The interaction of talin with the cell membrane is essential for integrin activation and focal adhesion formation

Krishna Chinthalapudi; Erumbi S. Rangarajan; Tina Izard

doi:10.1073/pnas.1806275115

New Research In

Edited by Barry Honig, Howard Hughes Medical Institute and Columbia University, New York, NY, and approved August 23, 2018 (received for review April 11, 2018)

Significance

Vertebrate cell growth, division, locomotion, morphogenesis, and development rely on the dynamic interactions of cells with extracellular matrix components via cell surface complexes termed focal adhesions that are composed of heterodimeric αβ integrin receptors, associated signaling molecules, and the large cytoskeletal protein talin. While it is known that talin activation and binding to β-integrin requires interactions with lipids, little is known regarding the structure and function of inactive vs. activated talin, and what is known is often disputed. Here we report that talin binding to the cell membrane seems necessary for integrin activation and focal adhesion formation, a finding that significantly advances our understanding of integrin activation and might aid the development of novel integrin therapeutic agents.

Abstract

Multicellular organisms have well-defined, tightly regulated mechanisms for cell adhesion. Heterodimeric αβ integrin receptors play central roles in this function and regulate processes for normal cell functions, including signaling, cell migration, and development, binding to the extracellular matrix, and senescence. They are involved in hemostasis and the immune response, participate in leukocyte function, and have biological implications in angiogenesis and cancer. Proper control of integrin activation for cellular communication with the external environment requires several physiological processes. Perturbation of these equilibria may lead to constitutive integrin activation that results in bleeding disorders. Furthermore, integrins play key roles in cancer progression and metastasis in which certain tumor types exhibit higher levels of various integrins. Thus, the integrin-associated signaling complex is important for cancer therapy development. During inside-out signaling, the cytoskeletal protein talin plays a key role in regulating integrin affinity whereby the talin head domain activates integrin by binding to the cytoplasmic tail of β-integrin and acidic membrane phospholipids. To understand the mechanism of integrin activation by talin, we determined the crystal structure of the talin head domain bound to the acidic phospholipid phosphatidylinositol 4,5-bisphosphate (PIP₂), allowing us to design a lipid-binding–deficient talin mutant. Our confocal microscopy with talin knockout cells suggests that the talin–cell membrane interaction seems essential for focal adhesion formation and stabilization. Basal integrin activation in Chinese hamster ovary cells suggests that the lipid-binding–deficient talin mutant inhibits integrin activation. Thus, membrane attachment of talin seems necessary for integrin activation and focal adhesion formation.

Talin is a key player in integrin activation. Vertebrates express two isoforms in which talin1 is ubiquitously expressed, while talin2 is found primarily in striated muscle and in the brain. As a multidomain cytoskeletal protein, talin contains discrete binding sites for acidic phospholipids, β-integrin, actin, and vinculin, as well as layilin, PIPK1γ90, and synemin. Talin links microfilaments to the cytoplasmic membrane at cell-extracellular matrix adhesion sites. This process depends critically on talin. Talin consists of a polypeptide chain of 2,541 amino acids and is often described as having an N-terminal FERM (four-point-one, ezrin, radixin, moesin) domain connected by a linker (residues 401–482) that harbors a calpain-II cleavage site to a large “rod” domain (residues 483–2,541). The talin head domain is different from all other FERM domain-containing proteins in that it has four subdomains, F0–F3 (instead of the typical three, F1–F3), and they adopt an extended structure (1) instead of the canonical cloverleaf conformation seen in the ERM family of proteins (2). As seen in other FERM domain-containing proteins, the talin FERM subdomains contain a ubiquitin-like F1, acyl-CoA–binding protein-like F2 and phosphotyrosine-binding–like F3 subdomain. Unlike all other F1 FERM subdomains, the talin F1 subdomain has an unstructured insert (F1 loop, residues 133–165, harboring two major phosphorylation sites, T144 and T150) (3), and the talin preceding F0 subdomain has a ubiquitin-like fold. The talin F3 subdomain harbors the primary β-integrin–binding site (4, 5). The talin rod domain consists of 13 domains, R1–R13, composed of 62 amphipathic α-helices arranged into four-helix (R2, R3, R4, and R8) and five-helix (R1, R5, R6, R7, R9, R10, R11, R12, and R13) bundle domains (6, 7) and a C-terminal dimerization domain (6). Each domain has unique properties, including binding to other talin domains, to integrin, and to vinculin (8, 9). A secondary integrin-binding site in the rod domain (residues 1,974–2,293) of uncertain function has also been identified (10). There are three actin-binding sites located on the head domain and R4–R8 and R13 subdomains (6, 11, 12), and 13 vinculin-binding sites that are single amphipathic α-helices (7, 13 ⇓⇓⇓⇓⇓⇓–20).

The role of lipids in integrin activation remains unclear despite a large body of literature and the known functional importance of talin attachment to the membrane (8). In the first stages of cell attachment, the talin F3 FERM domain binds to the NPxY motif of the integrin cytoplasmic β tail, thereby inducing reorganization of the integrin heterodimer and activating integrin (5, 21 ⇓⇓–24). Talin attachment to the plasma membrane is enhanced by phosphatidylinositol 4,5-bisphosphate (PIP₂), which induces a conformational change in talin to expose the integrin-binding site (22, 25 ⇓⇓–28). The role of PIP₂ in integrin activation is particularly interesting since PIP₂ is a major phosphoinositide of the inner membrane (29, 30), and because talin regulates the local PIP₂ concentration in the membrane by binding and activating PIPK1γ (31, 32). PIP₂ regulates important processes, such as vesicular trafficking, platelet activation, cytoskeleton organization (33 ⇓–35), and focal adhesion turnover (25, 26, 36). This process evolves by targeting proteins to the membrane, often through induction of a conformational change or oligomerization (36, 37).

In mammals, the heterodimeric integrin transmembrane receptors are composed of 18 distinct α and β chains (38, 39). By responding to extracellular and intracellular stimuli, integrins connect the extracellular matrix to the cytoskeleton, and transduce signals across the plasma membrane in both directions, termed outside-in and inside-out signaling, respectively (39). Integrin activation is important in platelets and leukocytes as well as many tissues in which extracellular matrix remodeling, angiogenesis, and cell migration are involved. These processes require tightly controlled integrin activation mechanisms that involve conformational changes of these receptors. Thus, understanding the molecular mechanisms of how talin activates integrin is fundamental for gaining insight into important pathological states and recognizing how integrin activation might aid the development of novel integrin antagonists.

Here we report the talin1 head/PIP₂ complex crystal structure together with biochemical and functional data that answer important questions, including how PIP₂ activates talin. Our data provide several surprises and answers to longstanding mechanistic questions and suggest a mechanism in which on recruitment of cytosolic talin by PIPK1γ to the plasma membrane (32), PIP₂ activates talin by severing the head–tail interaction, thereby exposing the integrin-binding site. Remarkably, our in vivo data suggest that the talin–PIP₂ interaction is crucial for talin localization to the cell membrane, affects the scaffolding of cells, and thus is likely key for cell spreading and adhesion. We further find that disrupting talin binding to the membrane affects integrin activation, and that this talin–PIP₂ interaction seems necessary for focal adhesion formation. Collectively, our study provides a major advance in our understanding of the dynamic control of focal adhesions by talin.

Results

PIP₂ Binding to Talin Allosterically Blocks the Integrin and Talin Tail-Binding Sites.

We determined the crystal structures of the N-terminal talin head domain (residues 1–400) and the deletion mutant of that domain Δ139–168 (talin residues 1–400), both bound to PIP₂ (Fig. 1 and SI Appendix, Tables S1 and S2). However, the full-length head domain structure did not show clear electron density for residues 139–168. The electron density map for the lipid was poorer compared with the PIP₂-bound Δ139–168 talin structure.

Fig. 1.

PIP₂ binding to talin allosterically blocks the integrin- and talin tail-binding sites. (A) Superposition of our talin/PIP₂ structure (F2, residues 209–304, green; F3, residues 311–398, blue) onto the talin head domain (residues 209–400; yellow) bound to the tail rod R9 subdomain (residues 1655–1824, cyan; PDB ID code 4F7G) (28). The red double arrow indicates how PIP₂ binding displaces the tail domain. (B) Superposition of our talin/PIP₂ structure (F2, green; F3, blue) onto the two talin2 F2-F3/integrin β1D (integrin, cyan; talin, yellow; PDB ID code 3G9W) (4) heterodimers in the asymmetric unit. The red double arrow indicates how PIP₂ binding prevents the membrane-proximal integrin binding. (C) Superposition of our talin/PIP₂ structure (colored spectrally: F0, residues 4–83, orange; F1, 85–195, yellow; F2, 209–304, green; F3, 311–398, blue; 139-171, disordered) onto the unbound talin1 structure (residues 1–400, Δ139–168, gray; PDB ID code 3IVF) (1) highlights the relative F0-F1 domain movements of approximately 7° and 3 Å.

The talin FERM domain architecture is linear compared with the classical cloverleaf structure. Our PIP₂/talin (1-400; Δ139–168) complex structure harbors one PIP₂-binding site that differs from the classical phosphoinositide-binding mode of other known modules (40), including the ERM protein radixin, where IP₃ bound between the interface between F1 and F3, making the molecular events for membrane-mediated spatiotemporal regulation of talin inhibition and activation unique. The PIP₂-binding site is lined by residues from the F2 (K272) and F3 (K316, K324, E342, and K343) FERM subdomains, in which the 4′-phosphate group of PIP₂ interacts with talin residues K272, E342, and K343 and the 5′-phosphate group interacts with E342 and K316. In addition, K272 interacts with the hydroxyl moiety of the inositol, while one of the carbonyls of the diacylglycerol moiety is within hydrogen-bonding distance to K324.

Superposition of our lipid-bound structure onto the talin head/tail (F2-F3/R9) structure [Protein Data Bank (PDB) ID code 4F7G] (28) reveals a large movement of approximately 10 Å of the F3 loop (residues 318–325) that is extensively involved in binding to the talin rod R9 subdomain as well as to integrin, but not to PIPK1γ (Fig. 1A). In our lipid-bound structure, the 318–325 main chain has temperature factors of approximately 81 Å², with the nearest crystal contacts occurring >4 Å between N323 and the symmetry-related T354. In the head/tail structure, N323 engages in hydrogen-bonding interactions with T1767 and D1809. Strikingly, K324 is approximately 6 Å closer to the lipid-binding site in the talin/PIP₂ structure compared with its position in the head/tail structure. This causes the 318–325 loop to move, whereby K320 releases the talin R9 rod subdomain by steric hindrance. Collectively, the structures show the molecular basis of how the binding of PIP₂ and the talin R9 rod subdomain seem mutually exclusive, particularly since the lipid-binding residues are not involved in crystal contacts (SI Appendix, Fig. S1). This finding is supported by solution studies showing that PIP₂ vesicles compete effectively with talin R9 binding to the talin F2F3 subdomains (28).

Similarly, superposition of our talin/PIP₂ structure onto the talin2 F2-F3/integrin β1D structure (PDB ID code 3G9W) shows that the 318–325 loop adopts a similar conformer in the integrin-bound state and the R9-bound state (Fig. 1B). Notably, the talin L325R mutation abolishes its binding to the integrin membrane-proximal region (4). The position of the loop in our lipid-bound structure causes K321 to occupy the integrin membrane-proximal binding site. In addition, the integrin and the talin R9 rod domain-binding sites overlap on the talin head domain. These findings are consistent with studies showing that the integrin membrane-distal site is necessary for talin-induced integrin activation (41). In our lipid-bound structure, N323 engages only in water-mediated crystal contacts with a symmetry-related T354. Thus, it seems unlikely that the distinct loop conformation is caused by crystal contacts, and more likely that it is caused by lipid binding (SI Appendix, Fig. S2).

The unbound structure (PDB ID code 3IVF) (1) is isomorphous to and almost identical to our PIP₂-bound structure except for relative domain movements. Significantly, superposition of the respective F2F3 domains shows relative F0F1 domain movements of approximately 7° and 3 Å (Fig. 1C). Notably, the 318–325 loop is in the PIP₂-bound conformer in the apo structure. PIP₂ also engages in crystal contacts with K334 and E335, and the E335 side chain moves to make room for the lipid (SI Appendix, Fig. S3). Collectively, this suggests that integrin or R9 binding causes the distinct 318–325 loop conformation.

The interpretation of comparisons with the PIPK1γ/talin structures (PDB ID code 2G35) (42) is less obvious, perhaps because these structures are from either the talin2 isoform or a talin1-PIPK1γ chimera. Furthermore, PIPK1γ is not in contact with the loop region that binds the integrin membrane-proximal site, but instead overlaps with the integrin membrane-distal site on talin.

The Talin Head Domain Harbors Only One PIP₂-Binding Site.

The F1 loop (residues 133–170) has been shown to be required for integrin activation but not for integrin binding. It can form an α-helix and as an isolated peptide, interacts with lipids. To determine if in the context of the talin head domain there is another lipid-binding site, we mutated K272Q, K316Q, K324Q, E342Q, and K343Q, residues that we had identified as lipid-binding amino acids in our complex crystal structure. We confirmed the integrity of our mutant proteins by thermal denaturation (SI Appendix, Fig. S4). This mutant talin exhibited similar melting temperatures (52.02 ± 0.27 °C) as was seen for wild-type talin (53.94 ± 0.27 °C). Thus, the mutations do not seem to affect the structure of the proteins.

We determined lipid binding via a lipid cosedimentation assay (Fig. 2A), which we used previously to detect micromolar lipid binding (43). The mutant showed approximately 12-fold less binding (as assessed using ImageJ) to the lipid vesicles compared with the wild-type talin.

Fig. 2.

The talin head domain harbors one PIP₂-binding site. (A) Structure-based mutagenesis confirms our talin1 lipid-binding sites by lipid cosedimentation assay. The 3 M mutant (Y377F, R358Q, K357Q) binds lipids as seen for wild type (WT) talin, while the minimal lipid-binding–deficient mutant 5M (K272Q, K316Q, K324Q, E342Q, K343Q) and the lipid-binding-deficient mutant 8M (K272Q, K316Q, K324Q, E342Q, K343Q, Y377F-K357Q-R358Q) show insignificant binding to PIP₂/PC vesicles. P, pellet. S, supernatant. (B) FRET experiment showing the fluorescence emission spectra of 1.5 μM CFP-talin (donor) and 4 μM YFP-talin (acceptor) in the absence and presence of increasing concentrations of PIP₂ micelles on excitation at 414 nm. The ordinate shows the relative fluorescence, and the horizontal axis shows the wavelength (nm) of the fluorescence emission scan plot. 0 and 200 μM PIP₂ are shown in black and red traces, respectively; traces for the other concentrations are colored spectrally. (Top) Wild type was predominantly monomeric up to approximately 20 μM PIP₂ and dimeric for all other four higher PIP₂ concentrations (50, 75, 100, and 200 μM). (Bottom) The lipid-binding–deficient mutant (K357Q, R358Q, Y377F, K272Q, K316Q, K324Q, E342Q, K343Q) does not dimerize at PIP₂ concentrations up to 200 μM.

We found a strong electron-dense feature near the side chain of talin residue R358 that is part of the membrane-distal binding site and has been identified as involved in the interaction with integrin by NMR studies (4). This feature is also near N285 and Q288 from a symmetry-related molecule (SI Appendix, Fig. S5). We initially interpreted this feature as a second PIP₂-binding site, since two PIP₂-binding sites (with affinities of 0.4 μM and 5 μM for PIP₂diC₈) were observed by isothermal titration calorimetry (28). However, another study using a phospholipid bilayer that contained 10% PIP₂ and was immobilized on a Biacore L1 chip found that the second, weaker site resided on F0F1 but did not affect the stoichiometry of the interaction of talin with acidic bilayers, and that the contribution of the second site was therefore negligible (26). Furthermore, our talin K358Q mutation resulted in similar binding to lipid vesicles as seen for wild-type talin (Fig. 2A). Thus, K272Q, K316Q, K324Q, E342Q, K343Q is a bonafide lipid-binding–deficient mutant.

We modeled two phosphates into this electron density. The only non-backbone interaction of the phosphate groups occurs with the guanidinium group of R358. The phosphates are also within hydrogen-bonding distance to the carbonyl of N285 and the amide of Q288 of a symmetry-related molecule (SI Appendix, Fig. S5). With respect to the two reported binding constants, it is interesting to note that the crystallographic twofold generates a talin dimer. A dimer is also detected by fluorescence resonance energy transfer of CFP (donor) and YFP (acceptor) fused wild-type talin proteins, but the same assay does not detect dimerization by our lipid-binding–deficient mutant talin (Fig. 2B).

Talin–PIP₂ Interactions Are Essential for Focal Adhesion Formation.

To elucidate how PIP₂ binding to the talin head domain contributes to focal adhesion formation, we generated mutant and wild-type constructs that were tagged with GFP at the N-terminus. These constructs were expressed in talin knockout cells that lack endogenous talin and do not adhere to the extracellular matrix (12). Exogenous expression of GFP-tagged full-length wild-type talin or the “3M” mutant that resides near the membrane-distal integrin-binding site (K357Q, R358Q, Y377F) rescued focal adhesion formation, cell spreading, and cells clearly displaying focal adhesions connected to prominent actin stress fibers (Fig. 3A). Surprisingly, the minimal lipid-binding–deficient mutant “5M” (K272Q, K316Q, K324Q, E342Q, K343Q) and the lipid-binding–deficient mutant “8M” (K272Q, K316Q, K324Q, E342Q, K343Q, K357Q, R358Q, Y377F) disrupted focal adhesion formation. The structure of both mutants seemed to be preserved, as judged by thermal denaturation (SI Appendix, Fig. S4). Furthermore, cells expressing these constructs showed diffused and chaotic actin stress fibers and much smaller cells, comparable to the talin-null cells. Importantly, large pools of GFP-tagged lipid-binding–deficient mutant talin (5M or 8M) accumulated in the cytosol compared with wild-type or mutant (3M) expressing cells. When we previously mutated our vinculin or metavinculin lipid-binding residues (9, 36, 44), focal adhesions were never completely disassembled; however, the focal adhesions were largely disrupted for cells expressing the lipid-binding–deficient talin mutant. Thus, lipid binding seems necessary for talin localization to the focal adhesion membrane sites and for talin regulation of the scaffolding effects. Furthermore, talin-null cells expressing lipid-binding–deficient talin mutants were approximately fivefold smaller (with a chaotic and diffused actin network) compared with talin-null cells expressing wild-type talin, in which a pronounced F-actin is visible, as the cellular integrity is maintained by intact talin–membrane interactions and proper cytoskeletal rearrangements.

Fig. 3.

Talin–PIP₂ interactions seem to be essential for focal adhesion formation. (A) Talin-null papillary collecting duct cells (PCDs) engineered to express full-length wild-type GFP-talin1 or mutant GFP-talin1 fusion proteins were analyzed by confocal laser scanning microscopy. Representative images, which define the localization of GFP-talin1 (green) at FAs decorating F-actin (red), along with the merged channels, are shown along with the nuclei stained with DAPI. Data shown are representative of five independent experiments. (Scale bars: 2 and 5 μm as indicated.) (B and C) Flow cytometry analyses of (B) PAC1 binding to wild-type talin1 and (C) the minimal lipid-binding–deficient mutant 5M (K272Q, K316Q, K324Q, E342Q, K343Q) represented as FACS dot plots. Abscissas, PAC1 binding; ordinates, full-length talin1 expression. Each experiment was repeated twice. Stably transfected wild-type talin1 increases integrin activation (correlated to PAC1 binding), but the lipid-binding–deficient mutant significantly reduces integrin activation.

Next, to determine the effects of talin binding to the plasma membrane, we assessed the PIP₂-mediated integrin activation in Chinese hamster ovary (CHO) cells that stably expressed integrin (αIIbβ3). When we transiently transfected talin1, we obtained too few cells expressing the protein to perform our integrin activation assays in triplicate. We overcame this by generating stable CHO cells expressing full-length talin1 tagged with EGFP. For each construct, we measured the mean fluorescence intensity (MFI) of the bound ligand in at least two independent experiments. The PAC1 antibody, which recognizes only activated αIIbβ3 receptors, bound to integrin αIIbβ3 with higher affinity in cells expressing wild-type talin1, with an MFI of 13.5% compared with the cell-expressing mutant talin1 (MFI of 1.57%) or the lipid-binding–deficient mutants (MFI of 0.98% for 5M and ≤0.26% for 8M) (Fig. 3 B and C and SI Appendix, Fig. S6). Collectively, our data show that disrupting talin binding to the membrane affects integrin activation.

Since focal adhesion formation was significantly affected by our talin lipid-binding–deficient mutant, we assessed the mobility of talin by FRAP in the talin-null cell background (Fig. 4A). These experiments were not possible with the lipid-binding–deficient talin mutants that lack distinct focal adhesions in which the major pool of GFP-talin proteins are present in the cytosol. The fitting of our full-length talin FRAP data with >99% confidence was possible only with the double exponential (Fig. 4B). The resulting recovery curve revealed that the fluorescence recovery was biphasic with an initial fast-phase t_1/2 of 1.7 ± 0.15 s and a slow-phase t_1/2 of 9.2 ± 0.36 s. Thus, talin is recruited to focal adhesions, and this localization is maintained in cells moving in a persistent manner, allowing determination of the dynamics for the association of talin with the membrane. The initial fast rate of recovery possibly accounts for the bulk of recovery and most likely reflects this rebinding of cytosolic talin to focal adhesions at the plasma membrane. A relatively small amount of recovery also may have occurred by lateral diffusion from the adjacent membrane, which would be consistent with the second observed, slower t_1/2. Collectively, talin binding to the membrane is a dynamic process important for its scaffolding effects at the focal adhesion membrane sites.

Fig. 4.

Talin1 influences focal adhesion dynamics at the plasma membrane. (A) Representative images of FRAP recovery of EGFP-tagged wild-type full-length talin1. Focal adhesions are indicated before and after photobleaching (arrows). (B) FRAP recovery curve of talin1 in talin^−/− PCD cells. A double-exponential model was used to fit these normalized fluorescence curves. The red line is the calculated curve that fits the experimental data and is the best fit of a nonlinear regression analysis with >99% confidence. The results represent the mean ± SEM of 20 independent measurements. Error bars are shown in the form of bands (light gray) to represent the SEM. (Scale bar: 5 μm.)

Discussion

We determined the talin head/PIP₂ complex crystal structure and confirmed the lipid-binding site seen in the crystal biochemically. Based on sequence similarity and mutagenesis, one of the bonafide residues involved in PIP₂ binding, K272, has previously been identified as the “membrane orientation patch.” Other postulated lipid-binding residues (K256, K274, and R277) (4) are not part of the lipid-binding site. Our structural and biochemical identification of K324 as another bonafide PIP₂-binding residue also agrees with NMR studies revealing a perturbation in the 268–278 and 318–324 regions on binding to IP₃ (28). Surface plasmon resonance data showed that the K324A mutant bound PIP₂ sixfold weaker compared with wild type. Molecular dynamics simulations predicted that K324 might be in hydrogen-bonding interactions with PIP₂ in addition to being in contact with residue R995 from the integrin α subunit and thus releasing the electrostatic interaction with D723 from the integrin β subunit. However, in the absence of our high-resolution talin/PIP₂ structure, functional studies had been difficult to interpret. Here we mutated the five lipid-binding residues that we confirmed structurally and biochemically to be involved in PIP₂ binding: K272Q, K316Q, K324Q, E342Q, and K343Q. We found that the talin–lipid interaction seems to be essential for focal adhesion formation and stabilization, and that this interaction increases integrin activation.

As shown in our high-resolution confocal imaging studies, the K357Q, R358Q, and Y377F talin mutant that targets the integrin membrane-distal binding site does not affect focal adhesion formation (Fig. 3A). Comparison of our lipid-bound structure with the integrin-bound talin structure revealed a movement of the side chain of R358 to stack with integrin W775, while K357 is in electrostatic interaction with integrin E779 at the membrane-distal region. In contrast, the lipid-binding–deficient mutant affected both focal adhesion formation and integrin activation (Fig. 3). Notably, the distal integrin membrane interaction with talin has been shown to provide the initial linkage between talin and integrin, while strong activation arises from the subsequent binding of talin to the integrin membrane-proximal region (5). Our studies suggest that both integrin-binding sites are involved in integrin activation, while only the membrane-proximal site is involved in focal adhesion formation.

The majority of previous in vivo studies have used the transiently transfected talin head domain (residues 1–400) or just the F2F3 (residues 203–400) or F3 (residues 309–400) talin FERM subdomain (5, 24), which mimic the activated talin form. In contrast, we used full-length inactive talin to measure integrin activation by stably expressing talin proteins and selective sorting via two rounds of fluorescent-activated cell sorting (FACS) before performing the integrin activation assays. These stable pools of talin-expressing cells enabled us to reliably and reproducibly measure integrin activation rates using A5 CHO cells.

PIP₂ activates talin by severing the head–tail interaction, thereby exposing the integrin-binding site, although simultaneous binding of talin to the integrin membrane-proximal site and to PIP₂ seem mutually exclusive. Mutating talin residues involved in binding to the integrin membrane-distal site did not affect focal adhesion formation (Fig. 3A), while integrin activation was reduced by approximately 8.5-fold compared with wild type (Fig. 3B and SI Appendix, Fig. S6). In contrast, the lipid-binding–deficient mutant affected both focal adhesion formation and integrin activation. This suggests that the integrin membrane-proximal region plays a role in integrin activation and focal adhesions, whereas the membrane-distal region impacts only integrin activation.

Talin recruitment to the membrane leads to integrin activation. These sequential events are closely linked because integrin activation requires the talin head domain to be positioned close to the integrin tail on the cytoplasmic face of the membrane. Talin-mediated integrin activation requires that the autoinhibitory interactions between the talin head and rod domains are released by PIP₂. The autoinhibitory talin F2F3/R9 structure identified the head–tail interface and suggested a negatively charged surface to repel the membrane, although the orientation of the lipid was unknown. However, our talin/PIP₂ structure shows that this negatively charged surface is not planar, but rather is almost perpendicular to the PIP₂-binding site (SI Appendix, Fig. S7). It remains to be seen what surfaces on full-length talin are actually solvent-exposed.

The unique F1 loop has been suggested to become helical when in contact with PIP₂-rich microdomains to decrease the distance between talin and the plasma membrane. Furthermore, two of the talin phosphorylation sites that have been mapped from activated human platelets (3) are located on this loop (T144 and T150). This suggests that phosphorylation of these sites might prevent talin–membrane interactions. However, our lipid cosedimentation results showed that the lipid-binding–deficient talin, which has an intact F1 loop, does not bind to lipids. This suggests that in the context of the entire talin head domain (residues 1–400), this loop does not interact with the membrane. It remains to be seen if the second integrin-binding site located in the talin tail domain (residues 1984–2,113) allows for simultaneous binding of full-length talin to the membrane via F2F3 and to integrin via the second integrin-binding site.

While the talin–integrin interaction was shown to be enhanced by PIP₂ (31), binding of integrin via its membrane-proximal site and PIP₂ seem to be mutually exclusive (Fig. 1B). In agreement with the earlier studies, we show how PIP₂ severs the talin head–tail interaction to expose the cryptic integrin-binding site. Collectively, these findings indicate that on talin recruitment to the cell membrane, lipid binding to the talin head domain releases the interaction of the talin head with the talin rod domains. Activated talin can then activate integrin, which releases the interaction of talin with the membrane. It remains to be seen if the integrin membrane-proximal binding site is solvent-exposed in the full-length talin structure.

Experimental Procedures

DNA Constructs and Protein Preparation.

All bacterial expression plasmids and mammalian expression plasmids of talin1 used in this study were cloned using mouse full-length talin1 as a template. Site-directed mutagenesis was performed to generate the talin1 mutants, and all DNA constructs were sequence-verified. Proteins were expressed in Rosetta 2 or BL21-CodonPlus (DE3)-RIL host cells and purified by nickel affinity and size exclusion chromatography.

In Vitro and in Vivo Functional Assays and Confocal Microscopy.

The talin1 head domain proteins were used for in vitro FRET assays as CFP and YFP FRET donor and acceptor pairs. In brief, FRET measurements were performed for 1.5 μM CFP-talin1 and 4 μM YFP-talin1 wild-type and mutant proteins in the absence and presence of increasing concentrations of PIP₂ micelles (0–200 μM).

High-resolution confocal microscopy and FRAP experiments were performed with talin-null epithelial cells. EGFP-tagged full-length talin1 proteins were transfected in A5 CHO cells that stably expressed integrin (αIIbβ3) receptors, and basal integrin activation assays were performed by FACS using A5 CHO cells.

X-Ray Crystallography.

Lipid-bound talin1 head domain (residues 1–400; Δ139–168) crystals were obtained by hanging-drop vapor diffusion by optimizing the PIP₂diC₈-to-talin ratio. X-ray diffraction data were collected at the Stanford Synchrotron Radiation Lightsource, beamline 12–2, and the X-ray diffraction data were indexed, integrated, and scaled using XDS and AIMLESS as implemented in autoPROC (45). The unbound talin1 structure (PDB ID code 3IVF) (1) was used to obtain phases for our talin/PIP₂ complex, and crystallographic refinement was performed using autoBuster (46).

Acknowledgments

We are indebted to the staff of the Stanford Synchrotron Radiation Lightsource for synchrotron support. We thank the Max Planck Florida Light Microscopy facility and Florida Atlantic University, Nikon Center of Excellence for imaging facilities, Dr. Roy Zent (Vanderbilt Center for Kidney Disease) for talin-null cells, and Dr. Mark Ginsberg (University of California, San Diego) for A5 CHO cells. We thank Marina Primi [The Scripps Research Institute (TSRI)] and Charmane Gabriel (Oxbridge Academy) for protein expression and Louis Shane (Palm Beach Gardens, FL) and Douglas Bingham (TSRI) for helpful discussions regarding the manuscript. T.I. is supported by grants from the National Institute of Health and the Department of Defense, and by startup funds provided to TSRI from the State of Florida. This is publication 29675 from The Scripps Research Institute.

Footnotes

↵¹To whom correspondence should be addressed. Email: cmorrow{at}scripps.edu.

Author contributions: K.C., E.S.R., and T.I. designed research; K.C. and E.S.R. performed research; K.C., E.S.R., and T.I. analyzed data; and K.C., E.S.R., and T.I. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The atomic coordinates and structure factors have been deposited in the Protein Data Bank, www.wwpdb.org (PDB ID code 6mfs).
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1806275115/-/DCSupplemental.

Published under the PNAS license.

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2. Chambers D,
3. Nguyen R,
4. Reczek D
(2000) ERM-Merlin and EBP50 protein families in plasma membrane organization and function. Annu Rev Cell Dev Biol 16:113–143.
OpenUrl CrossRef PubMed
↵
1. Ratnikov B, et al.
(2005) Talin phosphorylation sites mapped by mass spectrometry. J Cell Sci 118:4921–4923.
OpenUrl FREE Full Text
↵
1. Anthis NJ, et al.
(2009) The structure of an integrin/talin complex reveals the basis of inside-out signal transduction. EMBO J 28:3623–3632.
OpenUrl Abstract/FREE Full Text
↵
1. Wegener KL, et al.
(2007) Structural basis of integrin activation by talin. Cell 128:171–182.
OpenUrl CrossRef PubMed
↵
1. Gingras AR, et al.
(2008) The structure of the C-terminal actin-binding domain of talin. EMBO J 27:458–469.
OpenUrl Abstract/FREE Full Text
↵
1. Papagrigoriou E, et al.
(2004) Activation of a vinculin-binding site in the talin rod involves rearrangement of a five-helix bundle. EMBO J 23:2942–2951.
OpenUrl Abstract/FREE Full Text
↵
1. Brown DT,
2. Izard T
(2015) Vinculin-cell membrane interactions. Oncotarget 6:34043–34044.
OpenUrl PubMed
↵
1. Izard T,
2. Brown DT
(2016) Mechanisms and functions of vinculin interactions with phospholipids at cell adhesion sites. J Biol Chem 291:2548–2555.
OpenUrl Abstract/FREE Full Text
↵
1. Moes M, et al.
(2007) The integrin binding site 2 (IBS2) in the talin rod domain is essential for linking integrin β subunits to the cytoskeleton. J Biol Chem 282:17280–17288.
OpenUrl Abstract/FREE Full Text
↵
1. Hemmings L, et al.
(1996) Talin contains three actin-binding sites each of which is adjacent to a vinculin-binding site. J Cell Sci 109:2715–2726.
OpenUrl Abstract/FREE Full Text
↵
1. Atherton P, et al.
(2015) Vinculin controls talin engagement with the actomyosin machinery. Nat Commun 6:10038.
OpenUrl CrossRef PubMed
↵
1. Bois PR,
2. Borgon RA,
3. Vonrhein C,
4. Izard T
(2005) Structural dynamics of α-actinin–vinculin interactions. Mol Cell Biol 25:6112–6122.
OpenUrl Abstract/FREE Full Text
↵
1. Fillingham I, et al.
(2005) A vinculin binding domain from the talin rod unfolds to form a complex with the vinculin head. Structure 13:65–74.
OpenUrl CrossRef PubMed
↵
1. Izard T,
2. Vonrhein C
(2004) Structural basis for amplifying vinculin activation by talin. J Biol Chem 279:27667–27678.
OpenUrl Abstract/FREE Full Text
↵
1. Nhieu GT,
2. Izard T
(2007) Vinculin binding in its closed conformation by a helix addition mechanism. EMBO J 26:4588–4596.
OpenUrl Abstract/FREE Full Text
↵
1. Yogesha SD,
2. Rangarajan ES,
3. Vonrhein C,
4. Bricogne G,
5. Izard T
(2012) Crystal structure of vinculin in complex with vinculin binding site 50 (VBS50), the integrin binding site 2 (IBS2) of talin. Protein Sci 21:583–588.
OpenUrl CrossRef PubMed
↵
1. Yogesha SD,
2. Sharff A,
3. Bricogne G,
4. Izard T
(2011) Intermolecular versus intramolecular interactions of the vinculin binding site 33 of talin. Protein Sci 20:1471–1476.
OpenUrl CrossRef PubMed
↵
1. Izard T, et al.
(2004) Vinculin activation by talin through helical bundle conversion. Nature 427:171–175.
OpenUrl CrossRef PubMed
↵
1. Bois PR,
2. O’Hara BP,
3. Nietlispach D,
4. Kirkpatrick J,
5. Izard T
(2006) The vinculin binding sites of talin and α-actinin are sufficient to activate vinculin. J Biol Chem 281:7228–7236.
OpenUrl Abstract/FREE Full Text
↵
1. Tadokoro S, et al.
(2003) Talin binding to integrin β tails: A final common step in integrin activation. Science 302:103–106.
OpenUrl Abstract/FREE Full Text
↵
1. Critchley DR
(2009) Biochemical and structural properties of the integrin-associated cytoskeletal protein talin. Annu Rev Biophys 38:235–254.
OpenUrl CrossRef PubMed
↵
1. García-Alvarez B, et al.
(2003) Structural determinants of integrin recognition by talin. Mol Cell 11:49–58.
OpenUrl CrossRef PubMed
↵
1. Vinogradova O, et al.
(2002) A structural mechanism of integrin α(IIb)β(3) “inside-out” activation as regulated by its cytoplasmic face. Cell 110:587–597.
OpenUrl CrossRef PubMed
↵
1. Saltel F, et al.
(2009) New PI(4,5)P2- and membrane-proximal integrin-binding motifs in the talin head control beta3-integrin clustering. J Cell Biol 187:715–731.
OpenUrl Abstract/FREE Full Text
↵
1. Moore DT, et al.
(2012) Affinity of talin-1 for the β3-integrin cytosolic domain is modulated by its phospholipid bilayer environment. Proc Natl Acad Sci USA 109:793–798.
OpenUrl Abstract/FREE Full Text
↵
1. Goksoy E, et al.
(2008) Structural basis for the autoinhibition of talin in regulating integrin activation. Mol Cell 31:124–133.
OpenUrl CrossRef PubMed
↵
1. Song X, et al.
(2012) A novel membrane-dependent on/off switch mechanism of talin FERM domain at sites of cell adhesion. Cell Res 22:1533–1545.
OpenUrl CrossRef PubMed
↵
1. McLaughlin S,
2. Wang J,
3. Gambhir A,
4. Murray D
(2002) PIP(2) and proteins: Interactions, organization, and information flow. Annu Rev Biophys Biomol Struct 31:151–175.
OpenUrl CrossRef PubMed
↵
1. Xu C,
2. Watras J,
3. Loew LM
(2003) Kinetic analysis of receptor-activated phosphoinositide turnover. J Cell Biol 161:779–791.
OpenUrl Abstract/FREE Full Text
↵
1. Ling K,
2. Doughman RL,
3. Firestone AJ,
4. Bunce MW,
5. Anderson RA
(2002) Type I γ phosphatidylinositol phosphate kinase targets and regulates focal adhesions. Nature 420:89–93.
OpenUrl CrossRef PubMed
↵
1. Di Paolo G, et al.
(2002) Recruitment and regulation of phosphatidylinositol phosphate kinase type 1 γ by the FERM domain of talin. Nature 420:85–89.
OpenUrl CrossRef PubMed
↵
1. Sun Y,
2. Thapa N,
3. Hedman AC,
4. Anderson RA
(2013) Phosphatidylinositol 4,5-bisphosphate: Targeted production and signaling. BioEssays 35:513–522.
OpenUrl CrossRef PubMed
↵
1. Kwiatkowska K
(2010) One lipid, multiple functions: How various pools of PI(4,5)P(2) are created in the plasma membrane. Cell Mol Life Sci 67:3927–3946.
OpenUrl CrossRef PubMed
↵
1. van den Bout I,
2. Divecha N
(2009) PIP5K-driven PtdIns(4,5)P2 synthesis: Regulation and cellular functions. J Cell Sci 122:3837–3850.
OpenUrl Abstract/FREE Full Text
↵
1. Chinthalapudi K, et al.
(2014) Lipid binding promotes oligomerization and focal adhesion activity of vinculin. J Cell Biol 207:643–656.
OpenUrl Abstract/FREE Full Text
↵
1. Hirao M, et al.
(1996) Regulation mechanism of ERM (ezrin/radixin/moesin) protein/plasma membrane association: Possible involvement of phosphatidylinositol turnover and Rho-dependent signaling pathway. J Cell Biol 135:37–51.
OpenUrl Abstract/FREE Full Text
↵
1. Anthis NJ,
2. Campbell ID
(2011) The tail of integrin activation. Trends Biochem Sci 36:191–198.
OpenUrl CrossRef PubMed
↵
1. Hynes RO
(2002) Integrins: Bidirectional, allosteric signaling machines. Cell 110:673–687.
OpenUrl CrossRef PubMed
↵
1. Kutateladze TG
(2010) Translation of the phosphoinositide code by PI effectors. Nat Chem Biol 6:507–513.
OpenUrl CrossRef PubMed
↵
1. Liu J,
2. Wang Z,
3. Thinn AM,
4. Ma YQ,
5. Zhu J
(2015) The dual structural roles of the membrane-distal region of the α-integrin cytoplasmic tail during integrin inside-out activation. J Cell Sci 128:1718–1731.
OpenUrl Abstract/FREE Full Text
↵
1. Kong X,
2. Wang X,
3. Misra S,
4. Qin J
(2006) Structural basis for the phosphorylation-regulated focal adhesion targeting of type Igamma phosphatidylinositol phosphate kinase (PIPKIgamma) by talin. J Mol Biol 359:47–54.
OpenUrl CrossRef PubMed
↵
1. Chinthalapudi K, et al.
(2018) Lipid binding promotes the open conformation and tumor-suppressive activity of neurofibromin 2. Nat Commun 9:1338.
OpenUrl
↵
1. Chinthalapudi K,
2. Rangarajan ES,
3. Brown DT,
4. Izard T
(2016) Differential lipid binding of vinculin isoforms promotes quasi-equivalent dimerization. Proc Natl Acad Sci USA 113:9539–9544.
OpenUrl Abstract/FREE Full Text
↵
1. Vonrhein C, et al.
(2011) Data processing and analysis with the autoPROC toolbox. Acta Crystallogr D Biol Crystallogr 67:293–302.
OpenUrl CrossRef PubMed
↵
1. Bricogne G, et al.
(2011) BUSTER version 2.9 (Global Phasing Ltd, Cambridge, UK).

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Proceedings of the National Academy of Sciences: 115 (41)

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Article Classifications

Biological Sciences
Biophysics and Computational Biology

[1] ↵
Elliott PR, et al.
(2010) The structure of the talin head reveals a novel extended conformation of the FERM domain. Structure 18:1289–1299.
OpenUrl CrossRef PubMed

[2] Elliott PR, et al.

[3] ↵
Bretscher A,
Chambers D,
Nguyen R,
Reczek D
(2000) ERM-Merlin and EBP50 protein families in plasma membrane organization and function. Annu Rev Cell Dev Biol 16:113–143.
OpenUrl CrossRef PubMed

[4] Bretscher A,

[5] Chambers D,

[6] Nguyen R,

[7] Reczek D

[8] ↵
Ratnikov B, et al.
(2005) Talin phosphorylation sites mapped by mass spectrometry. J Cell Sci 118:4921–4923.
OpenUrl FREE Full Text

[9] Ratnikov B, et al.

[10] ↵
Anthis NJ, et al.
(2009) The structure of an integrin/talin complex reveals the basis of inside-out signal transduction. EMBO J 28:3623–3632.
OpenUrl Abstract/FREE Full Text

[11] Anthis NJ, et al.

[12] ↵
Wegener KL, et al.
(2007) Structural basis of integrin activation by talin. Cell 128:171–182.
OpenUrl CrossRef PubMed

[13] Wegener KL, et al.

[14] ↵
Gingras AR, et al.
(2008) The structure of the C-terminal actin-binding domain of talin. EMBO J 27:458–469.
OpenUrl Abstract/FREE Full Text

[15] Gingras AR, et al.

[16] ↵
Papagrigoriou E, et al.
(2004) Activation of a vinculin-binding site in the talin rod involves rearrangement of a five-helix bundle. EMBO J 23:2942–2951.
OpenUrl Abstract/FREE Full Text

[17] Papagrigoriou E, et al.

[18] ↵
Brown DT,
Izard T
(2015) Vinculin-cell membrane interactions. Oncotarget 6:34043–34044.
OpenUrl PubMed

[19] Brown DT,

[20] Izard T

[21] ↵
Izard T,
Brown DT
(2016) Mechanisms and functions of vinculin interactions with phospholipids at cell adhesion sites. J Biol Chem 291:2548–2555.
OpenUrl Abstract/FREE Full Text

[22] Izard T,

[23] Brown DT

[24] ↵
Moes M, et al.
(2007) The integrin binding site 2 (IBS2) in the talin rod domain is essential for linking integrin β subunits to the cytoskeleton. J Biol Chem 282:17280–17288.
OpenUrl Abstract/FREE Full Text

[25] Moes M, et al.

[26] ↵
Hemmings L, et al.
(1996) Talin contains three actin-binding sites each of which is adjacent to a vinculin-binding site. J Cell Sci 109:2715–2726.
OpenUrl Abstract/FREE Full Text

[27] Hemmings L, et al.

[28] ↵
Atherton P, et al.
(2015) Vinculin controls talin engagement with the actomyosin machinery. Nat Commun 6:10038.
OpenUrl CrossRef PubMed

[29] Atherton P, et al.

[30] ↵
Bois PR,
Borgon RA,
Vonrhein C,
Izard T
(2005) Structural dynamics of α-actinin–vinculin interactions. Mol Cell Biol 25:6112–6122.
OpenUrl Abstract/FREE Full Text

[31] Bois PR,

[32] Borgon RA,

[33] Vonrhein C,

[34] Izard T

[35] ↵
Fillingham I, et al.
(2005) A vinculin binding domain from the talin rod unfolds to form a complex with the vinculin head. Structure 13:65–74.
OpenUrl CrossRef PubMed

[36] Fillingham I, et al.

[37] ↵
Izard T,
Vonrhein C
(2004) Structural basis for amplifying vinculin activation by talin. J Biol Chem 279:27667–27678.
OpenUrl Abstract/FREE Full Text

[38] Izard T,

[39] Vonrhein C

[40] ↵
Nhieu GT,
Izard T
(2007) Vinculin binding in its closed conformation by a helix addition mechanism. EMBO J 26:4588–4596.
OpenUrl Abstract/FREE Full Text

[41] Nhieu GT,

[42] Izard T

[43] ↵
Yogesha SD,
Rangarajan ES,
Vonrhein C,
Bricogne G,
Izard T
(2012) Crystal structure of vinculin in complex with vinculin binding site 50 (VBS50), the integrin binding site 2 (IBS2) of talin. Protein Sci 21:583–588.
OpenUrl CrossRef PubMed

[44] Yogesha SD,

[45] Rangarajan ES,

[46] Vonrhein C,

[47] Bricogne G,

[48] Izard T

[49] ↵
Yogesha SD,
Sharff A,
Bricogne G,
Izard T
(2011) Intermolecular versus intramolecular interactions of the vinculin binding site 33 of talin. Protein Sci 20:1471–1476.
OpenUrl CrossRef PubMed

[50] Yogesha SD,

[51] Sharff A,

[52] Bricogne G,

[53] Izard T

[54] ↵
Izard T, et al.
(2004) Vinculin activation by talin through helical bundle conversion. Nature 427:171–175.
OpenUrl CrossRef PubMed

[55] Izard T, et al.

[56] ↵
Bois PR,
O’Hara BP,
Nietlispach D,
Kirkpatrick J,
Izard T
(2006) The vinculin binding sites of talin and α-actinin are sufficient to activate vinculin. J Biol Chem 281:7228–7236.
OpenUrl Abstract/FREE Full Text

[57] Bois PR,

[58] O’Hara BP,

[59] Nietlispach D,

[60] Kirkpatrick J,

[61] Izard T

[62] ↵
Tadokoro S, et al.
(2003) Talin binding to integrin β tails: A final common step in integrin activation. Science 302:103–106.
OpenUrl Abstract/FREE Full Text

[63] Tadokoro S, et al.

[64] ↵
Critchley DR
(2009) Biochemical and structural properties of the integrin-associated cytoskeletal protein talin. Annu Rev Biophys 38:235–254.
OpenUrl CrossRef PubMed

[65] Critchley DR

[66] ↵
García-Alvarez B, et al.
(2003) Structural determinants of integrin recognition by talin. Mol Cell 11:49–58.
OpenUrl CrossRef PubMed

[67] García-Alvarez B, et al.

[68] ↵
Vinogradova O, et al.
(2002) A structural mechanism of integrin α(IIb)β(3) “inside-out” activation as regulated by its cytoplasmic face. Cell 110:587–597.
OpenUrl CrossRef PubMed

[69] Vinogradova O, et al.

[70] ↵
Saltel F, et al.
(2009) New PI(4,5)P2- and membrane-proximal integrin-binding motifs in the talin head control beta3-integrin clustering. J Cell Biol 187:715–731.
OpenUrl Abstract/FREE Full Text

[71] Saltel F, et al.

[72] ↵
Moore DT, et al.
(2012) Affinity of talin-1 for the β3-integrin cytosolic domain is modulated by its phospholipid bilayer environment. Proc Natl Acad Sci USA 109:793–798.
OpenUrl Abstract/FREE Full Text

[73] Moore DT, et al.

[74] ↵
Goksoy E, et al.
(2008) Structural basis for the autoinhibition of talin in regulating integrin activation. Mol Cell 31:124–133.
OpenUrl CrossRef PubMed

[75] Goksoy E, et al.

[76] ↵
Song X, et al.
(2012) A novel membrane-dependent on/off switch mechanism of talin FERM domain at sites of cell adhesion. Cell Res 22:1533–1545.
OpenUrl CrossRef PubMed

[77] Song X, et al.

[78] ↵
McLaughlin S,
Wang J,
Gambhir A,
Murray D
(2002) PIP(2) and proteins: Interactions, organization, and information flow. Annu Rev Biophys Biomol Struct 31:151–175.
OpenUrl CrossRef PubMed

[79] McLaughlin S,

[80] Wang J,

[81] Gambhir A,

[82] Murray D

[83] ↵
Xu C,
Watras J,
Loew LM
(2003) Kinetic analysis of receptor-activated phosphoinositide turnover. J Cell Biol 161:779–791.
OpenUrl Abstract/FREE Full Text

[84] Xu C,

[85] Watras J,

[86] Loew LM

[87] ↵
Ling K,
Doughman RL,
Firestone AJ,
Bunce MW,
Anderson RA
(2002) Type I γ phosphatidylinositol phosphate kinase targets and regulates focal adhesions. Nature 420:89–93.
OpenUrl CrossRef PubMed

[88] Ling K,

[89] Doughman RL,

[90] Firestone AJ,

[91] Bunce MW,

[92] Anderson RA

[93] ↵
Di Paolo G, et al.
(2002) Recruitment and regulation of phosphatidylinositol phosphate kinase type 1 γ by the FERM domain of talin. Nature 420:85–89.
OpenUrl CrossRef PubMed

[94] Di Paolo G, et al.

[95] ↵
Sun Y,
Thapa N,
Hedman AC,
Anderson RA
(2013) Phosphatidylinositol 4,5-bisphosphate: Targeted production and signaling. BioEssays 35:513–522.
OpenUrl CrossRef PubMed

[96] Sun Y,

[97] Thapa N,

[98] Hedman AC,

[99] Anderson RA

[100] ↵
Kwiatkowska K
(2010) One lipid, multiple functions: How various pools of PI(4,5)P(2) are created in the plasma membrane. Cell Mol Life Sci 67:3927–3946.
OpenUrl CrossRef PubMed

[101] Kwiatkowska K

[102] ↵
van den Bout I,
Divecha N
(2009) PIP5K-driven PtdIns(4,5)P2 synthesis: Regulation and cellular functions. J Cell Sci 122:3837–3850.
OpenUrl Abstract/FREE Full Text

[103] van den Bout I,

[104] Divecha N

[105] ↵
Chinthalapudi K, et al.
(2014) Lipid binding promotes oligomerization and focal adhesion activity of vinculin. J Cell Biol 207:643–656.
OpenUrl Abstract/FREE Full Text

[106] Chinthalapudi K, et al.

[107] ↵
Hirao M, et al.
(1996) Regulation mechanism of ERM (ezrin/radixin/moesin) protein/plasma membrane association: Possible involvement of phosphatidylinositol turnover and Rho-dependent signaling pathway. J Cell Biol 135:37–51.
OpenUrl Abstract/FREE Full Text

[108] Hirao M, et al.

[109] ↵
Anthis NJ,
Campbell ID
(2011) The tail of integrin activation. Trends Biochem Sci 36:191–198.
OpenUrl CrossRef PubMed

[110] Anthis NJ,

[111] Campbell ID

[112] ↵
Hynes RO
(2002) Integrins: Bidirectional, allosteric signaling machines. Cell 110:673–687.
OpenUrl CrossRef PubMed

[113] Hynes RO

[114] ↵
Kutateladze TG
(2010) Translation of the phosphoinositide code by PI effectors. Nat Chem Biol 6:507–513.
OpenUrl CrossRef PubMed

[115] Kutateladze TG

[116] ↵
Liu J,
Wang Z,
Thinn AM,
Ma YQ,
Zhu J
(2015) The dual structural roles of the membrane-distal region of the α-integrin cytoplasmic tail during integrin inside-out activation. J Cell Sci 128:1718–1731.
OpenUrl Abstract/FREE Full Text

[117] Liu J,

[118] Wang Z,

[119] Thinn AM,

[120] Ma YQ,

[121] Zhu J

[122] ↵
Kong X,
Wang X,
Misra S,
Qin J
(2006) Structural basis for the phosphorylation-regulated focal adhesion targeting of type Igamma phosphatidylinositol phosphate kinase (PIPKIgamma) by talin. J Mol Biol 359:47–54.
OpenUrl CrossRef PubMed

[123] Kong X,

[124] Wang X,

[125] Misra S,

[126] Qin J

[127] ↵
Chinthalapudi K, et al.
(2018) Lipid binding promotes the open conformation and tumor-suppressive activity of neurofibromin 2. Nat Commun 9:1338.
OpenUrl

[128] Chinthalapudi K, et al.

[129] ↵
Chinthalapudi K,
Rangarajan ES,
Brown DT,
Izard T
(2016) Differential lipid binding of vinculin isoforms promotes quasi-equivalent dimerization. Proc Natl Acad Sci USA 113:9539–9544.
OpenUrl Abstract/FREE Full Text

[130] Chinthalapudi K,

[131] Rangarajan ES,

[132] Brown DT,

[133] Izard T

[134] ↵
Vonrhein C, et al.
(2011) Data processing and analysis with the autoPROC toolbox. Acta Crystallogr D Biol Crystallogr 67:293–302.
OpenUrl CrossRef PubMed

[135] Vonrhein C, et al.

[136] ↵
Bricogne G, et al.
(2011) BUSTER version 2.9 (Global Phasing Ltd, Cambridge, UK).

[137] Bricogne G, et al.

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The interaction of talin with the cell membrane is essential for integrin activation and focal adhesion formation

Significance

Abstract

Results

PIP₂ Binding to Talin Allosterically Blocks the Integrin and Talin Tail-Binding Sites.

The Talin Head Domain Harbors Only One PIP₂-Binding Site.

Talin–PIP₂ Interactions Are Essential for Focal Adhesion Formation.

Discussion

Experimental Procedures

DNA Constructs and Protein Preparation.

In Vitro and in Vivo Functional Assays and Confocal Microscopy.

X-Ray Crystallography.

Acknowledgments

Footnotes

References

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The interaction of talin with the cell membrane is essential for integrin activation and focal adhesion formation

Significance

Abstract

Results

PIP2 Binding to Talin Allosterically Blocks the Integrin and Talin Tail-Binding Sites.

The Talin Head Domain Harbors Only One PIP2-Binding Site.

Talin–PIP2 Interactions Are Essential for Focal Adhesion Formation.

Discussion

Experimental Procedures

DNA Constructs and Protein Preparation.

In Vitro and in Vivo Functional Assays and Confocal Microscopy.

X-Ray Crystallography.

Acknowledgments

Footnotes

References

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Information

PIP₂ Binding to Talin Allosterically Blocks the Integrin and Talin Tail-Binding Sites.

The Talin Head Domain Harbors Only One PIP₂-Binding Site.

Talin–PIP₂ Interactions Are Essential for Focal Adhesion Formation.