The interaction of talin with the cell membrane isessential for integrin activation and focaladhesion formationKrishna Chinthalapudia, Erumbi S. Rangarajana, and Tina Izarda,1

aCell Adhesion Laboratory, Department of Integrative Structural and Computational Biology, The Scripps Research Institute, Jupiter, FL 33458

Edited by Barry Honig, Howard Hughes Medical Institute and Columbia University, New York, NY, and approved August 23, 2018 (received for review April11, 2018)

Multicellular organisms have well-defined, tightly regulated mech-anisms for cell adhesion. Heterodimeric αβ integrin receptors playcentral roles in this function and regulate processes for normal cellfunctions, including signaling, cell migration, and development,binding to the extracellular matrix, and senescence. They are in-volved in hemostasis and the immune response, participate in leu-kocyte function, and have biological implications in angiogenesisand cancer. Proper control of integrin activation for cellular commu-nication with the external environment requires several physiolog-ical processes. Perturbation of these equilibria may lead to constitutiveintegrin activation that results in bleeding disorders. Furthermore,integrins play key roles in cancer progression and metastasis in whichcertain tumor types exhibit higher levels of various integrins. Thus, theintegrin-associated signaling complex is important for cancer therapydevelopment. During inside-out signaling, the cytoskeletal protein talinplays a key role in regulating integrin affinity whereby the talin headdomain activates integrin by binding to the cytoplasmic tail ofβ-integrin and acidic membrane phospholipids. To understand themechanism of integrin activation by talin, we determined the crystalstructure of the talin head domain bound to the acidic phospholipidphosphatidylinositol 4,5-bisphosphate (PIP2), allowing us to design alipid-binding–deficient talin mutant. Our confocal microscopy with talinknockout cells suggests that the talin–cell membrane interaction seemsessential for focal adhesion formation and stabilization. Basal integrinactivation in Chinese hamster ovary cells suggests that the lipid-binding–deficient talin mutant inhibits integrin activation. Thus, mem-brane attachment of talin seems necessary for integrin activation andfocal adhesion formation.

angiogenesis | cell adhesion | integrin activation | phospholipids |talin activation

Talin is a key player in integrin activation. Vertebrates expresstwo isoforms in which talin1 is ubiquitously expressed, while

talin2 is found primarily in striated muscle and in the brain. As amultidomain cytoskeletal protein, talin contains discrete bindingsites for acidic phospholipids, β-integrin, actin, and vinculin, aswell as layilin, PIPK1γ90, and synemin. Talin links microfilamentsto the cytoplasmic membrane at cell-extracellular matrix adhesionsites. This process depends critically on talin. Talin consists of apolypeptide chain of 2,541 amino acids and is often described ashaving an N-terminal FERM (four-point-one, ezrin, radixin,moesin) domain connected by a linker (residues 401–482) thatharbors a calpain-II cleavage site to a large “rod” domain (resi-dues 483–2,541). The talin head domain is different from all otherFERM domain-containing proteins in that it has four subdomains,F0–F3 (instead of the typical three, F1–F3), and they adopt anextended structure (1) instead of the canonical cloverleaf con-formation seen in the ERM family of proteins (2). As seen inother FERM domain-containing proteins, the talin FERM sub-domains contain a ubiquitin-like F1, acyl-CoA–binding protein-like F2 and phosphotyrosine-binding–like F3 subdomain. Unlikeall other F1 FERM subdomains, the talin F1 subdomain has anunstructured insert (F1 loop, residues 133–165, harboring two

major phosphorylation sites, T144 and T150) (3), and the talinpreceding F0 subdomain has a ubiquitin-like fold. The talin F3subdomain harbors the primary β-integrin–binding site (4, 5). Thetalin rod domain consists of 13 domains, R1–R13, composed of 62amphipathic α-helices arranged into four-helix (R2, R3, R4, andR8) 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 othertalin domains, to integrin, and to vinculin (8, 9). A secondaryintegrin-binding site in the rod domain (residues 1,974–2,293) ofuncertain function has also been identified (10). There are threeactin-binding sites located on the head domain and R4–R8 andR13 subdomains (6, 11, 12), and 13 vinculin-binding sites that aresingle 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 oftalin attachment to the membrane (8). In the first stages of cellattachment, the talin F3 FERM domain binds to the NPxY motifof the integrin cytoplasmic β tail, thereby inducing reorganizationof the integrin heterodimer and activating integrin (5, 21–24).Talin attachment to the plasma membrane is enhanced by phos-phatidylinositol 4,5-bisphosphate (PIP2), which induces a confor-mational change in talin to expose the integrin-binding site (22,25–28). The role of PIP2 in integrin activation is particularly in-teresting since PIP2 is a major phosphoinositide of the inner

Significance

Vertebrate cell growth, division, locomotion, morphogenesis,and development rely on the dynamic interactions of cells withextracellular matrix components via cell surface complexestermed focal adhesions that are composed of heterodimeric αβintegrin receptors, associated signaling molecules, and the largecytoskeletal protein talin. While it is known that talin activationand binding to β-integrin requires interactions with lipids, littleis known regarding the structure and function of inactive vs.activated talin, and what is known is often disputed. Here wereport that talin binding to the cell membrane seems necessaryfor integrin activation and focal adhesion formation, a findingthat significantly advances our understanding of integrin acti-vation and might aid the development of novel integrintherapeutic agents.

Author contributions: K.C., E.S.R., and T.I. designed research; K.C. and E.S.R. performedresearch; 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.

Published under the PNAS license.

Data deposition: The atomic coordinates and structure factors have been deposited in theProtein Data Bank, www.wwpdb.org (PDB ID code 6mfs).1To whom correspondence should be addressed. Email: cmorrow@scripps.edu.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1806275115/-/DCSupplemental.

Published online September 25, 2018.

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membrane (29, 30), and because talin regulates the local PIP2concentration in the membrane by binding and activating PIPK1γ(31, 32). PIP2 regulates important processes, such as vesiculartrafficking, platelet activation, cytoskeleton organization (33–35),and focal adhesion turnover (25, 26, 36). This process evolves bytargeting proteins to the membrane, often through induction of aconformational change or oligomerization (36, 37).In mammals, the heterodimeric integrin transmembrane re-

ceptors are composed of 18 distinct α and β chains (38, 39). Byresponding to extracellular and intracellular stimuli, integrinsconnect the extracellular matrix to the cytoskeleton, and trans-duce 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 aswell as many tissues in which extracellular matrix remodeling,angiogenesis, and cell migration are involved. These processesrequire tightly controlled integrin activation mechanisms thatinvolve conformational changes of these receptors. Thus, un-derstanding the molecular mechanisms of how talin activatesintegrin is fundamental for gaining insight into important path-ological states and recognizing how integrin activation might aidthe development of novel integrin antagonists.Here we report the talin1 head/PIP2 complex crystal structure

together with biochemical and functional data that answer im-portant questions, including how PIP2 activates talin. Our dataprovide several surprises and answers to longstanding mecha-nistic questions and suggest a mechanism in which on re-cruitment 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, ourin vivo data suggest that the talin–PIP2 interaction is crucial fortalin localization to the cell membrane, affects the scaffolding ofcells, and thus is likely key for cell spreading and adhesion. Wefurther find that disrupting talin binding to the membrane affectsintegrin activation, and that this talin–PIP2 interaction seemsnecessary for focal adhesion formation. Collectively, our studyprovides a major advance in our understanding of the dynamiccontrol of focal adhesions by talin.

ResultsPIP2 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 deletionmutant of that domain Δ139–168 (talin residues 1–400), bothbound to PIP2 (Fig. 1 and SI Appendix, Tables S1 and S2).However, the full-length head domain structure did not showclear electron density for residues 139–168. The electron densitymap for the lipid was poorer compared with the PIP2-boundΔ139–168 talin structure.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 differsfrom the classical phosphoinositide-binding mode of otherknown modules (40), including the ERM protein radixin, whereIP3 bound between the interface between F1 and F3, making themolecular events for membrane-mediated spatiotemporal regu-lation of talin inhibition and activation unique. The PIP2-bindingsite is lined by residues from the F2 (K272) and F3 (K316, K324,E342, and K343) FERM subdomains, in which the 4′-phosphategroup of PIP2 interacts with talin residues K272, E342, and K343and the 5′-phosphate group interacts with E342 and K316. Inaddition, K272 interacts with the hydroxyl moiety of the inositol,while one of the carbonyls of the diacylglycerol moiety is withinhydrogen-bonding distance to K324.Superposition of our lipid-bound structure onto the talin head/

tail (F2-F3/R9) structure [Protein Data Bank (PDB) ID code4F7G] (28) reveals a large movement of approximately 10 Å ofthe F3 loop (residues 318–325) that is extensively involved in

binding to the talin rod R9 subdomain as well as to integrin, butnot 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 N323and the symmetry-related T354. In the head/tail structure, N323

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 (residues209–400; yellow) bound to the tail rod R9 subdomain (residues 1655–1824,cyan; PDB ID code 4F7G) (28). The red double arrow indicates how PIP2binding 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. Thered double arrow indicates how PIP2 binding prevents the membrane-proximal integrin binding. (C) Superposition of our talin/PIP2 structure (col-ored spectrally: F0, residues 4–83, orange; F1, 85–195, yellow; F2, 209–304,green; F3, 311–398, blue; 139-171, disordered) onto the unbound talin1structure (residues 1–400, Δ139–168, gray; PDB ID code 3IVF) (1) highlights therelative F0-F1 domain movements of approximately 7° and 3 Å.

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engages in hydrogen-bonding interactions with T1767 andD1809. Strikingly, K324 is approximately 6 Å closer to the lipid-binding site in the talin/PIP2 structure compared with its positionin the head/tail structure. This causes the 318–325 loop to move,whereby K320 releases the talin R9 rod subdomain by sterichindrance. Collectively, the structures show the molecular basisof how the binding of PIP2 and the talin R9 rod subdomain seemmutually exclusive, particularly since the lipid-binding residuesare not involved in crystal contacts (SI Appendix, Fig. S1). Thisfinding is supported by solution studies showing that PIP2 vesi-cles compete effectively with talin R9 binding to the talin F2F3subdomains (28).Similarly, superposition of our talin/PIP2 structure onto the

talin2 F2-F3/integrin β1D structure (PDB ID code 3G9W) showsthat the 318–325 loop adopts a similar conformer in the integrin-bound state and the R9-bound state (Fig. 1B). Notably, the talinL325R mutation abolishes its binding to the integrin membrane-proximal region (4). The position of the loop in our lipid-boundstructure causes K321 to occupy the integrin membrane-proximal binding site. In addition, the integrin and the talin R9rod domain-binding sites overlap on the talin head domain.These findings are consistent with studies showing that theintegrin membrane-distal site is necessary for talin-inducedintegrin activation (41). In our lipid-bound structure, N323 en-gages only in water-mediated crystal contacts with a symmetry-related T354. Thus, it seems unlikely that the distinct loopconformation is caused by crystal contacts, and more likely that itis caused by lipid binding (SI Appendix, Fig. S2).The unbound structure (PDB ID code 3IVF) (1) is iso-

morphous to and almost identical to our PIP2-bound structureexcept for relative domain movements. Significantly, superposi-tion of the respective F2F3 domains shows relative F0F1 domainmovements of approximately 7° and 3 Å (Fig. 1C). Notably, the318–325 loop is in the PIP2-bound conformer in the apo struc-ture. PIP2 also engages in crystal contacts with K334 and E335,and the E335 side chain moves to make room for the lipid (SIAppendix, Fig. S3). Collectively, this suggests that integrin or R9binding causes the distinct 318–325 loop conformation.The interpretation of comparisons with the PIPK1γ/talin struc-

tures (PDB ID code 2G35) (42) is less obvious, perhaps becausethese structures are from either the talin2 isoform or a talin1-PIPK1γ chimera. Furthermore, PIPK1γ is not in contact with theloop region that binds the integrin membrane-proximal site, butinstead overlaps with the integrin membrane-distal site on talin.

The Talin Head Domain Harbors Only One PIP2-Binding Site. The F1loop (residues 133–170) has been shown to be required forintegrin activation but not for integrin binding. It can form anα-helix and as an isolated peptide, interacts with lipids. To de-termine if in the context of the talin head domain there is an-other 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 con-firmed the integrity of our mutant proteins by thermal denaturation(SI Appendix, Fig. S4). This mutant talin exhibited similar meltingtemperatures (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 thestructure of the proteins.We determined lipid binding via a lipid cosedimentation assay

(Fig. 2A), which we used previously to detect micromolar lipidbinding (43). The mutant showed approximately 12-fold lessbinding (as assessed using ImageJ) to the lipid vesicles comparedwith the wild-type talin.We found a strong electron-dense feature near the side chain

of talin residue R358 that is part of the membrane-distal bindingsite and has been identified as involved in the interaction withintegrin by NMR studies (4). This feature is also near N285 andQ288 from a symmetry-related molecule (SI Appendix, Fig. S5).

We initially interpreted this feature as a second PIP2-bindingsite, since two PIP2-binding sites (with affinities of 0.4 μM and5 μM for PIP2diC8) were observed by isothermal titration calo-rimetry (28). However, another study using a phospholipidbilayer that contained 10% PIP2 and was immobilized on a

Fig. 2. The talin head domain harbors one PIP2-binding site. (A) Structure-based mutagenesis confirms our talin1 lipid-binding sites by lipid cosedi-mentation assay. The 3 M mutant (Y377F, R358Q, K357Q) binds lipids as seenfor wild type (WT) talin, while the minimal lipid-binding–deficient mutant5M (K272Q, K316Q, K324Q, E342Q, K343Q) and the lipid-binding-deficientmutant 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 ofincreasing concentrations of PIP2 micelles on excitation at 414 nm. The or-dinate shows the relative fluorescence, and the horizontal axis shows thewavelength (nm) of the fluorescence emission scan plot. 0 and 200 μM PIP2are shown in black and red traces, respectively; traces for the other con-centrations are colored spectrally. (Top) Wild type was predominantly mo-nomeric up to approximately 20 μM PIP2 and dimeric for all other fourhigher 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.

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Biacore L1 chip found that the second, weaker site resided onF0F1 but did not affect the stoichiometry of the interaction of talinwith acidic bilayers, and that the contribution of the second sitewas therefore negligible (26). Furthermore, our talin K358Qmutation resulted in similar binding to lipid vesicles as seen forwild-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 occurswith the guanidinium group of R358. The phosphates are alsowithin hydrogen-bonding distance to the carbonyl of N285 andthe amide of Q288 of a symmetry-related molecule (SI Appendix,Fig. S5). With respect to the two reported binding constants, it isinteresting to note that the crystallographic twofold generates atalin dimer. A dimer is also detected by fluorescence resonanceenergy transfer of CFP (donor) and YFP (acceptor) fused wild-type talin proteins, but the same assay does not detect di-merization 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 contrib-utes 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 lackendogenous talin and do not adhere to the extracellular matrix(12). Exogenous expression of GFP-tagged full-length wild-typetalin or the “3M” mutant that resides near the membrane-distalintegrin-binding site (K357Q, R358Q, Y377F) rescued focal ad-hesion formation, cell spreading, and cells clearly displaying focaladhesions 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. Thestructure of both mutants seemed to be preserved, as judged bythermal denaturation (SI Appendix, Fig. S4). Furthermore, cellsexpressing 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–deficientmutant talin (5M or 8M) accumulated in the cytosol compared withwild-type or mutant (3M) expressing cells. When we previouslymutated 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 expressingthe lipid-binding–deficient talin mutant. Thus, lipid binding seemsnecessary for talin localization to the focal adhesion membrane sitesand for talin regulation of the scaffolding effects. Furthermore,talin-null cells expressing lipid-binding–deficient talin mutantswere approximately fivefold smaller (with a chaotic and diffusedactin network) compared with talin-null cells expressing wild-type talin, in which a pronounced F-actin is visible, as thecellular integrity is maintained by intact talin–membrane inter-actions and proper cytoskeletal rearrangements.Next, to determine the effects of talin binding to the plasma

membrane, we assessed the PIP2-mediated integrin activation inChinese hamster ovary (CHO) cells that stably expressed integrin(αIIbβ3). When we transiently transfected talin1, we obtained toofew cells expressing the protein to perform our integrin activationassays in triplicate. We overcame this by generating stable CHOcells expressing full-length talin1 tagged with EGFP. For eachconstruct, we measured the mean fluorescence intensity (MFI) ofthe bound ligand in at least two independent experiments. ThePAC1 antibody, which recognizes only activated αIIbβ3 receptors,bound to integrin αIIbβ3 with higher affinity in cells expressing wild-type talin1, with anMFI of 13.5% compared with the cell-expressingmutant talin1 (MFI of 1.57%) or the lipid-binding–deficient mu-tants (MFI of 0.98% for 5M and ≤0.26% for 8M) (Fig. 3 B and Cand SI Appendix, Fig. S6). Collectively, our data show that dis-rupting 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 oftalin by FRAP in the talin-null cell background (Fig. 4A). Theseexperiments were not possible with the lipid-binding–deficienttalin mutants that lack distinct focal adhesions in which the

Fig. 3. Talin–PIP2 interactions seem to be essentialfor focal adhesion formation. (A) Talin-null papillarycollecting duct cells (PCDs) engineered to expressfull-length wild-type GFP-talin1 or mutant GFP-talin1fusion proteins were analyzed by confocal laser scan-ning microscopy. Representative images, which definethe localization of GFP-talin1 (green) at FAs decoratingF-actin (red), along with the merged channels, areshown along with the nuclei stained with DAPI. Datashown are representative of five independent experi-ments. (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–deficientmutant 5M (K272Q, K316Q, K324Q, E342Q, K343Q)represented as FACS dot plots. Abscissas, PAC1 binding;ordinates, full-length talin1 expression. Each experi-ment was repeated twice. Stably transfected wild-typetalin1 increases integrin activation (correlated to PAC1binding), but the lipid-binding–deficient mutant sig-nificantly reduces integrin activation.

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major pool of GFP-talin proteins are present in the cytosol. Thefitting of our full-length talin FRAP data with >99% confidencewas possible only with the double exponential (Fig. 4B). Theresulting recovery curve revealed that the fluorescence recoverywas biphasic with an initial fast-phase t1/2 of 1.7 ± 0.15 s and a slow-phase t1/2 of 9.2 ± 0.36 s. Thus, talin is recruited to focal adhesions,and this localization is maintained in cells moving in a persistentmanner, allowing determination of the dynamics for the associationof talin with the membrane. The initial fast rate of recovery pos-sibly accounts for the bulk of recovery and most likely reflects thisrebinding of cytosolic talin to focal adhesions at the plasmamembrane. A relatively small amount of recovery also may haveoccurred by lateral diffusion from the adjacent membrane, whichwould be consistent with the second observed, slower t1/2. Collec-tively, talin binding to the membrane is a dynamic process impor-tant for its scaffolding effects at the focal adhesion membrane sites.

DiscussionWe determined the talin head/PIP2 complex crystal structure andconfirmed the lipid-binding site seen in the crystal biochemically.Based on sequence similarity and mutagenesis, one of the bona-fide residues involved in PIP2 binding, K272, has previously beenidentified as the “membrane orientation patch.” Other postulatedlipid-binding residues (K256, K274, and R277) (4) are not part ofthe lipid-binding site. Our structural and biochemical identifica-tion of K324 as another bonafide PIP2-binding residue also agreeswith NMR studies revealing a perturbation in the 268–278 and318–324 regions on binding to IP3 (28). Surface plasmon reso-nance data showed that the K324A mutant bound PIP2 sixfoldweaker compared with wild type. Molecular dynamics simulationspredicted that K324 might be in hydrogen-bonding interactionswith PIP2 in addition to being in contact with residue R995 fromthe integrin α subunit and thus releasing the electrostatic in-teraction with D723 from the integrin β subunit. However, in theabsence of our high-resolution talin/PIP2 structure, functionalstudies had been difficult to interpret. Here we mutated the fivelipid-binding residues that we confirmed structurally and bio-chemically to be involved in PIP2 binding: K272Q, K316Q,K324Q, E342Q, and K343Q. We found that the talin–lipid in-teraction seems to be essential for focal adhesion formation andstabilization, 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 integrinmembrane-distal binding site does not affect focal adhesionformation (Fig. 3A). Comparison of our lipid-bound structurewith the integrin-bound talin structure revealed a movement of

the side chain of R358 to stack with integrin W775, while K357 isin electrostatic interaction with integrin E779 at the membrane-distal region. In contrast, the lipid-binding–deficient mutant af-fected both focal adhesion formation and integrin activation(Fig. 3). Notably, the distal integrin membrane interaction withtalin has been shown to provide the initial linkage between talinand integrin, while strong activation arises from the subsequentbinding of talin to the integrin membrane-proximal region (5).Our studies suggest that both integrin-binding sites are involvedin integrin activation, while only the membrane-proximal site isinvolved in focal adhesion formation.The majority of previous in vivo studies have used the tran-

siently transfected talin head domain (residues 1–400) or just theF2F3 (residues 203–400) or F3 (residues 309–400) talin FERMsubdomain (5, 24), which mimic the activated talin form. Incontrast, we used full-length inactive talin to measure integrinactivation by stably expressing talin proteins and selective sortingvia two rounds of fluorescent-activated cell sorting (FACS) be-fore performing the integrin activation assays. These stable poolsof talin-expressing cells enabled us to reliably and reproduciblymeasure integrin activation rates using A5 CHO cells.PIP2 activates talin by severing the head–tail interaction,

thereby exposing the integrin-binding site, although simultaneousbinding of talin to the integrin membrane-proximal site and to PIP2seem mutually exclusive. Mutating talin residues involved in bindingto the integrin membrane-distal site did not affect focal adhesionformation (Fig. 3A), while integrin activation was reduced by ap-proximately 8.5-fold compared with wild type (Fig. 3B and SI Ap-pendix, Fig. S6). In contrast, the lipid-binding–deficient mutantaffected both focal adhesion formation and integrin activation.This suggests that the integrin membrane-proximal region playsa role in integrin activation and focal adhesions, whereas themembrane-distal region impacts only integrin activation.Talin recruitment to the membrane leads to integrin activa-

tion. These sequential events are closely linked because integrinactivation requires the talin head domain to be positioned closeto the integrin tail on the cytoplasmic face of the membrane.Talin-mediated integrin activation requires that the auto-inhibitory interactions between the talin head and rod domainsare released by PIP2. The autoinhibitory talin F2F3/R9 structureidentified the head–tail interface and suggested a negativelycharged surface to repel the membrane, although the orientationof the lipid was unknown. However, our talin/PIP2 structureshows that this negatively charged surface is not planar, butrather is almost perpendicular to the PIP2-binding site (SI Ap-pendix, 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 thedistance between talin and the plasma membrane. Furthermore,two of the talin phosphorylation sites that have been mappedfrom activated human platelets (3) are located on this loop(T144 and T150). This suggests that phosphorylation of thesesites might prevent talin–membrane interactions. However, ourlipid cosedimentation results showed that the lipid-binding–deficient talin, which has an intact F1 loop, does not bind tolipids. This suggests that in the context of the entire talin headdomain (residues 1–400), this loop does not interact with themembrane. It remains to be seen if the second integrin-bindingsite located in the talin tail domain (residues 1984–2,113) allowsfor simultaneous binding of full-length talin to the membrane viaF2F3 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 siteand PIP2 seem to be mutually exclusive (Fig. 1B). In agreementwith the earlier studies, we show how PIP2 severs the talin head–tail interaction to expose the cryptic integrin-binding site. Col-lectively, these findings indicate that on talin recruitment to the

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 photo-bleaching (arrows). (B) FRAP recovery curve of talin1 in talin−/− PCD cells. Adouble-exponential model was used to fit these normalized fluorescencecurves. The red line is the calculated curve that fits the experimental dataand 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.)

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cell membrane, lipid binding to the talin head domain releasesthe interaction of the talin head with the talin rod domains.Activated talin can then activate integrin, which releases theinteraction of talin with the membrane. It remains to be seen ifthe integrin membrane-proximal binding site is solvent-exposedin the full-length talin structure.

Experimental ProceduresDNA Constructs and Protein Preparation. All bacterial expression plasmids andmammalian expression plasmids of talin1 used in this study were cloned usingmouse full-length talin1 as a template. Site-directed mutagenesis was per-formed to generate the talin1 mutants, and all DNA constructs weresequence-verified. Proteins were expressed in Rosetta 2 or BL21-CodonPlus(DE3)-RIL host cells and purified by nickel affinity and size exclusionchromatography.

In Vitro and in Vivo Functional Assays and Confocal Microscopy. The talin1 headdomain proteins were used for in vitro FRET assays as CFP and YFP FRET donorand acceptor pairs. In brief, FRET measurements were performed for 1.5 μMCFP-talin1 and 4 μM YFP-talin1 wild-type and mutant proteins in the absenceand presence of increasing concentrations of PIP2 micelles (0–200 μM).

High-resolution confocal microscopy and FRAP experiments wereperformed 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 byFACS 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 op-timizing the PIP2diC8-to-talin ratio. X-ray diffraction data were collected atthe Stanford Synchrotron Radiation Lightsource, beamline 12–2, and theX-ray diffraction data were indexed, integrated, and scaled using XDS andAIMLESS 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 StanfordSynchrotron Radiation Lightsource for synchrotron support. We thank theMax Planck Florida Light Microscopy facility and Florida Atlantic University,Nikon Center of Excellence for imaging facilities, Dr. Roy Zent (VanderbiltCenter for Kidney Disease) for talin-null cells, and Dr. Mark Ginsberg (Uni-versity of California, San Diego) for A5 CHO cells. We thank Marina Primi[The Scripps Research Institute (TSRI)] and Charmane Gabriel (Oxbridge Acad-emy) for protein expression and Louis Shane (Palm Beach Gardens, FL) andDouglas Bingham (TSRI) for helpful discussions regarding the manuscript. T.I. issupported by grants from the National Institute of Health and the Departmentof Defense, and by startup funds provided to TSRI from the State of Florida.This is publication 29675 from The Scripps Research Institute.

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