, NRP1 also promotes endothelial FN fibrillogenesis [84] suggests that these integrins may, either by signaling or by direct binding or both, favor the exocytosis of newly synthesized FN from perinuclear trans-Golgi network (TGN) to replace MMP-cleaved FN fibrils. This hypothesis is also in agreement with the observation that endocytosed active α5 [6] and β1 [96] integrins are recycled, likely from vesicular compartments laid closer to the nucleus and farther from the plasma membrane [97], with considerably slower kinetics compared with their inactive counterparts in different cell types. Indeed, in ECs, Rab21, which may interact with NRP1 via adaptor protein containing a PH domain, PTB domain, and leucine zipper motif 1 (APPL1) and GIPC1 [42], promotes the internalization in early endosomes of FN fragment-bound active α5β1 integrins, which subsequently traffic to post-Golgi carrier (PGC) vesicles stemmed from the TGN in a phosphatidylinositol 4-kinase, catalytic, beta (PI4KB) and clathrin adaptor protein complex-1A (AP-1A)-dependent manner and containing freshly synthesized FN [6] (Figure 1). Albeit further studies are needed to elucidate this step, once in PGCs, active α5β1 integrins might exchange exhausted and to-be-degraded FN fragments for newly synthesized FN dimers (Figure 1). Whether internalized NRP1 reaches the TGN/PGC compartment and participates in the recycling of active α5β1 integrins is still open issue. Next, RAB11B routes PGCs containing newly synthesized FN-bound active α5β1 towards basolateral side of ECs, where a complex formed by protein tyrosine phosphatase receptor type f polypeptide (PTPRF, otherwise identified as leukocyte common antigen related — LAR protein) and PTPRF interacting protein a1 (PPFIA1), also known as liprin-α1, concentrates in close proximity of fibrillar adhesions [6] (Figure 1). Intriguingly, in neuronal pre-synapses PTPRF and PPFIA1/liprin-α1 synchronize endo-exocytic traffic and mediate the docking of neurotransmitter vesicles at the plasma membrane [98,99]. In addition, similarly to kindlins, the emergence of liprins has also been implicated in the rise of multicellular organisms [16]. Analogously to its neuronal function and due to its binding to the β1 cytodomain of active α5β1 integrin, in ECs PPFIA1/liprin-α1 promotes the docking of PGCs containing newly synthesized FN and active α5β1 integrins [6]. The fusion of these PGCs with the plasma membrane may bolster the local release of newly synthesized FN and active α5β1 integrins and the exchange of old for new fibrillar FN at the abluminal side of endothelium [6]. Of note, PPFIA1/liprin-α1 silencing also affects subendothelial FN deposition and vascular morphogenesis in developing Zebrafish embryo [6]. Supporting its role in defining EC polarity [6], PPFIA1/liprin-α1 was discovered to control the basolateral secretion of at least three additional ECM components that are pivotal regulators of vascular ECM organization and remodeling, namely lysyl oxidase, multimerin 2, and dystroglycan-1 [100]. Thus, ECs employ a synaptic-like machinery to control the coupling of active α5β1 integrin endo-exocytosis and the replenishment of degraded FN fibrils with the basolateral secretion of TGN-derived newly synthesized FN [6], along with proteins involved in FN rehandling [100]. It has been for long known that, during embryogenesis, dynamically remodeling angiogenic blood vessels develop in a FN-rich ECM, which, during the subsequent vascular maturation and stabilization phase, is substituted with a laminin-rich basement membrane [81,101]. To this matter, the discovery that PPFIA1/liprin-α1 also controls the basolateral secretion of the basement membrane organizing protein dystroglycan-1 [100] suggests that PPFIA1/liprin-α1 may participate in orchestrating the transition from angiogenic to quiescent blood vessels [81].

More recently, it has been reported that the other TGN-associated clathrin adaptor protein, known to participate together with AP-1 [102] in the secretory pathway, namely Golgi-localized gamma ear-containing Arf-binding protein 2 (GGA2) [102], is also involved in active β1 integrin recycling in human MDA-MB-231 breast cancer cells [103]. The association of GGA2 with active β1 integrin is stabilized by the small GTPase RAB13, which also supports the return of internalized active β1 integrin to the cell surface. Moreover, both GGA2 and RAB13 are required for efficient cancer cell migration and invasion [103]. Of note, along with RAB13, proximity biotinylation analyses also identified PTPRF as a GGA2 interactor [103]. In addition, since PTPRF was previously reported to drive active α5β1 integrin recycling in ECs [6,104], it may hence function as a multipurpose docking receptor that, localizing at ECM adhesion sites [105], allows the polarized and targeted recycling of active β1 integrins via different TGN-connected trafficking pathways. Indeed, GGAs and AP-1 TGN-associated clathrin adaptors may have overlapping [102], as well as distinct functions, GGAs, but not AP-1, transporting ubiquitinated protein cargos [106,107]

Internalized active integrins elicit endosomal signaling pathways

Rac1-stimulated actin branched polymerization drives the formation of plasma membrane extensions, known as lamellipodia, which need to be confined at the leading edge of migrating cells to effectively allow directional motility [108]. In this context, integrin-ECM engagement at the cell front triggers a self-sustaining Rac1-activating positive feedback loop that supports lamellipodium-driven cell motility [109]. In addition, Rac1 endocytosis, from and recycling to, the plasma membrane represents a strategy to selectively restrain and polarize, in space and time, the signaling of this small GTPase [110]. Early endosomes have been identified as further subcellular sites for Rac1 activation in addition to the plasma membrane [110]. Indeed, the major Rac1 GEF T-lymphoma invasion and metastasis-inducing protein 1 (TIAM1) was found to reside on early endosomes of HeLa cancer cells [110], likely because, via its pleckstrin homology (PH) domain [111], TIAM1 binds PI3P [112], a phospholipid produced by the key Rab5 effector PI3KC3, also known as VPS34 [113] (Figure 1). The small GTPase R-Ras, one of the master regulators of integrin function [114], is highly expressed in vascular ECs [64], localizes in lamellipodia-associated nascent ECM-adhesions, where it recruits RIN2 as a result of the interaction with its Ras association (RA) domain [64]. In ECs, the binding to R-Ras converts RIN2 from a Rab5 GEF to an adaptor that first interacts at high affinity with Rab5-GTP to selectively promote ECM-bound/active β1 integrin endocytosis and next causes R-Ras repositioning on early endosomes [64]. After the active β1 integrin/Rab5/RIN2-dependent transfer on early endosomes, R-Ras contacts the RA domain of TIAM1, thus promoting the GTP loading of Rac1 [64] (Figure 1) followed by its polarized relocation to the plasma membrane, likely via the small GTPase Arf6 [110,115]. In sum, it appears that, at the leading edge of migrating ECs, the endocytosis of active β1 integrins co-ordinates the RIN2-dependent translocation of R-Ras on early endosomes, where it triggers, via TIAM1, a self-sustaining Rac1-activating positive feedback loop that drives directed cell motility. Along this line, it was previously reported that in cancer cells Rab5 gets activated upon integrin-mediated cell spreading on FN [116] that, indeed, depends on a focal adhesion kinase (FAK)-Rab5-Rac1 signaling pathway that critically acts downstream of the mechanosensing protein vinculin [117]. More recently, it has been proposed that FAK may favor Rab5 GTP loading by binding and inhibiting the Rab GAP activity [118] of the p85α subunit of PI3K [119]. Furthermore, upon GTP loading, Rac1 elicits additional phosphorylation of FAK that in turn promotes further Rac1 activation, giving rise to a positive feedback mechanism [117]. Altogether these data support a model in which active integrin endocytosis is key in promoting the GTP loading of Rac1 to enable cell spreading.

FAK activation at the plasma membrane regulates cell proliferation, survival, migration, and invasion [120]. However, it has been proposed that endosomal FAK signaling may also support the resistance of normal cells to anoikis as well as breast cancer cell anchorage independent growth and metastatic dissemination [121]. In the cytoplasm, FAK exists in an autoinhibited conformation and the binding of its FERM domain to PI(4,5)P2, which at the plasma membrane is generated in close proximity to ECM-bound integrins, elicits FAK conformational activation [120]. Consistently, the FERM domain is sufficient to recruit FAK to active integrin containing endosomes through still unknown mechanisms [121] (Figure 1). While conformational activation is not required for vesicular targeting, FAK endosomal signaling strictly depends on its tyrosine kinase activity. Similarly to what observed at the plasma membrane [122], FAK, once activated, may promote talin recruitment and tension-independent activation on endosomes. Mechanistically, it has been proposed that FAK phosphorylates and activates type I phosphatidylinositol phosphate kinase (PIPKIγi2), generating PI(4,5)P2 promoting talin recruitment [123]. The presence of talin on endosomes may facilitate the maintenance of integrin active conformation during its recycling to the plasma membrane, even in the absence of ligands [123].

Perspective

  • Differently from what previously hypothesized, integrins do not need to be conformationally inactivated to be endocytosed. Dedicated transmembrane and cytosolic proteins contribute to ECM-bound active integrin internalization and recycling to the cell surface, along with the secretion of newly synthesized ECM.

  • Active integrin endocytosis and traffic control ECM and adhesion site turnover, metabolism, polarity, and endosomal signaling supporting motility and survival both of normal and cancer cells.

  • It will be crucial to identify and thoroughly dissect the molecular mechanisms responsible for those signaling aspects of active integrin traffic still poorly understood, such as those involved in the control of ECM-containing PGCs from the TGN or in the targeting of FAK on endosomes.

Competing Interests

The authors declare that there are no competing interests associated with the manuscript.

Funding

Supported by Fondazione AIRC (IG grants #16702 and 21315 to G.S.; #20366 to D.V.); FPRC-ONLUS Grant ‘FPRC - 5 per mille 2014 Ministero Salute' (to G.S.); Associazione ‘Augusto per la Vita’ (to G.S.); Fondazione Telethon [grant n. GGP15102] (to G.S.).

Author Contribution

All authors contributed substantially to the conception and design of work leading to this review, provided content and wrote the article. All authors have approved the final version for publication.

Acknowledgements

The authors thank Giulia Villari for critical reading of the manuscript and discussion.

Abbreviations

     
  • AMPK

    adenosine monophosphate activated protein kinase

  •  
  • AP-1A

    adaptor protein complex-1A

  •  
  • CLASPs

    cytoplasmic linker associated proteins

  •  
  • CLIC3

    chloride intracellular channel protein 3

  •  
  • CXCL12

    C-X-C motif chemokine 12

  •  
  • ECM

    extracellular matrix

  •  
  • ECs

    endothelial cells

  •  
  • FAK

    focal adhesion kinase

  •  
  • FERM

    four-point-one, ezrin, radixin, moesin

  •  
  • FN

    fibronectin

  •  
  • GAP

    GTPase activating protein

  •  
  • GEFs

    guanine nucleotide exchange factors

  •  
  • GGA2

    Golgi-localized gamma ear-containing Arf-binding protein 2

  •  
  • MMPs

    matrix metalloproteinases

  •  
  • MT1-MMP

    membrane-type 1 MMP

  •  
  • MTJs

    myotendinous junctions

  •  
  • MYO6

    myosin VI

  •  
  • NRP1

    neuropilin 1

  •  
  • PGC

    post-Golgi carrier

  •  
  • PI4KB

    phosphatidylinositol 4-kinase, catalytic, beta

  •  
  • PPFIA1

    PTPRF interacting protein a1

  •  
  • RA

    Ras association

  •  
  • TGN

    trans-Golgi network

  •  
  • TIAM1

    T-lymphoma invasion and metastasis-inducing protein 1

  •  
  • VEGF-A

    vascular endothelial growth factor-A

  •  
  • βPS

    β position-specific

References

References
1
Bachmann
,
M.
,
Kukkurainen
,
S.
,
Hytönen
,
V.P.
and
Wehrle-Haller
,
B.
(
2019
)
Cell adhesion by integrins
.
Physiol. Rev.
99
,
1655
1699
2
Byron
,
A.
and
Frame
,
M.C.
(
2016
)
Adhesion protein networks reveal functions proximal and distal to cell-matrix contacts
.
Curr. Opin. Cell Biol.
39
,
93
100
3
Humphries
,
J.D.
,
Chastney
,
M.R.
,
Askari
,
J.A.
and
Humphries
,
M.J.
(
2019
)
Signal transduction via integrin adhesion complexes
.
Curr. Opin. Cell Biol.
56
,
14
21
4
Hynes
,
R.O.
(
2012
)
Evolution: the evolution of metazoan extracellular matrix
.
J. Cell Biol.
196
,
671
679
5
Li
,
J.
,
Su
,
Y.
,
Xia
,
W.
,
Qin
,
Y.
,
Humphries
,
M.J.
,
Vestweber
,
D.
et al (
2017
)
Conformational equilibria and intrinsic affinities define integrin activation
.
EMBO J.
36
,
629
645
6
Mana
,
G.
,
Clapero
,
F.
,
Panieri
,
E.
,
Panero
,
V.
,
Böttcher
,
R.T.
,
Tseng
,
H.Y.
et al (
2016
)
PPFIA1 drives active α5β1 integrin recycling and controls fibronectin fibrillogenesis and vascular morphogenesis
.
Nat. Commun.
7
,
13546
7
Goult
,
B.T.
,
Yan
,
J.
and
Schwartz
,
M.A.
(
2018
)
Talin as a mechanosensitive signaling hub
.
J. Cell Biol.
217
,
3776
3784
8
Sun
,
Z.
,
Costell
,
M.
and
Fässler
,
R.
(
2019
)
Integrin activation by talin, kindlin and mechanical forces
.
Nat. Cell Biol.
21
,
25
31
9
Rognoni
,
E.
,
Ruppert
,
R.
and
Fässler
,
R.
(
2016
)
The kindlin family: functions, signaling properties and implications for human disease
.
J. Cell Sci.
129
,
17
27
10
Theodosiou
,
M.
,
Widmaier
,
M.
,
Böttcher
,
R.T.
,
Rognoni
,
E.
,
Veelders
,
M.
,
Bharadwaj
,
M.
et al (
2016
)
Kindlin-2 cooperates with talin to activate integrins and induces cell spreading by directly binding paxillin
.
eLife
5
,
e10130
11
Klapproth
,
S.
,
Bromberger
,
T.
,
Türk
,
C.
,
Krüger
,
M.
and
Moser
,
M.
(
2019
)
A kindlin-3-leupaxin-paxillin signaling pathway regulates podosome stability
.
J. Cell Biol.
218
,
3436
3454
12
Zhu
,
L.
,
Liu
,
H.
,
Lu
,
F.
,
Yang
,
J.
,
Byzova
,
T.V.
and
Qin
,
J.
(
2019
)
Structural basis of paxillin recruitment by kindlin-2 in regulating cell adhesion
.
Structure
27
,
1686
1697.e5
13
Sebé-Pedrós
,
A.
,
Degnan
,
B.M.
and
Ruiz-Trillo
,
I.
(
2017
)
The origin of metazoa: a unicellular perspective
.
Nat. Rev. Genet.
18
,
498
512
14
King
,
N.
,
Westbrook
,
M.J.
,
Young
,
S.L.
,
Kuo
,
A.
,
Abedin
,
M.
,
Chapman
,
J.
et al (
2008
)
The genome of the choanoflagellate monosiga brevicollis and the origin of metazoans
.
Nature
451
,
783
788
15
Sebé-Pedrós
,
A.
,
Roger
,
A.J.
,
Lang
,
F.B.
,
King
,
N.
and
Ruiz-Trillo
,
I.
(
2010
)
Ancient origin of the integrin-mediated adhesion and signaling machinery
.
Proc. Natl Acad. Sci. U.S.A.
107
,
10142
7
16
Paps
,
J.
and
Holland
,
P.W.H.
(
2018
)
Reconstruction of the ancestral metazoan genome reveals an increase in genomic novelty
.
Nat. Commun.
9
,
1730
17
Serini
,
G.
,
Valdembri
,
D.
,
Zanivan
,
S.
,
Morterra
,
G.
,
Burkhardt
,
C.
,
Caccavari
,
F.
et al (
2003
)
Class 3 semaphorins control vascular morphogenesis by inhibiting integrin function
.
Nature
424
,
391
397
18
Valdembri
,
D.
,
Regano
,
D.
,
Maione
,
F.
,
Giraudo
,
E.
and
Serini
,
G.
(
2016
)
Class 3 semaphorins in cardiovascular development
.
Cell Adh. Migr.
10
,
641
651
19
Moreno-Layseca
,
P.
,
Icha
,
J.
,
Hamidi
,
H.
and
Ivaska
,
J.
(
2019
)
Integrin trafficking in cells and tissues
.
Nat. Cell Biol.
21
,
122
132
20
Zhu
,
L.
,
Yang
,
J.
,
Bromberger
,
T.
,
Holly
,
A.
,
Lu
,
F.
,
Liu
,
H.
et al (
2017
)
Structure of Rap1b bound to talin reveals a pathway for triggering integrin activation
.
Nat. Commun.
8
,
1744
21
Camp
,
D.
,
Haage
,
A.
,
Solianova
,
V.
,
Castle
,
W.M.
,
Xu
,
Q.A.
,
Lostchuck
,
E.
et al (
2018
)
Direct binding of talin to Rap1 is required for cell-ECM adhesion in drosophila
.
J. Cell Sci.
131
,
jcs225144
22
Bromberger
,
T.
,
Klapproth
,
S.
,
Rohwedder
,
I.
,
Zhu
,
L.
,
Mittmann
,
L.
,
Reichel
,
C.A.
et al (
2018
)
Direct Rap1/Talin1 interaction regulates platelet and neutrophil integrin activity in mice
.
Blood
132
,
2754
2762
23
Gingras
,
A.R.
,
Lagarrigue
,
F.
,
Cuevas
,
M.N.
,
Valadez
,
A.J.
,
Zorovich
,
M.
,
McLaughlin
,
W.
et al (
2019
)
Rap1 binding and a lipid-dependent helix in talin F1 domain promote integrin activation in tandem
.
J. Cell Biol.
218
,
1799
1809
24
Bromberger
,
T.
,
Zhu
,
L.
,
Klapproth
,
S.
,
Qin
,
J.
and
Moser
,
M.
(
2019
)
Rap1 and membrane lipids cooperatively recruit talin to trigger integrin activation
.
J. Cell Sci.
132
,
jcs235531
25
Lagarrigue
,
F.
,
Kim
,
C.
and
Ginsberg
,
M.H.
(
2016
)
The Rap1-RIAM-talin axis of integrin activation and blood cell function
.
Blood
128
,
479
487
26
Chrzanowska-Wodnicka
,
M.
(
2013
)
Distinct functions for Rap1 signaling in vascular morphogenesis and dysfunction
.
Exp. Cell Res.
319
,
2350
2359
27
Ellis
,
S.J.
,
Goult
,
B.T.
,
Fairchild
,
M.J.
,
Harris
,
N.J.
,
Long
,
J.
,
Lobo
,
P.
et al (
2013
)
Talin autoinhibition is required for morphogenesis
.
Curr. Biol.
23
,
1825
1833
28
Hogg
,
N.
,
Patzak
,
I.
and
Willenbrock
,
F.
(
2011
)
The insider's guide to leukocyte integrin signalling and function
.
Nat. Rev. Immunol.
11
,
416
426
29
Strazza
,
M.
,
Azoulay-Alfaguter
,
I.
,
Peled
,
M.
,
Smrcka
,
A.V.
,
Skolnik
,
E.Y.
,
Srivastava
,
S.
et al (
2017
)
PLCε1 regulates SDF-1α-induced lymphocyte adhesion and migration to sites of inflammation
.
Proc. Natl Acad. Sci. U.S.A.
114
,
2693
2698
30
Chrzanowska-Wodnicka
,
M.
(
2017
)
Rap1 in endothelial biology
.
Curr. Opin. Hematol.
24
,
248
255
31
Gloerich
,
M.
and
Bos
,
J.L.
(
2011
)
Regulating Rap small G-proteins in time and space
.
Trends Cell Biol.
21
,
615
623
32
Wang
,
Y.
,
He
,
H.
,
Srivastava
,
N.
,
Vikarunnessa
,
S.
,
Chen
,
Y.B.
,
Jiang
,
J.
et al (
2012
)
Plexins are GTPase-activating proteins for Rap and are activated by induced dimerization
.
Sci. Signal.
5
,
ra6
33
Bos
,
J.L.
and
Pannekoek
,
W.J.
(
2012
)
Semaphorin signaling meets rap
.
Sci. Signal.
5
,
pe6
34
Gioelli
,
N.
,
Maione
,
F.
,
Camillo
,
C.
,
Ghitti
,
M.
,
Valdembri
,
D.
,
Morello
,
N.
et al (
2018
)
A rationally designed NRP1-independent superagonist SEMA3A mutant is an effective anticancer agent
.
Sci. Transl. Med.
10
,
eaah4807
35
Worzfeld
,
T.
,
Swiercz
,
J.M.
,
Sentürk
,
A.
,
Genz
,
B.
,
Korostylev
,
A.
,
Deng
,
S.
et al (
2014
)
Genetic dissection of plexin signaling in vivo
.
Proc. Natl Acad. Sci. U.S.A.
111
,
2194
2199
36
Serini
,
G.
and
Bussolino
,
F.
(
2004
)
Common cues in vascular and axon guidance
.
Physiology (Bethesda)
19
,
348
354
37
Bussolino
,
F.
,
Valdembri
,
D.
,
Caccavari
,
F.
and
Serini
,
G.
(
2006
)
Semaphoring vascular morphogenesis
.
Endothelium
13
,
81
91
38
Bouvard
,
D.
,
Pouwels
,
J.
,
De Franceschi
,
N.
and
Ivaska
,
J.
(
2013
)
Integrin inactivators: balancing cellular functions in vitro and in vivo
.
Nat. Rev. Mol. Cell Biol.
14
,
430
442
39
Bussolino
,
F.
,
Caccavari
,
F.
,
Valdembri
,
D.
and
Serini
,
G.
(
2009
)
Angiogenesis: a balancing act between integrin activation and inhibition?
Eur. Cytokine Netw.
20
,
191
196
40
Bridgewater
,
R.E.
,
Norman
,
J.C.
and
Caswell
,
P.T.
(
2012
)
Integrin trafficking at a glance
.
J. Cell Sci.
125
(
Pt 16
),
3695
3701
41
Valdembri
,
D.
and
Serini
,
G.
(
2012
)
Regulation of adhesion site dynamics by integrin traffic
.
Curr. Opin. Cell Biol.
24
,
582
591
42
Valdembri
,
D.
,
Sandri
,
C.
,
Santambrogio
,
M.
and
Serini
,
G.
(
2011
)
Regulation of integrins by conformation and traffic: it takes two to tango
.
Mol. Biosyst.
7
,
2539
2546
43
Santambrogio
,
M.
,
Valdembri
,
D.
and
Serini
,
G.
(
2011
)
Increasing traffic on vascular routes
.
Mol. Aspects Med.
32
,
112
122
44
Bretscher
,
M.S.
(
1989
)
Endocytosis and recycling of the fibronectin receptor in CHO cells
.
EMBO J.
8
,
1341
1348
45
Lawson
,
M.A.
and
Maxfield
,
F.R.
(
1995
)
Ca2+- and calcineurin-dependent recycling of an integrin to the front of migrating neutrophils
.
Nature
377
,
75
79
46
Puklin-Faucher
,
E.
and
Sheetz
,
M.P.
(
2009
)
The mechanical integrin cycle
.
J. Cell Sci.
122
(
Pt 2
),
179
186
47
Byron
,
A.
,
Humphries
,
J.D.
,
Askari
,
J.A.
,
Craig
,
S.E.
,
Mould
,
A.P.
and
Humphries
,
M.J.
(
2009
)
Anti-integrin monoclonal antibodies
.
J. Cell Sci.
122
(
Pt 22
),
4009
4011
48
Schwarzbauer
,
J.E.
and
DeSimone
,
D.W.
(
2011
)
Fibronectins, their fibrillogenesis, and in vivo functions
.
Cold Spring Harb. Perspect. Biol.
3
,
a005041
49
Vogel
,
V.
(
2018
)
Unraveling the mechanobiology of extracellular matrix
.
Annu. Rev. Physiol.
80
,
353
387
50
Humphrey
,
J.D.
,
Dufresne
,
E.R.
and
Schwartz
,
M.A.
(
2014
)
Mechanotransduction and extracellular matrix homeostasis
.
Nat. Rev. Mol. Cell Biol.
15
,
802
812
51
Bonnans
,
C.
,
Chou
,
J.
and
Werb
,
Z.
(
2014
)
Remodelling the extracellular matrix in development and disease
.
Nat. Rev. Mol. Cell Biol.
15
,
786
801
52
Stehbens
,
S.J.
,
Paszek
,
M.
,
Pemble
,
H.
,
Ettinger
,
A.
,
Gierke
,
S.
and
Wittmann
,
T.
(
2014
)
CLASPs link focal-adhesion-associated microtubule capture to localized exocytosis and adhesion site turnover
.
Nat. Cell Biol.
16
,
561
573
53
Wang
,
Y.
and
McNiven
,
M.A.
(
2012
)
Invasive matrix degradation at focal adhesions occurs via protease recruitment by a FAK-p130Cas complex
.
J. Cell Biol.
196
,
375
385
54
Shi
,
F.
and
Sottile
,
J.
(
2011
)
MT1-MMP regulates the turnover and endocytosis of extracellular matrix fibronectin
.
J. Cell Sci.
124
(
Pt 23
),
4039
4050
55
Shi
,
F.
and
Sottile
,
J.
(
2008
)
Caveolin-1-dependent beta1 integrin endocytosis is a critical regulator of fibronectin turnover
.
J. Cell Sci.
121
,
2360
2371
56
Singh
,
P.
,
Carraher
,
C.
and
Schwarzbauer
,
J.E.
(
2010
)
Assembly of fibronectin extracellular matrix
.
Annu. Rev. Cell Dev. Biol.
26
,
397
419
57
Shi
,
F.
,
Harman
,
J.
,
Fujiwara
,
K.
and
Sottile
,
J.
(
2010
)
Collagen I matrix turnover is regulated by fibronectin polymerization
.
Am. J. Physiol. Cell Physiol.
298
,
C1265
C1275
58
Memmo
,
L.M.
and
McKeown-Longo
,
P.
(
1998
)
The alphavbeta5 integrin functions as an endocytic receptor for vitronectin
.
J. Cell Sci.
111
(
Pt 4
),
425
433
PMID:
[PubMed]
59
Leonoudakis
,
D.
,
Huang
,
G.
,
Akhavan
,
A.
,
Fata
,
J.E.
,
Singh
,
M.
,
Gray
,
J.W.
et al (
2014
)
Endocytic trafficking of laminin is controlled by dystroglycan and is disrupted in cancers
.
J. Cell Sci.
127
(
Pt 22
),
4894
4903
60
Yuan
,
L.
,
Fairchild
,
M.J.
,
Perkins
,
A.D.
and
Tanentzapf
,
G.
(
2010
)
Analysis of integrin turnover in fly myotendinous junctions
.
J. Cell Sci.
123
(
Pt 6
),
939
946
61
Hu
,
K.
,
Ji
,
L.
,
Applegate
,
K.T.
,
Danuser
,
G.
and
Waterman-Storer
,
C.M.
(
2007
)
Differential transmission of actin motion within focal adhesions
.
Science
315
,
111
115
62
Rossier
,
O.
,
Octeau
,
V.
,
Sibarita
,
J.B.
,
Leduc
,
C.
,
Tessier
,
B.
,
Nair
,
D.
et al (
2012
)
Integrins β1 and β3 exhibit distinct dynamic nanoscale organizations inside focal adhesions
.
Nat. Cell Biol.
14
,
1057
1067
63
Spiess
,
M.
,
Hernandez-Varas
,
P.
,
Oddone
,
A.
,
Olofsson
,
H.
,
Blom
,
H.
,
Waithe
,
D.
et al (
2018
)
Active and inactive β1 integrins segregate into distinct nanoclusters in focal adhesions
.
J. Cell Biol.
217
,
1929
1940
64
Sandri
,
C.
,
Caccavari
,
F.
,
Valdembri
,
D.
,
Camillo
,
C.
,
Veltel
,
S.
,
Santambrogio
,
M.
et al (
2012
)
The R-Ras/RIN2/Rab5 complex controls endothelial cell adhesion and morphogenesis via active integrin endocytosis and Rac signaling
.
Cell Res.
22
,
1479
1501
65
Palamidessi
,
A.
,
Frittoli
,
E.
,
Ducano
,
N.
,
Offenhauser
,
N.
,
Sigismund
,
S.
,
Kajiho
,
H.
et al (
2013
)
The GTPase-activating protein RN-tre controls focal adhesion turnover and cell migration
.
Curr. Biol.
23
,
2355
2364
66
Dong
,
J.M.
,
Tay
,
F.P.
,
Swa
,
H.L.
,
Gunaratne
,
J.
,
Leung
,
T.
,
Burke
,
B.
et al (
2016
)
Proximity biotinylation provides insight into the molecular composition of focal adhesions at the nanometer scale
.
Sci. Signal.
9
,
rs4
67
Kechagia
,
J.Z.
,
Ivaska
,
J.
and
Roca-Cusachs
,
P.
(
2019
)
Integrins as biomechanical sensors of the microenvironment
.
Nat. Rev. Mol. Cell Biol.
20
,
457
473
68
Pellinen
,
T.
,
Arjonen
,
A.
,
Vuoriluoto
,
K.
,
Kallio
,
K.
,
Fransen
,
J.A.
and
Ivaska
,
J.
(
2006
)
Small GTPase Rab21 regulates cell adhesion and controls endosomal traffic of beta1-integrins
.
J. Cell Biol.
173
,
767
780
69
Kuo
,
J.C.
,
Han
,
X.
,
Hsiao
,
C.T.
,
Yates Iii
,
J.R.
and
Waterman
,
C.M.
(
2011
)
Analysis of the myosin-II-responsive focal adhesion proteome reveals a role for beta-Pix in negative regulation of focal adhesion maturation
.
Nat. Cell Biol.
13
,
383
393
70
Mai
,
A.
,
Veltel
,
S.
,
Pellinen
,
T.
,
Padzik
,
A.
,
Coffey
,
E.
,
Marjomäki
,
V.
et al (
2011
)
Competitive binding of Rab21 and p120RasGAP to integrins regulates receptor traffic and migration
.
J. Cell Biol.
194
,
291
306
71
Pellinen
,
T.
,
Tuomi
,
S.
,
Arjonen
,
A.
,
Wolf
,
M.
,
Edgren
,
H.
,
Meyer
,
H.
et al (
2008
)
Integrin trafficking regulated by Rab21 is necessary for cytokinesis
.
Dev. Cell
15
,
371
385
72
Rainero
,
E.
,
Howe
,
J.D.
,
Caswell
,
P.T.
,
Jamieson
,
N.B.
,
Anderson
,
K.
,
Critchley
,
D.R.
et al (
2015
)
Ligand-occupied integrin internalization links nutrient signaling to invasive migration
.
Cell Rep.
10
,
398
413
73
Dozynkiewicz
,
M.A.
,
Jamieson
,
N.B.
,
Macpherson
,
I.
,
Grindlay
,
J.
,
van den Berghe
,
P.V.
,
von Thun
,
A.
et al (
2012
)
Rab25 and CLIC3 collaborate to promote integrin recycling from late endosomes/lysosomes and drive cancer progression
.
Dev. Cell
22
,
131
145
74
Muranen
,
T.
,
Iwanicki
,
M.P.
,
Curry
,
N.L.
,
Hwang
,
J.
,
DuBois
,
C.D.
,
Coloff
,
J.L.
et al (
2017
)
Starved epithelial cells uptake extracellular matrix for survival
.
Nat. Commun.
8
,
13989
75
Olivares
,
O.
,
Mayers
,
J.R.
,
Gouirand
,
V.
,
Torrence
,
M.E.
,
Gicquel
,
T.
,
Borge
,
L.
et al (
2017
)
Collagen-derived proline promotes pancreatic ductal adenocarcinoma cell survival under nutrient limited conditions
.
Nat. Commun.
8
,
16031
76
Georgiadou
,
M.
,
Lilja
,
J.
,
Jacquemet
,
G.
,
Guzmán
,
C.
,
Rafaeva
,
M.
,
Alibert
,
C.
et al (
2017
)
AMPK negatively regulates tensin-dependent integrin activity
.
J. Cell Biol.
216
,
1107
1121
77
Rodriguez-Boulan
,
E.
and
Macara
,
I.G.
(
2014
)
Organization and execution of the epithelial polarity programme
.
Nat. Rev. Mol. Cell Biol.
15
,
225
242
78
Datta
,
A.
,
Bryant
,
D.M.
and
Mostov
,
K.E.
(
2011
)
Molecular regulation of lumen morphogenesis
.
Curr. Biol.
21
,
R126
R136
79
Zovein
,
A.C.
,
Luque
,
A.
,
Turlo
,
K.A.
,
Hofmann
,
J.J.
,
Yee
,
K.M.
,
Becker
,
M.S.
et al (
2010
)
Beta1 integrin establishes endothelial cell polarity and arteriolar lumen formation via a Par3-dependent mechanism
.
Dev. Cell
18
,
39
51
80
Iruela-Arispe
,
M.L.
and
Davis
,
G.E.
(
2009
)
Cellular and molecular mechanisms of vascular lumen formation
.
Dev. Cell
16
,
222
231
81
Senger
,
D.R.
and
Davis
,
G.E.
(
2011
)
Angiogenesis
.
Cold Spring Harb. Perspect. Biol.
3
,
a005090
82
Turner
,
C.J.
,
Badu-Nkansah
,
K.
and
Hynes
,
R.O.
(
2017
)
Endothelium-derived fibronectin regulates neonatal vascular morphogenesis in an autocrine fashion
.
Angiogenesis
20
,
519
531
83
Guo
,
H.F.
and
Vander Kooi
,
C.W.
(
2015
)
Neuropilin functions as an essential cell surface receptor
.
J. Biol. Chem.
290
,
29120
6
84
Valdembri
,
D.
,
Caswell
,
P.T.
,
Anderson
,
K.I.
,
Schwarz
,
J.P.
,
König
,
I.
,
Astanina
,
E.
et al (
2009
)
Neuropilin-1/GIPC1 signaling regulates α5β1 integrin traffic and function in endothelial cells
.
PLoS Biol.
7
,
e1000025
85
Ellison
,
T.S.
,
Atkinson
,
S.J.
,
Steri
,
V.
,
Kirkup
,
B.M.
,
Preedy
,
M.E.
,
Johnson
,
R.T.
et al (
2015
)
Suppression of β3-integrin in mice triggers a neuropilin-1-dependent change in focal adhesion remodelling that can be targeted to block pathological angiogenesis
.
Dis. Model. Mech.
8
,
1105
1119
86
Schiller
,
H.B.
,
Friedel
,
C.C.
,
Boulegue
,
C.
and
Fässler
,
R.
(
2011
)
Quantitative proteomics of the integrin adhesome show a myosin II-dependent recruitment of LIM domain proteins
.
EMBO Rep.
12
,
259
266
87
Teesalu
,
T.
,
Sugahara
,
K.N.
,
Kotamraju
,
V.R.
and
Ruoslahti
,
E.
(
2009
)
C-end rule peptides mediate neuropilin-1-dependent cell, vascular, and tissue penetration
.
Proc. Natl Acad. Sci. U.S.A.
106
,
16157
16162
88
Pang
,
H.B.
,
Braun
,
G.B.
,
Friman
,
T.
,
Aza-Blanc
,
P.
,
Ruidiaz
,
M.E.
,
Sugahara
,
K.N.
et al (
2014
)
An endocytosis pathway initiated through neuropilin-1 and regulated by nutrient availability
.
Nat. Commun.
5
,
4904
89
O'Loughlin
,
T.
,
Masters
,
T.A.
and
Buss
,
F.
(
2018
)
The MYO6 interactome reveals adaptor complexes coordinating early endosome and cytoskeletal dynamics
.
EMBO Rep.
19
,
e44884
90
Lanahan
,
A.
,
Zhang
,
X.
,
Fantin
,
A.
,
Zhuang
,
Z.
,
Rivera-Molina
,
F.
,
Speichinger
,
K.
et al (
2013
)
The neuropilin 1 cytoplasmic domain is required for VEGF-A-dependent arteriogenesis
.
Dev. Cell
25
,
156
168
91
Chittenden
,
T.W.
,
Claes
,
F.
,
Lanahan
,
A.A.
,
Autiero
,
M.
,
Palac
,
R.T.
,
Tkachenko
,
E.V.
et al (
2006
)
Selective regulation of arterial branching morphogenesis by synectin
.
Dev. Cell
10
,
783
795
92
Lanahan
,
A.A.
,
Hermans
,
K.
,
Claes
,
F.
,
Kerley-Hamilton
,
J.S.
,
Zhuang
,
Z.W.
,
Giordano
,
F.J.
et al (
2010
)
VEGF receptor 2 endocytic trafficking regulates arterial morphogenesis
.
Dev. Cell
18
,
713
724
93
Gelfand
,
M.V.
,
Hagan
,
N.
,
Tata
,
A.
,
Oh
,
W.J.
,
Lacoste
,
B.
,
Kang
,
K.T.
et al (
2014
)
Neuropilin-1 functions as a VEGFR2 co-receptor to guide developmental angiogenesis independent of ligand binding
.
eLife
3
,
e03720
94
Wang
,
S.
,
Sekiguchi
,
R.
,
Daley
,
W.P.
and
Yamada
,
K.M.
(
2017
)
Patterned cell and matrix dynamics in branching morphogenesis
.
J. Cell Biol.
216
,
559
570
95
Astrof
,
S.
and
Hynes
,
R.O.
(
2009
)
Fibronectins in vascular morphogenesis
.
Angiogenesis
12
,
165
175
96
Arjonen
,
A.
,
Alanko
,
J.
,
Veltel
,
S.
and
Ivaska
,
J.
(
2012
)
Distinct recycling of active and inactive β1 integrins
.
Traffic
13
,
610
625
97
Neefjes
,
J.
,
Jongsma
,
M.M.L.
and
Berlin
,
I.
(
2017
)
Stop or go? endosome positioning in the establishment of compartment architecture, dynamics, and function
.
Trends Cell Biol.
27
,
580
594
98
Haucke
,
V.
,
Neher
,
E.
and
Sigrist
,
S.J.
(
2011
)
Protein scaffolds in the coupling of synaptic exocytosis and endocytosis
.
Nat. Rev. Neurosci.
12
,
127
138
99
Südhof
,
T.C.
(
2012
)
The presynaptic active zone
.
Neuron
75
,
11
25
100
Wei
,
H.
,
Sundararaman
,
A.
,
Dickson
,
E.
,
Rennie-Campbell
,
L.
,
Cross
,
E.
,
Heesom
,
K.J.
et al (
2019
)
Characterization of the polarized endothelial secretome
.
FASEB J.
33
,
12277
12287
101
Risau
,
W.
and
Lemmon
,
V.
(
1988
)
Changes in the vascular extracellular matrix during embryonic vasculogenesis and angiogenesis
.
Dev. Biol
125
,
441
450
102
Hirst
,
J.
,
Borner
,
G.H.
,
Antrobus
,
R.
,
Peden
,
A.A.
,
Hodson
,
N.A.
,
Sahlender
,
D.A.
et al (
2012
)
Distinct and overlapping roles for AP-1 and GGAs revealed by the “knocksideways” system
.
Curr. Biol.
22
,
1711
1716
103
Sahgal
,
P.
,
Alanko
,
J.
,
Icha
,
J.
,
Paatero
,
I.
,
Hamidi
,
H.
,
Arjonen
,
A.
et al (
2019
)
GGA2 and RAB13 promote activity-dependent β1-integrin recycling
.
J. Cell Sci.
132
,
jcs233387
104
Hamidi
,
H.
and
Ivaska
,
J.
(
2017
)
Vascular morphogenesis: an integrin and fibronectin highway
.
Curr Biol.
27
,
R158
R161
105
Serra-Pagès
,
C.
,
Kedersha
,
N.L.
,
Fazikas
,
L.
,
Medley
,
Q.
,
Debant
,
A.
and
Streuli
,
M.
(
1995
)
The LAR transmembrane protein tyrosine phosphatase and a coiled-coil LAR-interacting protein co-localize at focal adhesions
.
EMBO J.
14
,
2827
2838
106
Puertollano
,
R.
and
Bonifacino
,
J.S.
(
2004
)
Interactions of GGA3 with the ubiquitin sorting machinery
.
Nat. Cell Biol.
6
,
244
251
107
Scott
,
P.M.
,
Bilodeau
,
P.S.
,
Zhdankina
,
O.
,
Winistorfer
,
S.C.
,
Hauglund
,
M.J.
,
Allaman
,
M.M.
et al (
2004
)
GGA proteins bind ubiquitin to facilitate sorting at the trans-Golgi network
.
Nat. Cell Biol.
6
,
252
259
108
Krause
,
M.
and
Gautreau
,
A.
(
2014
)
Steering cell migration: lamellipodium dynamics and the regulation of directional persistence
.
Nat. Rev. Mol. Cell Biol.
15
,
577
590
109
Ridley
,
A.J.
(
2015
)
Rho GTPase signalling in cell migration
.
Curr. Opin. Cell Biol.
36
,
103
112
110
Palamidessi
,
A.
,
Frittoli
,
E.
,
Garré
,
M.
,
Faretta
,
M.
,
Mione
,
M.
,
Testa
,
I.
et al (
2008
)
Endocytic trafficking of Rac is required for the spatial restriction of signaling in cell migration
.
Cell
134
,
135
147
111
Snyder
,
J.T.
,
Rossman
,
K.L.
,
Baumeister
,
M.A.
,
Pruitt
,
W.M.
,
Siderovski
,
D.P.
,
Der
,
C.J.
et al (
2001
)
Quantitative analysis of the effect of phosphoinositide interactions on the function of Dbl family proteins
.
J. Biol. Chem.
276
,
45868
45875
112
Huotari
,
J.
and
Helenius
,
A.
(
2011
)
Endosome maturation
.
EMBO J.
30
,
3481
3500
113
Bilanges
,
B.
,
Posor
,
Y.
and
Vanhaesebroeck
,
B.
(
2019
)
PI3K isoforms in cell signalling and vesicle trafficking
.
Nat. Rev. Mol. Cell Biol.
20
,
515
534
114
Lilja
,
J.
,
Zacharchenko
,
T.
,
Georgiadou
,
M.
,
Jacquemet
,
G.
,
De Franceschi
,
N.
,
Peuhu
,
E.
et al (
2017
)
SHANK proteins limit integrin activation by directly interacting with Rap1 and R-Ras
.
Nat. Cell Biol.
19
,
292
305
115
Goldfinger
,
L.E.
,
Ptak
,
C.
,
Jeffery
,
E.D.
,
Shabanowitz
,
J.
,
Hunt
,
D.F.
and
Ginsberg
,
M.H.
(
2006
)
RLIP76 (RalBP1) is an R-Ras effector that mediates adhesion-dependent Rac activation and cell migration
.
J. Cell Biol.
174
,
877
888
116
Torres
,
V.A.
,
Mielgo
,
A.
,
Barbero
,
S.
,
Hsiao
,
R.
,
Wilkins
,
J.A.
and
Stupack
,
D.G.
(
2010
)
Rab5 mediates caspase-8-promoted cell motility and metastasis
.
Mol. Biol. Cell
21
,
369
376
117
Atherton
,
P.
,
Lausecker
,
F.
,
Harrison
,
A.
and
Ballestrem
,
C.
(
2017
)
Low-intensity pulsed ultrasound promotes cell motility through vinculin-controlled Rac1 GTPase activity
.
J. Cell Sci.
130
,
2277
2291
118
Arriagada
,
C.
,
Silva
,
P.
,
Millet
,
M.
,
Solano
,
L.
,
Moraga
,
C.
and
Torres
,
V.A.
(
2019
)
Focal adhesion kinase-dependent activation of the early endocytic protein Rab5 is associated with cell migration
.
J. Biol. Chem.
294
,
12836
12845
119
Mellor
,
P.
,
Marshall
,
J.D.S.
,
Ruan
,
X.
,
Whitecross
,
D.E.
,
Ross
,
R.L.
,
Knowles
,
M.A.
et al (
2018
)
Patient-derived mutations within the N-terminal domains of p85α impact PTEN or Rab5 binding and regulation
.
Sci. Rep.
8
,
7108
120
Kleinschmidt
,
E.G.
and
Schlaepfer
,
D.D.
(
2017
)
Focal adhesion kinase signaling in unexpected places
.
Curr. Opin. Cell Biol.
45
,
24
30
121
Alanko
,
J.
,
Mai
,
A.
,
Jacquemet
,
G.
,
Schauer
,
K.
,
Kaukonen
,
R.
,
Saari
,
M.
et al (
2015
)
Integrin endosomal signalling suppresses anoikis
.
Nat. Cell Biol.
17
,
1412
1421
122
Lawson
,
C.
,
Lim
,
S.T.
,
Uryu
,
S.
,
Chen
,
X.L.
,
Calderwood
,
D.A.
and
Schlaepfer
,
D.D.
(
2012
)
FAK promotes recruitment of talin to nascent adhesions to control cell motility
.
J. Cell Biol.
196
,
223
232
123
Nader
,
G.P.
,
Ezratty
,
E.J.
and
Gundersen
,
G.G.
(
2016
)
Talin and PIPKIγ regulate endocytosed integrin activation to polarize focal adhesion assembly
.
Nat. Cell Biol.
18
,
491
503
124
Cullen
,
P.J.
and
Steinberg
,
F.
(
2018
)
To degrade or not to degrade: mechanisms and significance of endocytic recycling
.
Nat. Rev. Mol. Cell Biol.
19
,
679
696
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