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Using two-colour imaging and high resolution TIRF microscopy, we investigated the assembly and maturation of nascent adhesions in migrating cells. We show that nascent adhesions assemble and are stable within the lamellipodium. The assembly is independent of myosin II but its rate is proportional to the protrusion rate and requires actin polymerization. At the lamellipodium back, the nascent adhesions either disassemble or mature through growth and elongation. Maturation occurs along an α-actinin–actin template that elongates centripetally from nascent adhesions. α-Actinin mediates the formation of the template and organization of adhesions associated with actin filaments, suggesting that actin crosslinking has a major role in this process. Adhesion maturation also requires myosin II. Rescue of a myosin IIA knockdown with an actin-bound but motor-inhibited mutant of myosin IIA shows that the actin crosslinking function of myosin II mediates initial adhesion maturation. From these studies, we have developed a model for adhesion assembly that clarifies the relative contributions of myosin II and actin polymerization and organization.
Adhesion assembly and turnover are highly dynamic and coordinated processes essential for cell migration1,2. Adhesions serve as traction points for cell translocation and mediate a network of signalling events that regulate protrusion, contractility and attachment1–7. Although much is known about the functions of adhesions in developing and responding to traction and the signalling networks they regulate, less is known about the mechanisms by which adhesions assemble and turnover.
In migrating cells, protrusions are generated by actin polymerization at the front1,8. Protrusions consist of two structurally and kinetically distinct actin networks9,10. The lamellipodium comprises a treadmilling network of Arp2/3-mediated, branched actin filaments, whereas the lamellum consists of actin filament bundles9,10. The location and mechanism of adhesion assembly are unclear. Recent evidence suggests that adhesions form at the base of the lamellipodium11,12 in response to waves of actomyosin-generated force and halted protrusive activity12. However, periodic contractions are not observed in all cells13,14, particularly highly protrusive or rapidly migrating cells, suggesting alternative mechanisms of adhesion assembly.
Adhesions are thought to mature by a sequential mechanism coupled to tension or myosin II activity12,15. Inhibition of several signalling components, including FAK, Src and ERK kinases, stops adhesion turnover and promotes maturation1, suggesting a role for phosphorylation-mediated affinity changes of adhesion components. However, contractile force has also emerged as a major regulator. Application of force induces adhesion growth, whereas inhibition of actomyosin contractility decreases adhesion size16–18. Myosin II is also an endpoint of the pathways regulated by Rho GTPases, which are downstream hubs for migration-related signalling pathways19,20.
Our goals in this study were to define the steps and mechanisms underlying the early events in adhesion formation during migration and determine the role of the actin cytoskeleton and myosin II contractility in these processes. These studies were facilitated by the identification of small adhesions near the leading edge of motile cells6,14,21,22 and rapid two-colour TIRF microscopy. We show that nascent adhesions assemble in the lamellipodium, are stable only within the lamellipodial dendritic actin network, and do not require myosin II activity. These adhesions grow and elongate at the lamellipodium–lamellum interface along an actin–α-actinin template. Finally, we show that the actin crosslinking property of myosin II contributes prominently to adhesion maturation. On the basis of these observations, we developed a model of adhesion assembly that provides a conceptual framework for the formation of adhesions in protrusions.
We and others have reported that small, dynamic adhesions are present near the leading edge of protrusions in motile cells6,14,21,22. These structures colocalize with integrins (Fig. 1a), contain molecules commonly associated with adhesions, contain phosphorylated FAK (Tyr 397) and paxillin (Tyr 31), and associate closely with the substratum6,14,21. They also follow the edge of the protrusion as it moves forward. To determine whether the adhesions were sliding outward during protrusion or undergoing rapid assembly and disassembly (turnover), we observed them at high temporal and spatial resolution using TIRF microscopy.
CHO.K1 cells expressing paxillin–mEGFP and plated onto fibronectin (2 μg ml−1) generated broad protrusions with an array of small, punctate adhesions near the leading edge (Fig. 1b; Supplementary Information, Movie 1). These adhesions were stationary relative to the substratum and disassembled as new, nascent adhesions formed in front of them. As a result, the periphery of active protrusions was always decorated with small adhesions undergoing continuous turnover.
The kinetics of formation and disassembly of these adhesions were quantified by measuring the fluorescence intensity for marker proteins, such as paxillin or vinculin1. Using a frame rate of 1 s and correcting for photobleaching, the kinetics revealed three distinct phases: assembly, stability and disassembly. The average life of these adhesions was 76.1 ± 22.0 s (Fig. 1b) and the rates of assembly and disassembly were 1.26 ± 0.10 min−1 and 0.73 ± 0.07 min−1, respectively (Fig. 1d). Higher fibronectin concentrations and highly protrusive phenotypes 6,14 had negligible effects on these rates.
The transient stability (11.8 ± 6.2 s) suggests that nascent adhesions do not grow indefinitely but instead stop at a target state. In support of this hypothesis, the relative intensity of the adhesions within a region peaked and remained at this plateau until disassembly. The nascent adhesions also reached a plateau at a common size of 0.19 ± 0.01 μm2, which is close to the diffraction limit, suggesting that their true size may be smaller than their observed size.
Finally, all components studied so far enter and leave nascent adhesions simultaneously. Paxillin, vinculin, FAK, G protein-coupled receptor kinase-interacting protein 1 (GIT1) and zyxin, for example, showed indistinguishable relative assembly and disassembly kinetics (Fig. 1e). This suggests that they enter and exit nascent adhesions either as preformed clusters or individually in response to a common or kinetically indistinguishable event(s).
Recently, we reported that knockdown of myosin II by RNA interference (RNAi) produces protrusions that have only a rim of small adhesions near the leading edge15. This prompted us to investigate whether nascent adhesions can form in the absence of myosin II activity. Knockdown of myosin IIA (MIIA) resulted in adhesions near the leading edge that turned over continuously, showing the same three discrete phases described above for nascent adhesions in wild-type cells. The average life was 71.3 ± 21.5 s, with assembly and disassembly rates of 1.49 ± 0.10 min−1 and 0.85 ± 0.11 min−1, respectively, a stability time of 12.2 ± 6.4 s, and an average size of 0.20 ± 0.01 μm2. These properties are comparable to nascent adhesions from wild-type cells, indicating that they are similar, if not identical, adhesions.
To demonstrate more rigorously that nascent adhesions can form without myosin II activity, we generated double RNAi knockdown of MIIA and MIIB. More than 90% of the total myosin II expression was inhibited, as assessed by residual fluorescence intensity in analysed cells (Fig. 2a). The cells no longer had large, stable adhesions in central regions, but the protrusions still contained small, dynamic adhesions at the periphery (Fig. 2b). We also treated MIIA knockdown cells with blebbistatin (an inhibitor of myosin II ATPase activity) to inhibit residual activity of MIIA and MIIB, the other isoform present in these cells. Blebbistatin-treated MIIA knockdown cells retained a rim of dynamic adhesions near the leading edge (Fig. 2c). Their life (74.1 ± 28.2 s) and size (0.22 ± 0.01 μm2) were comparable to those of both wild-type and myosin II knockdown cells. Finally, wild-type or MIIA knockdown cells plated in the presence of blebbistatin also had nascent adhesions near the leading edge of protrusions (data not shown). Taken together, these data show that the formation and turnover of the small, dynamic adhesions near the leading edge are independent of myosin II.
Finally, we compared the nascent adhesions with focal complexes, which are small adhesions at the cell periphery induced by constitutively activated (CA) Rac23. The focal complexes were about two times larger than nascent adhesions (data not shown), appeared mainly at the lamellipodium–lamellum interface (Supplementary Information, Fig. S1a) and did not turn over. Moreover, addition of blebbistatin induced disassembly of focal complexes into smaller adhesions that are indistinguishable from nascent adhesions (Supplementary Information, Fig. S1b). Thus, CA Rac-induced focal complexes are myosin II-dependent24 and distinct from the smaller, myosin II-independent nascent adhesions. This also shows that not all punctate, peripheral adhesions are nascent adhesions.
We next investigated where nascent adhesions form and what determines their lifespan. The lamellipodium was identified as the dense, 1–3-μm actin-rich band near the leading edge that contains barbed-end actin, Arp2/3 and cofilin10,11 (Supplementary Information, Fig. S2). When paxillin–mOrange (or vinculin–mOrange) and GFP–actin were co-expressed, time-lapse TIRF microscopy revealed that the nascent adhesions form and reside in the lamellipodium (Fig. 3a; Supplementary Information, Movie 2).
Dual-colour imaging showed that the fluorescence intensity of paxillin (or vinculin) increased 10–35 s after the actin began accumulating in the lamellipodium (Fig. 3b) but not at its outer edge. After assembly, the nascent adhesions were transiently stable for 11.8 ± 6.2 s, which correlates with the time a fully assembled adhesion resides within the boundaries of the lamellipodium. Moreover, the adhesions began to disassemble at the rear of the lamellipodium as the dense actin band passed by them (Fig. 3a, b; Supplementary Information, Movie 2).
To determine whether the rates of nascent adhesion assembly, disassembly and protrusion were coupled, we compared the rate of increase or decrease in paxillin–mEGFP fluorescence in nascent adhesions with the rate of leading edge extension adjacent to the adhesions during phases of protrusion. The protrusion rates varied 3–5-fold within and among cells. The rate of adhesion assembly was linearly proportional to the local rate of leading edge extension (Fig. 3c). In contrast, the rate of adhesion disassembly was not affected by the speed of protrusion, indicating that the kinetics of adhesion disassembly and actin polymerization are not mechanistically linked. Interestingly, the relationship between the adhesion assembly and local protrusion rates was sustained under a variety of conditions, including plating in high fibronectin concentrations and myosin II inhibition, where retrograde flow is inhibited (data not shown). Taken together, these data show that the rate of nascent adhesion formation is directly coupled to the rate of protrusion and suggest that there is a link between nascent adhesion assembly and actin polymerization in the lamellipodium.
To ascertain whether these adhesions required dendritic actin for their stability, we disrupted the lamellipodium with cytochalasin-D, which caps barbed ends and inhibits actin polymerization. Addition of cytochalasin-D (1 μM) inhibited protrusion immediately; no new adhesions formed and those remaining in the lamellipodium were stabilized. However, as the dense actin band characterizing the lamellipodium narrowed and collapsed, the adhesions in the back disassembled as the band passed behind them (Fig. 3d). This shows that continued adhesion assembly requires actin polymerization, that these adhesions are only stable within the lamellipodium and that the adhesions disassemble when the dendritic actin of the lamellipodium moves past them.
Whereas some nascent adhesions diassemble as the lamellipodium moves past them, others mature by growth and elongation at the lamellum–lamellipodium interface during pauses in protrusion (Fig. 4a; Supplementary Information, Movie 3). These maturing adhesions arise from the nascent adhesions (Supplementary Information, Movie 4). The oriented, centripetal nature of their growth suggests that the adhesions mature along a structural track or template. To investigate this, we imaged CHO.K1 cells expressing GFP–actin. We observed small, linear actin filaments emanating centripetally from the halted protrusion (Fig. 4a; Supplementary Information, Movie 5). This thin line of actin was most readily observed with a promoter-truncated GFP–actin vector25 that results in low expression levels, minimizing cytoplasmic background, suggesting that these structures consist of only a few actin filaments. Dual-colour imaging using GFP–actin and paxillin–mOrange revealed that short actin filaments seemed to grow from the nascent adhesions, which subsequently underwent elongation, raising the possibility that they arise from adhesion-associated actin polymerization (Fig. 4b; Supplementary Information, Movie 6). Dual-colour imaging of cells expressing GFP–α-tubulin and paxillin–mOrange did not show any colocalization of microtubules with the elongating adhesions (data not shown).
To determine the mechanism of adhesion maturation, we first compared the relative kinetics of elongation using GFP- and mOrange- or mCherry-labelled pairs of some core adhesion molecules. These studies revealed a hierarchy of entry into the elongating adhesion. α-Actinin and actin elongated simultaneously (Fig. 4c). Paxillin and talin entered after actin and α-actinin but slightly before vinculin, suggesting that vinculin entry into elongating adhesions requires the pre-assembly and/or activation of another adhesion component(s) (Fig. 4c). Tensin entered the adhesions after vinculin and its concentration increased as the adhesion matured further (data not shown). Finally, MIIA approached elongating actin filaments from the central part of the cell and linked up with the filaments as they elongated and thickened (Supplementary Information, Movie 7).
The simultaneous, initial accumulation of α-actinin and actin at sites of adhesion elongation points to a key role for α-actinin in the maturation process. To investigate this, we inhibited α-actinin expression in CHO.K1 cells using RNAi. Immunoblots revealed that the RNAi efficiency was close to 85% (Fig. 5a), similar to the transfection efficiency. Immunofluorescence microscopy showed that knockdown cells expressed undetectable amounts of α-actinin (Fig. 5b). The protrusion rate in α-actinin-deficient cells was reduced and discontinuous (Supplementary Information, Fig. 3). The actin filament bundles in protrusions no longer showed a centripetal orientation, and instead, were short and more randomly oriented (Fig. 5c, d, top panels). Small, punctate adhesions formed near the leading edge but they neither turned over nor matured into larger, elongated adhesions, even when the protrusion was halted (Fig. 5d, e; Supplementary Information, Movie 8). Interestingly, paxillin was distributed in punctate structures along the actin filaments throughout the cell rather than at the ends of large actin bundles (Fig. 5c). Expression of an RNAi-insensitive α-actinin–GFP rescued the phenotype produced by α-actinin knockdown (Fig. 5e and data not shown). These observations indicate that α-actinin has a crucial role in actin organization and adhesion maturation.
As α-actinin organizes F-actin through its crosslinking activity26, its marked effect on adhesion assembly prompted us to examine whether the crosslinking property of myosin II has a similar role. To separate the contribution of myosin II-dependent actin crosslinking from its contractile activity on adhesion maturation, we expressed paxillin–mOrange in MIIA knockdown CHO.K1 cells together with MIIAN93K27. In this myosin IIA mutant, ATPase activity is inhibited by 90%, and motor activity is inhibited completely27; but it constitutively binds to actin14,28. The adhesions elongated centripetally and were comparable in size and morphology to the control cells rescued with wild-type GFP–MIIA (Fig. 6b, c, f; Supplementary Information, Movie 9). This contrasts with the phenotype in MIIA knockdown cells in which adhesion maturation was inhibited14 (Fig. 6a; Supplementary Information, Movie 9). To inhibit residual MIIB contractility in MIIAN93K-rescued cells, we used blebbistatin, which inhibits the ATPase activity of myosin II but, unlike the mutant, does not sustain strong actin binding. We still observed elongated adhesions in the protrusions (Fig. 6d; Supplementary Information, Movie 10). This suggests that myosin II-mediated actin crosslinking has an important role in adhesion maturation.
To further establish the importance of actin crosslinking, we investigated whether overexpressing one of the two crosslinking proteins, α-actinin and myosin II, can compensate for absence of the other. Overexpression of α-actinin–GFP in MIIA-deficient cells induced thick α-actinin bundles (Fig. 6e, green), which is consistent with its actin crosslinking activity29. This treatment also restored growth and centripetal elongation of the adhesions, although not to the levels seen in controls (Fig. 6e, f). In other respects, however, the cells still exhibited properties of the MIIA-deficient phenotype, for example, an inability to retract at the cell rear (data not shown).
In the reciprocal experiment, overexpression of MIIAN93K, which bundles actin but is not contractile, restored the maturation of adhesions in α-actinin knockdown cells (Fig. 7a, b, d; Supplementary Information, Movie 11). Similar results were observed when wild-type MIIA was overexpressed in the α-actinin knockdown cells (Fig. 7c; Supplementary Information, Movie 11). Together, these observations show that actin crosslinking promotes adhesion maturation.
Our observations support a working model for adhesion assembly during cell migration (Fig. 8a). Nascent adhesions assemble in the lamellipodium in a single concerted step as the protrusion advances. The assembly rate is linked to the protrusion rate and probably to actin polymerization. The nascent adhesions are stable only within dendritic actin and disassemble as the wave of depolymerizing actin at the rear of the lamellipodium passes by them with the advancing protrusion. Both the assembly and stability of the nascent adhesions in the lamellipodium are myosin II-independent. When protrusion pauses, a subset of nascent adhesions grow and elongate centripetally from the base of the lamellipodium. The elongation is directed by actin filaments. α-Actinin associates with the emerging actin filaments and organizes and orients them centripetally. These α-actinin–actin filaments function as a template for the hierarchical addition of other adhesion components. The actin crosslinking properties of myosin II also have a major role in the formation of the template and initial stages of adhesion maturation.
Our data show that nascent adhesions assemble in the lamellipodium as diffraction-limited puncta and then undergo myosin II-dependent maturation near the lamellipodium–lamellum interface. The prominent, readily visualized adhesions (for example, focal complexes reported to be present at the lamellipodium–lamellum interface11,12) differ from nascent adhesions as they are larger, more stable and myosin-II-dependent. The CHO cell is particularly useful for studies of adhesion assembly as the nascent adhesions have long lifespans and only a fraction of them mature. Presumably, this is due in part to lowered or localized myosin II activity. In contrast, nascent adhesions in MEFs or U2OS cells reside in the dendritic actin for only a few seconds (Supplementary Information, Fig. S4, Movie 12 and data not shown); most stabilize and mature at the lamellipodium–lamellum interface. Myosin II is a determining factor as blebbistatin inhibits the probability of adhesion maturation and promotes the assembly and turnover of the nascent adhesions in both cell types (Supplementary Information, Fig. S4 and data not shown). Conversely, MIIA overexpression in CHO.K1 cells inhibits protrusion and increases the probability that nascent adhesions grow and elongate, which occurs almost immediately after they form14 (Supplementary Information, Movie 13). In all of these cells, however, the elongating adhesions in protrusions arise from nascent adhesions. Previous observations also implicate myosin II in adhesion assembly12,20 and the periodic interruption and retraction of protrusions12,14,30.
The formation and stability of nascent adhesions within the lamellipodium, the correlation between the assembly and protrusion rates, and the inhibition of both protrusion and nascent adhesion assembly by cytochalasin-D suggest that adhesion assembly is mechanistically and kinetically linked to actin polymerization in the lamellipodium; this is also observed when net protrusion is driven without myosin II mediated retrograde flow. Two components of nascent adhesions, vinculin and FAK, interact with the Arp2/3 complex, which nucleates actin polymerization within dendritic actin and thereby provide a potential mechanism for the coupling31–33. A recent study showed actin-polymerization-based, protrusion-independent lateral movement of integrins to filopodia-like ripples in the lamellipodium34; however, they did not report a similar mechanism for adhesions outside of ripples.
The lamellipodium–lamellum interface emerges as a critical region where adhesion fate is determined. Nascent adhesions either disassemble or mature as the dendritic actin passes by them. It is also the region where the dendritic actin turns over35, suggesting that nascent adhesions are physically linked to dendritic actin and disassemble in response to its turnover.
We present a mathematical model, which assumes that an adhesion precursor binding to the dendritic actin is a limiting step for adhesion assembly, and that the adhesion disassembly is mechanically coupled to dendritic actin disassembly (Supplementary Information, Text). We also assumed that the Arp2/3 mediated branching takes place near the leading edge, either on adhesions, where the branching points are firmly anchored to the substratum, or in the immediate vicinity of adhesions (so that ‘daughter’ filaments branch off ‘mother’ filaments anchored to the substratum). The solutions of mathematical equations derived from these assumptions reproduced our quantitative observations. That is, as the branching rate is almost constant near the leading edge, total dendritic actin filament length builds up almost linearly from the leading edge towards the rear. Adhesion assembly lags by the characteristic time of the precursor binding to actin behind the front of the actin band. Actin–ATP hydrolysis, cofilin action and decrease of the branching activity behind the leading edge determine the rear of the dendritic actin band, where both actin and adhesions disassemble in synchrony (Fig. 8b). Consistent with our data, the model further predicts that the adhesion assembly rate is proportional to the leading edge extension rate, independent of the disassembly rate from the speed of protrusion, and correlates with the inverse duration of the adhesion stability phase with the speed of protrusion.
Nascent adhesions at the lamellipodium–lamellum interface can also grow and elongate, presumably in response to a changing or different organization of actin. Elongated, centripetal adhesions at the periphery are a hallmark of maturing adhesions22,36. Short, linear actin filaments emanate from this region and provide a template for the maturation of nascent adhesions. These filaments could arise from either the reorganization of existing filaments in the dendritic actin or from local polymerization.
α-Actinin and myosin II are essential for the formation and organization of the actin template. In the absence of α-actinin, actin filaments are abnormally short, discontinuous and misoriented. α-Actinin also positions adhesions along actin filaments as adhesion components in α-actinin knockdown cells are no longer restricted to the ends of actin filaments and appear as puncta spread along the entire filament. α-Actinin has been reported previously to participate in the later stages of adhesion maturation by forming large stress fibres and adhesions at their ends29,37,38. An earlier observation that α-actinin is a late entry into the larger adhesions was made with wide-field optics, which would not have seen the templates and early events described here38.
Myosin II is also required for the growth and elongation of nascent adhesions. Neither actin templates nor the elongation of nascent adhesions are observed in MIIA-deficient or inhibited cells14. Several reports suggest that myosin II-mediated contraction has a major role in the maturation of adhesions by tension-induced alterations in the conformation of adhesion-related proteins14,15,24,39,40. Our study shows that the actin crosslinking activity of myosin II is important in the initial stages of adhesion maturation, presumably by organizing and clustering actin and actin-associated adhesion components. Thus, although myosin II-mediated contractility seems to promote the formation of thick actin filament bundles and large adhesions at later stages of maturation, the contractile activity could function synergistically with myosin II-mediated actin crosslinking at early stages by organizing actin filaments. Myosin II-mediated actin crosslinking can also transmit distally generated actomyosin contractility to adhesions. Others have also ascribed roles for myosin II crosslinking: an ATPase-deficient myosin II restores cortical integrity in Dictyostelium discoideum41; a motor-impaired MIIB mutant rescues hydrocephalus in MIIB knockout mice42; the bundling function of myosin in adhesion assembly was proposed previously43.
In summary, the data presented here provide new insights into the mechanism of adhesion assembly. They identify and characterize a new class of adhesions, ‘nascent adhesions’, which reside in the lamellipodium and serve as precursors for other adhesions in the protrusion. Moreover, they clarify the role of myosin II in adhesion maturation, lead to a ‘template’ model for centripetal adhesion elongation along actin–α-actinin filaments, and demonstrate the importance of the actin-bundling activity of myosin II.
To generate α-actinin siRNA, the oligonucleotide GGAGATCAATGGCAAATGG, corresponding to nucleotides 2003–2021 of rat α-actinin1 (NM_031005) was inserted into the appropriate pSUPER cassette according to the vector manufacturer’s instructions (Oligoengine). pSUPER-MIIA and pSUPER-MIIB have been described previously14. siRNA-insensitive α-actinin was generated by site-directed mutagenesis (Quickchange kit, Stratagene) introducing two silent mutations (ATC to ATT: Ile to Ile; AAC to AAT: Asn to Asn) in the RNAi target region of human α-actinin1, which shares 100% homology with rat.
Promoter-truncated GFP–actin, GFP–MIIA, GFP–cofilin, GFP–tensin, GFP–zyxin and human β1 integrin cDNA were gifts from Tim Mitchison (Harvard Medical School, Boston, MA)25, Robert Adelstein (National Institutes of Health, Bethesda, MD)44, John Condeelis (Albert Einstein College of Medicine, New York, NY)45, David Brautigan (University of Virginia, Charlottesville, VA)46, Klemens Rottner (Ludwig-Maximilians-Universität München, Germany)47 and Martin Humphries (University of Manchester, UK), respectively. GFP–MIIA-N93K14, paxillin–GFP and α-actinin–GFP38, GFP–vinculin and GFP–FAK and GIT1 have been described previously1. Where indicated, GFP was replaced by mCherry from Roger Tsien48 or CoralHue monomeric Kusabira Orange (mOrange, MBL). Rabbit polyclonal antibodies against MIIA (1:1,000), MIIB (1:1,000) and GIT1 (1:1,000) were obtained from Covance; α-actinin (1:100, mouse, IgG1) from SantaCruz Biotechnology and TS2/16 (β1 integrin, 1mg ml−1) from Biolegend.
CHO.K1 cells, mouse embryonic fibroblasts and U2OS osteosarcoma cells were cultured under standard conditions and transfected using Lipofectamine (Invitrogen)14. For co-transfection experiments, plasmids containing the siRNA sequences were used in 10:1 excess to GFP or mCherry-containing plasmids to ensure knockdown in fluorescence-positive cells.
Cells were plated onto fibronectin-coated cover-slips (2 μg ml−1) for 60 min, fixed using 4% paraformaldehyde and permeabilized with 0.5% Triton X-100 for 5 min. Coverslips were incubated with primary antibodies and a species-appropriate secondary antibody coupled to either Alexa488 or Alexa568 (Invitrogen). Barbed-end staining was performed as described elsewhere49.
Cells were plated on 2 μg ml−1 fibronectin-coated glass-bottomed dishes (migration-promoting conditions) in CCM1 for 1 h and maintained at 37 °C at pH 7.4. Confocal images were collected on an Olympus Fluoview 300 microscope (1.45 NA (oil) PlanApo ×60 TIRFM objective (Olympus)). GFP and mCherry/mOrange were excited using the 488-nm laser line of an Ar ion laser and the 543-nm laser line of a He-Ne laser (Melles Griot), respectively. A Q500LP dichroic mirror (Chroma Technology) was used for GFP-labelled cells. For dual-colour imaging, a green-red cube (488/543/633) with a DM570 dichroic mirror (Chroma Technology) was used. Fluorescence images were acquired using Fluoview software (Olympus).
TIRF images were acquired using an Olympus IX70 inverted microscope (1.45 NA (oil) PlanApo ×60 TIRFM objective), fitted with a Ludl modular automation controller (Ludl Electronic Products) and controlled by Metamorph (Molecular Devices). The excitation laser lines used were as described for confocal microscopy. Mirrors and filters were supplied by Chroma Technology. A dichroic mirror (HQ485/30) was used for GFP-labelled cells. For dual GFP–mCherry/mOrange acquisition, a polychroic mirror (Z488/543rpc) and a dual emission filter (Z488/543) were used. Also, HQ525/50 and HQ620/60 emission filters were used for GFP and mCherry/mOrange, respectively. For simultaneous GFP–mCherry acquisition, Dual-View (MAG Biosystems) was utilized. All images were acquired with a charge-coupled device camera (Retiga Exi; Qimaging) and analysed using Metamorph or ImageJ (NIH).
ImageJ was used to measure changes in fluorescent intensity of individual nascent adhesions over time in cells expressing fluorescent-tagged adhesion proteins1. Background and photobleaching corrections were applied to obtain true intensities of the adhesions. Assembly and disassembly rates were plotted and calculated using Microsoft Excel (Microsoft Corporation) or SigmaPlot (SPSS)1. Mean lifespan with standard deviation were measured from 30–50 individual adhesions in seven to fourteen cells. The elongation index of maturing adhesions was determined by measuring the long axis of the adhesions (that is, perpendicular to the membrane) and dividing it by the maximal perpendicular axis.
Protrusion was quantified using kymography14. Images were captured every second for 3 min. Kymographs were generated using Metamorph software along 1-pixel-wide regions oriented in the protrusion direction and perpendicular to the leading edge.
Figure S1 Activated Rac induced focal complexes differ from nascent adhesions. CHO.K1 cells expressing V12Rac, either without (A) or with 20 μM blebbistatin (B), were co-transfected with GFP-actin (left) and paxillin-mOrange (right). Scale bar = 10 μm. Color inserts show the relative positioning of paxillin-containing focal complexes (A) or nascent adhesions (B) with respect to the lamellipodium. Green, actin; magenta, paxillin. Note that the focal complexes reside the lamellipodium-lamellum interface and disassemble into smaller, lamellipodial localized nascent-like adhesions the presence of blebbistatin.
Figure S2 Molecular markers of the lamellipodium of migrating CHO.K1 cells. Characterization of dendritic actin of the lamellipodia. mRFP-actin (magenta) colocalize with GFP-cofilin and immunostaining of Arp3 (green). Barbed-end actin (green) outlines fluorescent phalloidin (magenta) at the leading edge.
Figure S3 Quantification of protrusion rates in α-actinin-knockdown cells. (A) Kymographs from control (pSUPER) and α-actinin-depleted cells. The knockdown cells were identified by coexpression of a GFP marker, imaged in brightfield, and then analyzed by kymography. (B) Quantification of protrusion rates from kymographs from (A). Average protrusion rates measured from >12 cells (three to five protrusions/cell) from four independent experiments. Error bars represent ±SEM.
Figure S4 The rapid assembly of nascent adhesions in MEFs. TIRF time-lapse images of paxillin-GFP in MEFs show the rapid assembly (green arrows) of new nascent adhesions near the leading edge. Most nascent adhesions undergo immediate maturation and do not turnover (red arrows). Scale bar = 3 μm. Elapsed time is in seconds. TIRF images of GFP-actin (green) and paxillin-mOrange (magenta) show nascent adhesions (green arrows) forming in MEFs and residing exclusively in the lamellipodium. Scale bar = 3μm.
Paxillin-containing nascent adhesions assemble and disassemble rapidly during protrusions. This movie corresponds to Fig. 1A. Images of paxillin-GFP collected every 1 sec using TIRF microscopy. Magenta and green arrows indicate representative assembling and disassembling adhesions, respectively. 10 frames/sec shown.
Rescue of the MIIA knockdown with the non-contractile MIIA mutant N93K and blebbistatin. This movie corresponds to Fig. 6D. Images of paxillin-mOrange collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
Rescue of the α-actinin knockdown by the high actin affinity, non-contractile mutant N93K-MIIA or over expression of MIIA wild-type. This movie corresponds to Fig. 7A–C. Images of paxillin-mOrange collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
Paxillin-containing nascent adhesions assemble and mature rapidly during protrusions in MEFs. This movie corresponds to Supplementary Fig. 40. Images of paxillin-GFP collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
Myosin II regulates the probability and rate of nascent adhesion maturation. Left, MIIA-deficient CHO.K1cell. This movie is from Vicente-Manzanares et al., J. Cell Biol. 176(5)573–580, Suppl. Movie 7; center, wild-type; right, overexpression of MIIA in a CHO.K1 cell. Images of paxillin-GFP were collected every 5 s using TIRF microscopy. 24 frames/s are shown.
Nascent adhesions reside in the lamellipodium. This movie corresponds to Fig. 3A. Green indicates the lamellipodium marked by GFP-actin; magenta indicates nascent adhesions marked by paxillin-mOrange. Images collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
Elongation of adhesions when a protrusion halts. This movie corresponds to Fig. 4A. Images of paxillin-GFP collected every 5 sec using TIRF microscopy. 24 frames/sec shown.
Long-term imaging of nascent adhesions showing formation, turnover and maturation. Images of paxillin-GFP collected every 5 sec using TIRF microscopy. 24 frames/sec shown.
Elongation of actin filaments when a protrusion halts. This movie corresponds to Fig. 4A. Images of GFP-actin collected every 1 sec using TIRF microscopy. 24 frames/sec shown.
Actin template elongates from maturing nascent adhesions. This movie corresponds to Fig. 4B. Merged images of GFP-actin (green) and paxillin-mOrange (magenta) collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
MIIA links to actin templates distal to the initial α-actinin. Merged images of GFP-actin (green) and mCherry-MIIA (magenta) collected every 5 s using TIRF microscopy. 24 frames/sec shown.
Adhesion turnover and maturation are inhibited by α-actinin knockdown. This movie corresponds to Fig. 5D. Images of GFP-actin and paxillin-mOrange collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
Rescue of MIIA-knockdowns with wild-type-MIIA or the high actin affinity, non-contractile mutant, N93K-MIIA. This movie corresponds to Figs. 6A-C. Images of paxillin-mOrange collected every 3 sec using TIRF microscopy. 24 frames/sec shown.
We thank Laura E. Chopko for helping to characterize the α-actinin knockdown. Also, we thank Hannelore Asmussen for technical assistance with substrate preparation. This work was supported by NIH grants GM23244 (AFH), the Cell Migration Consortium (U54 GM064346) and NSF grant DMS-0715729 (AM).
Note: Supplementary Information is available on the Nature Cell Biology website.
AUTHOR CONTRIBUTIONSC.K.C and M.V.-M. designed and performed the experiments and wrote the paper; J.Z. and l.A.W. assisted with the research; A.M. developed the mathematical model and its presentation; A.R.H. designed the experiments and wrote the paper.
COMPETING FINANCIAL INTERESTS
The authors declare no competing financial interests.