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. 2012 Jul 23;109(32):12992–12997. doi: 10.1073/pnas.1208296109

Talin couples the actomyosin cortex to the plasma membrane during rear retraction and cytokinesis

Masatsune Tsujioka a,1,2,3, Shigehiko Yumura b, Kei Inouye c, Hitesh Patel d, Masahiro Ueda e,1,2, Shigenobu Yonemura a
PMCID: PMC3420178  PMID: 22826231

Abstract

Contraction of the cortical actin cytoskeleton underlies both rear retraction in directed cell migration and cytokinesis. Here, we show that talin, a central component of focal adhesions, has a major role in these processes. We found that Dictyostelium talin A colocalized with myosin II in the rear of migrating cells and the cleavage furrow. During directed cell migration, talin A-null cells displayed a long thin tail devoid of actin filaments, whereas additional depletion of SibA, a transmembrane adhesion molecule that binds to talin A, reverted this phenotype, suggesting a requirement of the link between actomyosin and SibA by talin A for rear retraction. Disruptions of talin A also resulted in detachment of the actomyosin contractile ring from the cell membrane and concomitant regression of the cleavage furrow under certain conditions. The C-terminal actin-binding domain (ABD) of talin A exhibited a localization pattern identical to that of full-length talin A. The N-terminal FERM domain was found to bind phosphatidylinositol 4,5-bisphosphate [PtdIns(4,5)P2] and phosphatidylinositol 3,4,5-trisphosphate [PtdIns(3,4,5)P3] in vitro. In vivo, however, PtdIns(4,5)P2, which is known to activate talin, is believed to be enriched in the rear of migrating cells and the cleavage furrow in Dictyostelium. From these results, we propose that talin A activated by PtdIns(4,5)P2 in the cell posterior or cleavage furrow links actomyosin cytoskeleton to adhesion molecules or other membrane proteins, and that the force is transmitted through these links to retract the tail during cell migration or to cause efficient ingression of the equator during cytokinesis.

Contraction of cortical actomyosin, which is composed of filamentous actin and myosin II, mediates the changes in cell shape that occur during diverse biological processes. For example, this contraction drives the apical constriction of cells in epithelial sheets, which results in sheet folding, an important step in neural tube formation in the early stages of vertebrate development (1). In this case, contractile forces are transmitted across the sheet through adherence junctions to induce the deformation of the entire tissue. Cell shape changes driven by actomyosin contraction are also essential for important processes in isolated cells, including rear retraction in migrating cells and ingression of the cleavage furrow in dividing cells (2, 3). In these processes, however, little is known about which molecules link the actomyosin cortex to the plasma membrane to transmit the contractile forces.

One such candidate molecule is talin (4). In mammals, this large cytoskeletal protein (∼270 kDa) is a major component of integrin-mediated adhesion structures, named focal adhesions (5), where talin has an important role of linking adhesion molecules to the actin cytoskeleton. Accordingly, the depletion of talin alters cell-substrate adhesion in mouse undifferentiated ES cells and fibroblast cells (6, 7). Talin possesses two conserved domains: the FERM domain at the N terminus, which interacts with a variety of proteins including integrins and lipids within the plasma membrane, and the I/LWEQ module at the C terminus, which is an actin-binding domain (ABD).

In this study, we investigated the role of the Dictyostelium homolog of talin (talin A) in the transmission of contractile forces produced by actomyosin to the plasma membrane in rear retraction and cytokinesis (8, 9). Dictyostelium, a eukaryotic haploid organism, is an ideal system for investigating the mechanisms of myosin II-related events. Thanks to having a single copy of the myosin II gene (10), myosin II-depleted cells can be readily obtained and have been extensively studied (11). Dictyostelium cells grow as solitary amoebae in the vegetative stage. Upon starvation, however, they stop proliferation and aggregate to form multicellular structures. During aggregation, cells move toward the aggregation centers by chemotaxis to cAMP, leading to the formation of streams composed of highly polarized cells. As in mammalian cells, the contraction of cortical actomyosin provides the forces necessary for rear retraction in cell migration and ingression of the cleavage furrow in cytokinesis (2, 11, 12).

Talin A has the same structural features as mammalian talins and appears to have similar functions (8, 9). It accumulates at cell–substrate adhesion sites (13, 14), whereas its absence reduces adhesion to the substratum. Talin A also appears to function at locations other than adhesion sites. A micropipette aspiration assay revealed weakening of the otherwise strong coupling between the plasma membrane and the cytoskeletal cortex at the cell posterior in talin A-null cells (15), suggesting that talin A acts as a linker between them at the posterior of cells where myosin II accumulates. Additionally, talin A-null cells show cytokinesis defects under nonadherent conditions (8). In this study, we examined the subcellular localization of talin A under various conditions using the C-terminal GFP fusion (talin A–GFP), which has been shown to rescue the defects of talin A-null cells and accumulate at cell–substrate adhesion sites (14, 16). We next examined talin A-null cells in detail, focusing on the tail region during cell migration and the cleavage furrow during cytokinesis. The results provide evidence for the requirement of talin A for the transmission of contractile forces from actomyosin to the cell membrane for rear retraction in cell migration and equatorial furrowing in cytokinesis.

Results

Talin A Colocalizes with Myosin II at the Rear of Migrating Cells.

In steadily migrating polarized cells such as the cells in aggregation streams, talin A–GFP was primarily enriched at the rear of the cells, a distribution pattern very similar to myosin II (Fig. 1A, a and b and Movie S1). This posterior enrichment was further confirmed with a red fluorescent protein (RFP) fusion protein and also by staining of wild-type cells with an anti-talin A antibody (Fig. S1 A and B). The spatial correlation between talin A and myosin II was further investigated on a finer scale by total internal reflection fluorescence (TIRF) imaging of talin A–GFP-expressing cells double stained with antibodies against myosin II and GFP. As shown in Fig. 1B, a large fraction of talin A–GFP molecules in the cell cortex are located near myosin II, suggesting the possibility of direct binding. However, the results of coimmunoprecipitation experiments using an antimyosin II antiserum did not support this possibility. Although talin A–GFP as well as actin did coprecipitate with myosin II, neither of them did under conditions where myosin II is dissociated from actin filaments, indicating the interactions between talin A–GFP and myosin II being mediated by actin filaments (Fig. S2 A and B). In less polarized cells, talin A–GFP was often enriched in pseudopodia that were extending for a limited period before retraction (Fig. S1C and Movie S2). Photobleaching experiments indicated that talin A in migrating cells was almost stationary with respect to the substratum (Fig. S1D and Table S1), consistent with the posterior accumulation due to retrograde flow of talin A in polarized cells showing sustained unidirectional locomotion.

Fig. 1.

Fig. 1.

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Subcellular distribution of talin A and myosin II in migrating cells. (A) Confocal images of aggregating talin A-null cells expressing talin A–GFP (a), myosin II-null cells expressing GFP–myosin II (b), myosin II-null cells expressing talin A–GFP (c), talin A-null cells expressing GFP–myosin II (d). Large arrows indicate the direction of migration. In c, a dragged tail and the leading edge rich in talin A–GFP are indicated by arrowheads and a small arrow, respectively. (B) High magnification TIRF images of a small area of a talin A–GFP-expressing cell fixed and immunostained with antibodies against GFP (green) and myosin II (red). A significant colocalization of the two signals is evident in the merged image. (Scale bars, 10 μm in A and 1 μm in B.)

Talin A–GFP introduced into a talin A/myosin II double knockout mutant was present throughout the whole plasma membrane of polarized cells. Although it still strongly concentrated in the dragged tail frequently shown in myosin II-null cells (Fig. 1A, c; arrowheads), the distribution is in contrast to its posterior localization in polarized myosin II-expressing cells (compare Fig. 1 A, a with A, c and Movie S3). Notably, talin A–GFP often accumulated at the leading edge of myosin II-null cells (Fig. 1A, c; small arrow). These results indicate that myosin II is required for the proper posterior localization of talin A but not for its association with cell cortex. Conversely, GFP–myosin II introduced into double mutant cells was found at the rear of migrating cells (Fig. 1A, d), just as in talin A-expressing cells (Fig. 1A, b), indicating that the distribution of myosin II is independent of talin A.

Talin A Is Required for Normal Tail Retraction in Directed Cell Migration.

To gain insight into possible functions of talin A in the rear of migrating cells, we closely examined the rear part of mutant cells migrating toward a microcapillary filled with a cAMP solution. Talin A-null cells were often seen to trail a long thin tail, sometimes longer than the cell body (Fig. 2B, arrowheads). This was apparently caused by the cell membrane failing to detach from the substratum at the rear edge of the cells (Fig. 2E and Movie S4). Observation by interference reflection microscopy (IRM) indicated that small areas of the long tails were in close contact with the substratum (Fig. 2F, arrowheads). In some cells, the tail was detached from the substratum and retracted into the cell body, whereas in others the tail continued to be stretched (Fig. 2E). This is consistent with a model in which some adhesion molecules normally linked to the cortical cytoskeleton by talin A are not pulled off the substratum in talin A-null cells that keep advancing. SibA (17, 18) is a candidate of such adhesion molecules. The Sib proteins are proposed to be functional homologs of the β-chain of integrin heterodimer in Dictyostelium, and talin A binds to all of the five Sib proteins in vitro (18). Specifically, SibA is shown to be important in cell–substrate adhesion in the unicellular stage (17, 18). We produced a double knockout mutant lacking both talin A and SibA (talin A/SibA-null) and found that they seldom produced long tails during chemotactic locomotion (Fig. 2C). This finding suggests that SibA is the adhesion molecule that remained attached to the substratum and caused the long tails in talin A-null cells. Myosin II-null cells also dragged a long thin tail in the microcapillary assay, as observed in the aggregation streams shown in Fig. 1A, c (Fig. 2D, arrowheads). We counted the number of cells migrating toward the capillary tip showing a tail longer than half the length of the cell body in mutant and wild-type strains. Whereas 23.5% of talin A-null (8/34) and 32.1% of myosin II-null cells (9/28) exhibited such long tails, no wild-type (0/24) and only 2.6% of talin A/SibA-null cells (1/39) did.

Fig. 2.

Fig. 2.

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Altered morphologies of the tails of talin A-null and myosin II-null cells. (A–D) Phase contrast images showing wild-type (A), talin A-null (B), talin A/SibA-null (C), and myosin II-null (D) cells moving toward the tips of microcapillaries filled with a cAMP solution. (E) Phase-contrast time-lapse images of a talin A-null cell moving toward a tip of a microcapillary filled with a cAMP solution, demonstrating the elongation of a long tail. Time is indicated at the Lower Left in seconds. (F) Migrating talin A-null cell observed by interference reflection microscopy (IRM). The elongated tail closely apposed to the substrate appears dark (arrowheads). (G and H) Phase contrast and conventional fluorescent images of a migrating myosin II-null cell (G) and two examples of talin A-null cells (H) expressing GFP–ABD. F-actin is present in the tail of myosin II-null but not of the talin A-null cells. The elongated tails are indicated by arrowheads (B and D –F). (Scale bars, 10 μm in A–H.)

Some differences were noted in the elongated tails of myosin II-null and talin A-null cells. They tend to be thicker in myosin II-null than talin A-null cells. When F-actin was visualized in mutants with a fluorescent probe (GFP–ABD) (19), fluorescent signals were detected in the tails of myosin II-null cells but not talin A-null cells (Fig. 2 G and H), indicating that the trailing tails in myosin II-null cells are due to the lack of cortical contraction, whereas those in talin A-null cells are a consequence of detachment of the cell membrane from the contracting actomyosin cortex.

Talin A Colocalizes with Myosin II at the Cleavage Furrow in Dividing Cells.

Because there is a similarity in the role of actomyosin contraction between tail retraction and furrow ingression, we investigated the distribution and possible functions of talin A in dividing cells. During cytokinesis, both myosin II and talin A–GFP accumulated around the equatorial region in a highly coordinated manner right before furrowing began and continued to be located in the cleavage furrow throughout cell division (Fig. 3A). In contrast, in the absence of myosin II, talin A–GFP was distributed in the periphery of the daughter cells (compare Fig. 3A to Fig. S1E), indicating that the enrichment of talin A in the equator of dividing cells is dependent on myosin II. On the other hand, the accumulation of myosin II in the cleavage furrow did not require talin A (Fig. S1F).

Fig. 3.

Fig. 3.

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Function of talin A in cytokinesis. (A) Phase contrast and TIRF images of talin A–GFP-expressing cells undergoing cytokinesis show the subcellular distribution of talin A and myosin II, which was determined by immunostaining using antibodies against GFP and myosin II. (B) Time-lapse images of GFP–myosin II-expressing wild-type and talin A-null cells attempting cytokinesis under a sheet of 3% (wt/vol) agarose show cytokinesis defects of talin A-null cells. Superimposed fluorescence and phase-contrast images are shown. Time in minutes is indicated at the Lower Left of each panel. Of the 10 wild-type cells examined, all completed furrow ingression, although it took much longer than in uncovered or nonadherent conditions where it usually takes less than 5 min (Upper) (21). Furrowing failed in the early stage in all 10 talin A-null cells examined (Lower). (Scale bars, 10 μm.)

Talin A Is Required for Normal Linkage Between the Plasma Membrane and the Contractile Ring in Cytokinesis.

The localization of talin A in the cleavage furrow suggests its role in furrow ingression. To test this possibility, we performed morphological analyses by dividing wild-type and talin A-null cells. Under adhesive conditions, talin A-null cells underwent cytokinesis following a time course similar to wild-type cells. However, the cleavage furrow was consistently broader, and the pole-to-pole distance was significantly larger, in talin A-null cells (Fig. S3 A and B). By analogy to the cytokinesis of myosin II-null cells, in which the division process is driven by opposite movement of the daughter cells (2), the above results suggest that the active polar elongation by the pseudopodia at both polar regions substantially contributes to the separation of the daughter cells in talin A-null cells. When the polar elongation was inhibited by pressing the cells under a sheet of 3% (wt/vol) agarose (20), wild-type cells still completed furrow ingression (Fig. 3B, Upper), but talin A-null cells failed to constrict the furrow. As shown in the Lower panels in Fig. 3B, GFP–myosin II accumulated in the equatorial region, where a cleavage furrow formed. However, the cell membrane on either side of the furrow soon regressed, whereas GFP–myosin II continued to contract, resulting in the separation of the contractile ring from the plasma membrane and eventual failure of cytokinesis. From these observations, we infer that the link between the contractile ring and the plasma membrane is impaired in talin A-null cells.

Possible Molecular Mechanisms of Talin A Function.

To gain insight into the molecular mechanisms of talin A in the membrane–cortex interface at the cell posterior and cleavage furrow, we investigated the interaction of N- and C-terminal domains with membrane lipids and cortical actin filaments.

To examine which lipid components of the plasma membrane bind to the FERM domain of talin A, a membrane strip spotted with 15 kinds of lipids known to be constituents of the plasma membrane was incubated with a lysate of bacteria expressing the GST-tagged FERM domain of talin A. The signals of the GST-tagged protein bound to the spotted lipids were detected with an anti-GST antibody. The talin A FERM domain was found to bind phosphatidylinositol 4,5-biphosphate [PtdIns(4,5)P2] and phosphatidylinositol 3,4,5-triphosphate [PtdIns(3,4,5)P3], with approximately four times stronger affinity to PtdIns(3,4,5)P3 than to PtdIns(4,5)P2, whereas no binding was detected with any of the other lipids examined (Fig. 4A and Fig. S4A), suggesting the importance of interaction with phosphoinositides in talin A function. In accord with this interaction, it has been shown that PtdIns(3,4,5)P3 is enriched in the pseudopodia, whereas PTEN, a phosphatase that converts PtdIns(3,4,5)P3 to PtdIns(4,5)P2, is distributed in the posterior and furrow region in migrating and dividing cells, respectively (22–24). It should be noted, however, that the distribution of talin A within the cells is independent of the phosphoinositides; talin A localization to pseudopodia in nonpolarized cells was unaffected in PI3K1–5-null cells, which are unable to phosphorylate PtdIns(4,5)P2 to produce PtdIns(3,4,5)P3, due to the lack of all five phosphatidylinositide-3 kinases (PI3Ks), or in cells treated with the PI3K inhibitor LY294002 (Fig. S4 B and C). By contrast, disruption of the actin structures using latrunculin A completely abolished talin A localization. Wild-type cells treated with latrunculin A become round, but still respond to external cAMP signal, showing PtdIns(3,4,5)P3 and PTEN accumulation toward the up and down gradient, respectively (Fig. 4B, Upper) (25). In such cells, talin A–GFP diffusely distributed in the cytoplasm in the gradient of cAMP (Fig. 4B, Lower).

Fig. 4.

Fig. 4.

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Properties of membrane- and actin-binding domains of talin A. (A) The binding activities of the FERM domain to the plasma membrane components. GST-tagged FERM domain of talin A bound to the indicated lipids spotted on the membrane was immunologically detected. TG, triglyceride; DAG, diacylglycerol; PA, phosphatidic acid; PS, phosphatidylserine; PE, phosphatidylethanolamine; PC, phosphatidylcholine; PG, phosphatidylglycerol; CL, cardiolipin; PI, phosphatidylinositol; PIP, PtdIns(4)P; PIP2, PtdIns(4,5)P2; PIP3, PtdIns(3,4,5)P3; CHO, cholesterol; SPH, sphingomyelin; and SUL, 3-sulfogalactosylceramide. The FERM domain bound to PtdIns(4,5)P2 and PtdIns(3,4,5)P3. (B) Localization of the GFP–PH domain of CRAC (26), a marker for PtdIns(3,4,5)P3, and talin A–GFP in cells in a cAMP gradient after treatment with 3 μM latrunculin A. The GFP–PH domain accumulated at the membrane domain facing the source of cAMP, whereas talin A–GFP was diffusely distributed in cytoplasm. (C) Confocal images of wild-type cells expressing GFP–I/LWEQ module. Fluorescence signals are located in the posterior of strongly polarized cells (a), in the furrow region of a cytokinetic cell (b), and in pseudopodia of nonpolarized cells (c). The arrow in a indicates the direction of migration. (Scale bars, 10 μm.)

A GFP fusion of the C-terminal actin-binding domain of talin A (I/LWEQ module, amino acids 2255–2492) showed subcellular distributions indistinguishable from those of the full-length talin A–GFP; localization in the posterior of migrating polarized cells (Fig. 4C, a), the furrow region during cytokinesis (Fig. 4C, b), and in the pseudopodia of nonpolarized cells (Fig. 4C, c). It did not cause artifactual accumulation (sequestration) of itself and actin filaments as reported by Weber et al. (27) with a longer C-terminal domain of talin A (amino acids 1908–2492). The I/LWEQ module alone could not rescue the talin A mutant phonotype.

Discussion

Subcellular Localization of Talin A.

In this study, we found talin A to be localized in the posterior of highly polarized migrating cells and in the equatorial region of dividing cells. The posterior enrichment is consistent with the report of Merkel et al. (15), who suggested that talin A serves as a linker between the cortical cytoskeleton and the plasma membrane at the cell posterior. We also observed the accumulation of talin A–GFP in extending pseudopodia of less polarized starved cells and vegetative cells, in agreement with Kreitmeier et al. (9). In either case, talin A was closely associated with the cell membrane. Domain analysis indicated that the I/LWEQ module is required for talin A localization, whereas the rest of the protein including the FERM domain is also essential for its function.

Three possibilities may be considered for the mechanism of the posterior/equator localization of talin A: accumulation by cortical actin flow (28), specific binding to a subpopulation of actin filaments, and recruitment by direct binding to myosin II. The first possibility is consistent with higher occurrence of the posterior accumulation of talin A in steadily migrating cells but not in less motile cells or myosin II-null cells and is supported by the results of photobleaching experiments demonstrating retrograde flow of talin A. Backward movement of talin A in migrating cells has also been reported by Hibi et al. (13). The second possibility gains support from the requirement of myosin II for the specific localization of talin A and their colocalization in the posterior of migrating cells and in the cleavage furrow. Thus, talin A appears to bind to cortical actin filaments that are interacting with myosin II. Uyeda et al. (29) proposed that the head domain of myosin II preferentially binds to actin filaments stretched by interaction with myosin II and that this binding property of the head domain is responsible for the accumulation of myosin II itself in the posterior cortex of migrating cells and cleavage furrow of dividing cells. Washington and Knecht (30) reported that the actin-binding domains of filamin and α-actinin determine the distribution of the proteins, which show distinct, almost complementary localization patterns, suggesting that different actin-binding domains have different affinities for actin filaments in functionally distinct regions of the cytoskeleton. In the case of talin A, its actin-binding domain (the I/LWEQ module) could conceivably have a binding property similar to the myosin head to give rise to the distribution patterns very similar to myosin II. The third possibility is implicated by the prominent colocalization between myosin II and talin A, as has been shown with animal talin (31). So far, however, we have obtained no evidence supporting their direct binding in biochemical assays (Fig. S2). Any of these three mechanisms may be working together as in the case of myosin II localization in which multiple mechanisms including retrograde flow and direct recruitment are involved (32).

Roles of Talin A in Rear Retraction and Cytokinesis.

From its localization and mutant phenotype, as well as the essential role of the C-terminal domain in its localization and the requirement of the N-terminal domain in its function, we propose a model in which talin A links the cortical actomyosin to the cell membrane and transmits the necessary forces to retract the tail during cell migration or to constrict the cleavage furrow in cytokinesis (Fig. 5 A and B). Because PTEN colocalizes with myosin II (33), the chance of talin A to encounter PtdIns(4,5)P2 would be high. Then, PtdIns(4,5)P2, which is known to be an activator of talin (34–36), activates talin A by binding to the N-terminal FERM domain, enabling talin A to associate with the binding partners within the plasma membrane. In cells migrating on a substratum, the contraction of actomyosin cytoskeleton provides the force to detach adhesion molecules such as SibA from the substratum through talin A, pulling the rear membrane back to the cell body (Fig. 5B). In talin A-null cells, however, contracting actomyosin is not connected to adhesion molecules well enough for their detachment from the substratum, resulting in long thin tails devoid of F-actin (Fig. 5B). On the other hand, in myosin II-null cells, the lack of a contraction force results in the formation of a long tail in which loose actin filaments remain connected with the cell membrane and adhesion molecules (Fig. 5B). Laevsky and Knecht (20) have also reported that myosin II-null cells drag a long tail containing GFP–ABD, which is especially prominent under an agarose sheet but also present in liquid media. In certain mammalian cell lines, depletion of myosin II results in the formation of similar F-actin–containing long tails (37).

Fig. 5.

Fig. 5.

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Models for the function of talin A. Illustrations depicting a functional model of talin A in cytokinesis (A) and in tail retraction (B). Talin A is activated by PtdIns(4,5)P2 in the posterior region of migrating cells and the furrow region in cytokinesis, where PtdIns(4,5)P2 is believed to be enriched. Then, it connects the binding partners within the plasma membrane to actomyosin. (A) Contraction of a contractile ring linked to the plasma membrane by talin A provides the main force for ingression of the cleavage furrow, leading to the formation of the deep furrow in wild-type cells. In talin A-null cells, on the other hand, the contraction force is not sufficiently transmitted to the furrowing membrane. On the substrate, therefore, outward movement of the daughter cells contributes more to the cytokinetic furrow in talin A-null than in wild-type cells, resulting in the shallow furrow as shown in Fig. S3A. In nonadherent talin A-null cells, the lack of traction forces causes more severe cytokinetic defects (8). (B) In wild-type (Top), talin A transmits a contraction force by the actomyosin cortex (thick solid lines) to tear SibA from the substratum and pull the plasma membrane inward. In the absence of talin A (Middle), the contraction force of the actomyosin cortex is inadequately transmitted to the adhesion molecules or cell membrane, resulting in the formation of a long thin tail. Without myosin II (Bottom), no pulling force is generated in the cortical actin cytoskeleton (thick dashed lines), which also results in the formation of a long thin tail containing actin filaments.

Considering the linking role of talin A at locations other than adhesion sites, talin A may bind to membrane proteins other than adhesion molecules. Murine megakaryocytes lacking talin 1 exhibit active membrane blebbing (38), suggestive of the dissociation between the cortical cytoskeleton and the plasma membrane, implying that mammalian talin also plays a role in regions other than focal adhesions.

As with animal talin (4), talin A may function as a core component for protein complexes that enhance association between the cortical cytoskeleton and the plasma membrane. Consistent with this notion, talin A is known to interact with myosin VII, which is a submembranous protein (39, 40). In preliminary experiments, we have also identified several proteins that bind to talin A, including submembranous and cytoskeletal proteins. Further studies are needed to elucidate the structural details of the linkage between the cell membrane and actomyosin cytoskeleton via talin A.

Materials and Methods

Strains, Constructs, and Microscopy.

Detailed information about constructs, Dictyostelim strains, and microscopy used in this study can be found in SI Materials and Methods.

Observation of Cell Motility and Cytokinesis.

To observe directed migration of single cells, cells growing in culture dishes were collected and resuspended in KK2 phosphate buffer at a density of 5 × 104 cells/μL. The cell suspension was spotted on nonnutrient agar as 10-μL aliquots. After 6 h, the cells, which had initiated aggregation by chemotaxis to cAMP, were collected in KK2 phosphate buffer and plated in glass-bottom dishes (35 mm; Iwaki). A microcapillary (sterile Femtotips; Eppendorf) filled with 1 mM cAMP solution was placed onto the glass surface. The phase contrast or fluorescent images of the cells moving toward the tip of the capillary were taken with an inverted microscope.

For time-lapse imaging of aggregating cells expressing GFP fusion proteins in the streams, cells were allowed to develop on nonnutrient agar up to the aggregation stage. Small pieces of agar carrying streaming cells were cut out and inverted onto glass-bottom dishes (35 mm; Iwaki). Fluorescent images of aggregation streams were captured at 5-s intervals using confocal microscopy.

For the observations of cytokinesis under an agarose sheet, the vegetative cells cultured in axenic medium in a glass-bottom dish were overlaid with an agarose sheet, which was cut out from an agarose plate made by pouring 2 mL of melted 3% (wt/vol) agarose into a 3-cm culture dish (Iwaki). To tightly cover the cells with the sheet, the setup was dried with the lid opened for 30 min after the removal of the media. Cells undergoing cytokinesis were identified and the phase contrast and fluorescent images were manually taken at 1- or 2-min intervals.

Membrane–Lipid Binding Assay.

Escherichia coli strain BL21 (pLys) was transformed with the GST–FERM construct. Expression of the GST-tagged protein was induced by adding isopropyl-Inline graphic-d-1-thiogalactopyranoside at a final concentration of 1 mM. Induction was confirmed by Coomassie Brilliant Blue staining of bacterial lysates separated by SDS/PAGE. Bacterial cells expressing the GST-tagged protein were collected in TBS-T (10 mM Tris, 100 mM NaCl, 0.05% Tween 20, pH 7.4) containing 0.5% gelatin, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, and protease inhibitor mixture (Complete Mini, EDTA-free; Roche), and lysed by sonication. After the cell lysate was cleared by centrifugation (MX-300; Tomy) at 9,100 × g for 10 min, hydrophobic membranes that had been spotted with 15 kinds of lipids at 100 pmol/spot (P-6002; Echelon Biosciences) were preincubated in TBS-T containing 0.5% gelatin for 30 min and incubated with the cell lysates for 12 h at 4 °C. After washing in TBS-T three times, the strip was incubated for 1 h in TBS-T with an anti-GST monoclonal antibody (B-14; Santa Cruz). The strip was washed as before, then incubated for 1 h in TBS-T with a horseradish peroxidase-conjugated antimouse secondary antibody (Jackson ImmunoResearch). After washing the strip in TBS-T five times, GST proteins bound to lipids spotted on the strip were detected by chemiluminescence assay (Immobilon Western; Millipore).

Supplementary Material

Supporting Information
supp_109_32_12992__index.html (1.4KB, html)

Acknowledgments

This work was supported by the Special Postdoctoral Researchers program, RIKEN, and a grant-in-aid for scientific research by the Ministry of Education, Culture, Sports, Science and Technology.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1208296109/-/DCSupplemental.

References

  • 1.Martin AC. Pulsation and stabilization: Contractile forces that underlie morphogenesis. Dev Biol. 2010;341:114–125. doi: 10.1016/j.ydbio.2009.10.031. [DOI] [PubMed] [Google Scholar]
  • 2.Uyeda TQ, Nagasaki A, Yumura S. Multiple parallelisms in animal cytokinesis. Int Rev Cytol. 2004;240:377–432. doi: 10.1016/S0074-7696(04)40004-7. [DOI] [PubMed] [Google Scholar]
  • 3.Pollard TD, Cooper JA. Actin, a central player in cell shape and movement. Science. 2009;326:1208–1212. doi: 10.1126/science.1175862. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Critchley DR, Gingras AR. Talin at a glance. J Cell Sci. 2008;121:1345–1347. doi: 10.1242/jcs.018085. [DOI] [PubMed] [Google Scholar]
  • 5.Wehrle-Haller B. Structure and function of focal adhesions. Curr Opin Cell Biol. 2012;24:116–124. doi: 10.1016/j.ceb.2011.11.001. [DOI] [PubMed] [Google Scholar]
  • 6.Priddle H, et al. Disruption of the talin gene compromises focal adhesion assembly in undifferentiated but not differentiated embryonic stem cells. J Cell Biol. 1998;142:1121–1133. doi: 10.1083/jcb.142.4.1121. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Zhang X, et al. Talin depletion reveals independence of initial cell spreading from integrin activation and traction. Nat Cell Biol. 2008;10:1062–1068. doi: 10.1038/ncb1765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Niewöhner J, Weber I, Maniak M, Müller-Taubenberger A, Gerisch G. Talin-null cells of Dictyostelium are strongly defective in adhesion to particle and substrate surfaces and slightly impaired in cytokinesis. J Cell Biol. 1997;138:349–361. doi: 10.1083/jcb.138.2.349. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Kreitmeier M, Gerisch G, Heizer C, Müller-Taubenberger A. A talin homologue of Dictyostelium rapidly assembles at the leading edge of cells in response to chemoattractant. J Cell Biol. 1995;129:179–188. doi: 10.1083/jcb.129.1.179. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Eichinger L, et al. The genome of the social amoeba Dictyostelium discoideum. Nature. 2005;435:43–57. doi: 10.1038/nature03481. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Yumura S, Uyeda TQ. Myosins and cell dynamics in cellular slime molds. Int Rev Cytol. 2003;224:173–225. doi: 10.1016/s0074-7696(05)24005-6. [DOI] [PubMed] [Google Scholar]
  • 12.Iwadate Y, Yumura S. Actin-based propulsive forces and myosin-II-based contractile forces in migrating Dictyostelium cells. J Cell Sci. 2008;121:1314–1324. doi: 10.1242/jcs.021576. [DOI] [PubMed] [Google Scholar]
  • 13.Hibi M, Nagasaki A, Takahashi M, Yamagishi A, Uyeda TQ. Dictyostelium discoideum talin A is crucial for myosin II-independent and adhesion-dependent cytokinesis. J Muscle Res Cell Motil. 2004;25:127–140. doi: 10.1023/b:jure.0000035842.71415.f3. [DOI] [PubMed] [Google Scholar]
  • 14.Patel H, et al. The multi-FERM-domain-containing protein FrmA is required for turnover of paxillin-adhesion sites during cell migration of Dictyostelium. J Cell Sci. 2008;121:1159–1164. doi: 10.1242/jcs.021725. [DOI] [PubMed] [Google Scholar]
  • 15.Merkel R, et al. A micromechanic study of cell polarity and plasma membrane cell body coupling in Dictyostelium. Biophys J. 2000;79:707–719. doi: 10.1016/S0006-3495(00)76329-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Tsujioka M, et al. Overlapping functions of the two talin homologues in Dictyostelium. Eukaryot Cell. 2008;7:906–916. doi: 10.1128/EC.00464-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Cornillon S, Froquet R, Cosson P. Involvement of Sib proteins in the regulation of cellular adhesion in Dictyostelium discoideum. Eukaryot Cell. 2008;7:1600–1605. doi: 10.1128/EC.00155-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Cornillon S, et al. An adhesion molecule in free-living Dictyostelium amoebae with integrin beta features. EMBO Rep. 2006;7:617–621. doi: 10.1038/sj.embor.7400701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Pang KM, Lee E, Knecht DA. Use of a fusion protein between GFP and an actin-binding domain to visualize transient filamentous-actin structures. Curr Biol. 1998;8:405–408. doi: 10.1016/s0960-9822(98)70159-9. [DOI] [PubMed] [Google Scholar]
  • 20.Laevsky G, Knecht DA. Cross-linking of actin filaments by myosin II is a major contributor to cortical integrity and cell motility in restrictive environments. J Cell Sci. 2003;116:3761–3770. doi: 10.1242/jcs.00684. [DOI] [PubMed] [Google Scholar]
  • 21.Nagasaki A, Kanada M, Uyeda TQ. Cell adhesion molecules regulate contractile ring-independent cytokinesis in Dictyostelium discoideum. Cell Res. 2009;19:236–246. doi: 10.1038/cr.2008.318. [DOI] [PubMed] [Google Scholar]
  • 22.Funamoto S, Meili R, Lee S, Parry L, Firtel RA. Spatial and temporal regulation of 3-phosphoinositides by PI 3-kinase and PTEN mediates chemotaxis. Cell. 2002;109:611–623. doi: 10.1016/s0092-8674(02)00755-9. [DOI] [PubMed] [Google Scholar]
  • 23.Janetopoulos C, Borleis J, Vazquez F, Iijima M, Devreotes P. Temporal and spatial regulation of phosphoinositide signaling mediates cytokinesis. Dev Cell. 2005;8:467–477. doi: 10.1016/j.devcel.2005.02.010. [DOI] [PubMed] [Google Scholar]
  • 24.Iijima M, Devreotes P. Tumor suppressor PTEN mediates sensing of chemoattractant gradients. Cell. 2002;109:599–610. doi: 10.1016/s0092-8674(02)00745-6. [DOI] [PubMed] [Google Scholar]
  • 25.Hoeller O, Kay RR. Chemotaxis in the absence of PIP3 gradients. Curr Biol. 2007;17:813–817. doi: 10.1016/j.cub.2007.04.004. [DOI] [PubMed] [Google Scholar]
  • 26.Loovers HM, et al. Distinct roles of PI(3,4,5)P3 during chemoattractant signaling in Dictyostelium: A quantitative in vivo analysis by inhibition of PI3-kinase. Mol Biol Cell. 2006;17:1503–1513. doi: 10.1091/mbc.E05-09-0825. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Weber I, Niewöhner J, Du A, Röhrig U, Gerisch G. A talin fragment as an actin trap visualizing actin flow in chemotaxis, endocytosis, and cytokinesis. Cell Motil Cytoskeleton. 2002;53:136–149. doi: 10.1002/cm.10065. [DOI] [PubMed] [Google Scholar]
  • 28.Fukui Y, Kitanishi-Yumura T, Yumura S. Myosin II-independent F-actin flow contributes to cell locomotion in dictyostelium. J Cell Sci. 1999;112:877–886. doi: 10.1242/jcs.112.6.877. [DOI] [PubMed] [Google Scholar]
  • 29.Uyeda TQ, Iwadate Y, Umeki N, Nagasaki A, Yumura S. Stretching actin filaments within cells enhances their affinity for the myosin II motor domain. PLoS ONE. 2011;6:e26200. doi: 10.1371/journal.pone.0026200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Washington RW, Knecht DA. Actin binding domains direct actin-binding proteins to different cytoskeletal locations. BMC Cell Biol. 2008;9:10. doi: 10.1186/1471-2121-9-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Lin Y, Kishi H, Nakamura A, Takagi T, Kohama K. N-terminal myosin-binding fragment of talin. Biochem Biophys Res Commun. 1998;249:656–659. doi: 10.1006/bbrc.1998.9000. [DOI] [PubMed] [Google Scholar]
  • 32.Yumura S, Ueda M, Sako Y, Kitanishi-Yumura T, Yanagida T. Multiple mechanisms for accumulation of myosin II filaments at the equator during cytokinesis. Traffic. 2008;9:2089–2099. doi: 10.1111/j.1600-0854.2008.00837.x. [DOI] [PubMed] [Google Scholar]
  • 33.Pramanik MK, Iijima M, Iwadate Y, Yumura S. PTEN is a mechanosensing signal transducer for myosin II localization in Dictyostelium cells. Genes Cells. 2009;14:821–834. doi: 10.1111/j.1365-2443.2009.01312.x. [DOI] [PubMed] [Google Scholar]
  • 34.Saltel F, et al. New PI(4,5)P2- and membrane proximal integrin-binding motifs in the talin head control beta3-integrin clustering. J Cell Biol. 2009;187:715–731. doi: 10.1083/jcb.200908134. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Goksoy E, et al. Structural basis for the autoinhibition of talin in regulating integrin activation. Mol Cell. 2008;31:124–133. doi: 10.1016/j.molcel.2008.06.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Martel V, et al. Conformation, localization, and integrin binding of talin depend on its interaction with phosphoinositides. J Biol Chem. 2001;276:21217–21227. doi: 10.1074/jbc.M102373200. [DOI] [PubMed] [Google Scholar]
  • 37.Even-Ram S, et al. Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat Cell Biol. 2007;9:299–309. doi: 10.1038/ncb1540. [DOI] [PubMed] [Google Scholar]
  • 38.Wang Y, et al. Loss of PIP5KIgamma, unlike other PIP5KI isoforms, impairs the integrity of the membrane cytoskeleton in murine megakaryocytes. J Clin Invest. 2008;118:812–819. doi: 10.1172/JCI34239. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Galdeen SA, Stephens S, Thomas DD, Titus MA. Talin influences the dynamics of the myosin VII-membrane interaction. Mol Biol Cell. 2007;18:4074–4084. doi: 10.1091/mbc.E06-07-0586. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Tuxworth RI, Stephens S, Ryan ZC, Titus MA. Identification of a myosin VII-talin complex. J Biol Chem. 2005;280:26557–26564. doi: 10.1074/jbc.M503699200. [DOI] [PubMed] [Google Scholar]

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