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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
J Biol Chem. Author manuscript; available in PMC Nov 17, 2008.
Published in final edited form as:
PMCID: PMC2583367
NIHMSID: NIHMS76707

Structure/Function Analysis of the Interaction of Phosphatidylinositol 4,5-Bisphosphate with Actin-capping Protein

IMPLICATIONS FOR HOW CAPPING PROTEIN BINDS THE ACTIN FILAMENT*,S

Abstract

The heterodimeric actin-capping protein (CP) can be inhibited by polyphosphoinositides, which may be important for actin polymerization at membranes in cells. Here, we have identified a conserved set of basic residues on the surface of CP that are important for the interaction with phosphatidylinositol 4,5-bisphosphate (PIP2). Computational docking studies predicted the identity of residues involved in this interaction, and functional and physical assays with site-directed mutants of CP confirmed the prediction. The PIP2 binding site overlaps with the more important of the two known actin-binding sites of CP. Correspondingly, we observed that loss of PIP2 binding correlated with loss of actin binding among the mutants. Using TIRF (total internal reflection fluorescence) microscopy, we observed that PIP2 rapidly converted capped actin filaments to a growing state, consistent with uncapping. Together, these results extend our understanding of how CP binds to the barbed end of the actin filament, and they support the idea that CP can “wobble” when bound to the barbed end solely by the C-terminal “tentacle” of its β-subunit.

Actin polymerization is essential for many biological phenomena, including the cell migration that occurs as part of chemotaxis, development, and cancer. Cells regulate the location and timing of actin polymerization with remarkable precision. In cells, actin filaments grow primarily by the addition of subunits to free, barbed ends of filaments. Free barbed ends can be created by actin nucleation factors, such as Arp2/3 complex and formins, and by actin-severing proteins, such as ADF/cofilin and gelsolin. Capping proteins bind to free barbed ends and thereby halt the polymerization process.

The α/β heterodimeric capping protein, referred to here as CP,5 is a capper found in essentially every eukaryotic organism and in every vertebrate cell and tissue type (reviewed in Ref. 1). In vertebrate striated muscle, a sarcomere-specific isoform of CP at the Z-line caps the barbed ends of the actin-based thin filaments, leading to the name CapZ. Cells lacking CP show a range of morphologic and developmental defects, based on improper actin assembly and cell motility. In Drosophila, loss of CP is lethal during embryogenesis.

Typical vertebrate cultured cells contain lamellipodial and filopodial processes on their surface, which are based on actin filaments organized into branched networks and straight bundles, respectively. In each case, barbed ends are oriented predominantly toward the plasma membrane, and the local capping of barbed ends is crucial. The branched networks of filaments in lamellipodia are nucleated from Arp2/3 complex, as described in the dendritic nucleation model (2). Capping of the barbed ends over time is a necessary feature of this model in order to keep the filaments short, which makes the branched network stiff, and to restrict or “funnel” polymerization to the newly created barbed ends near the membrane. Inhibition of CP essentially abolishes the formation of lamellipodia and branched networks (3). In such cells, filopodia formation is greatly enhanced, which contributes to the notion that localized inhibition of CP at one place on the membrane may be essential for a filopodium to form.

In vitro, actin-based motility can be reconstituted from pure proteins with a mixture of Arp2/3 complex, ADF/cofilin, and CP (4). Here, the dependence of motility on CP concentration was a bell-shaped curve, suggesting that optimal levels of capping may be crucial for actin-based motility in cells. Indeed, Arp2/3-mediated actin assembly as part of endocytosis in yeast does depend on optimal CP activity (5).

CP is active in the absence of calcium and under a wide range of buffer conditions. However, cells contain a number of proteins that are able to bind to and inhibit CP, such as V-1/myotrophin, CKIP-1, CD2AP, and CARMIL (6-9). In addition, polyphosphoinositides such as phosphatidylinositol 4,5-bisphosphate (PIP2) can bind and inhibit CP (10). Of note, PIP2 appears to be able to remove CP from capped barbed ends (11), which may help to stimulate actin assembly in certain situations in cells. During platelet activation, an initial step appears to be release of CP from the actin cytoskeleton by PIP2 (12); later in the process, CP appears to return to the actin cytoskeleton, presumably binding newly formed barbed ends. Association of CP with the actin cytoskeleton is also seen with actin assembly in Dictyostelium cells responding to chemoattractant (13).

The crystal structure of CP shows the α- and β-subunits arranged with a pseudo-2-fold rotational axis of symmetry (14). In molecular dynamics simulations, the C-terminal region of the β-subunit is highly mobile, and the C-terminal region of the α-subunit remains closely apposed to the surface of the protein (6), as suggested by the crystal structure. A recent cryo-electron microscopy analysis of CP bound to the barbed end of the actin filament shows the top surface of CP in contact with actin, with the C-terminal regions of each subunit as likely sites of close contact (15). In biochemical studies, the C-terminal region of each subunit appears to be able to bind to the barbed end, and those interactions are independent of each other (16, 17). When both actin-binding sites are intact, CP binds to the barbed end with subnanomolar affinity because of a low off-rate constant (17).

Our current view of the interaction of CP with the filament barbed end includes the hypothesis that when CP is attached solely by its β-tentacle, it is able to wobble. This exposes the top surface of the protein, which includes the actin-binding site near the α-subunit C terminus. Recent findings with the protein V-1/myotrophin support this view (6). V-1 appears to bind the β-subunit C terminus, and V-1 inhibits capping but does not uncap, as predicted by the hypothesis. Another prediction of the hypothesis is that a molecule that is able to uncap should bind to the top surface of the protein, and this interaction should be independent of the β-tentacle.

In this study, to gain insight into how PIP2 binds CP, how PIP2 binding inhibits CP, and how CP binds actin, we wanted to perform a structure/function analysis with site-directed mutagenesis to identify regions of CP necessary for interaction with PIP2. We also performed computational modeling of CP/PIP2 interactions to help guide the mutagenesis and to provide independent and complementary evidence regarding the nature of the interaction. In addition, we sought new evidence for the uncapping effect of PIP2, with direct visualization of actin filament growth in real time by TIRF microscopy. Finally, we wanted to use these results to test the wobble model for the interaction of CP with the filament barbed end.

EXPERIMENTAL PROCEDURES

Chemicals and reagents were from Fisher Scientific and Sigma unless stated otherwise. PIP2 and 1,2-di-oleyoyl ethylene glycol (diacylglycerol) were obtained from Avanti (Alabaster, AL). Synthetic acyl-chain variants of PIP2 with diC4, diC8, and diC16 were obtained from Echelon Biosciences (Salt Lake City, UT). Concentrated phospholipid stock solutions were prepared and handled as described (28).

Plasmid Construction and Mutagenesis

Site-directed mutations were created in a chicken CP α1β1 pET bacterial expression plasmid (pBJ994) (18) by PCR using the QuikChange method (Stratagene, La Jolla, CA). Primers and plasmids are listed in supplemental Tables I and II, respectively. pET-3d/CPα(K256A)β, encoding a K256A mutation in the α-subunit, was constructed using forward primer KKT 255 and reverse primer KKT 256. pET-3d/CPα(R260A)β, encoding an R260A mutation in the α-subunit, was constructed using KKT 216 and 217. pET-3d/CPαβ(K95A), encoding a K95A mutation in the β-subunit, was constructed using KKT 205 and KKT 206. pET-3d/CPα(KR256,260AA)β, encoding a R256A,K260A mutation in the α-chain, was constructed using pET-3d/CPα(K256A)β as template and primers KKT 216 and KKT 217. pET3d/CPα(RK266, 268AA)β, encoding an RK266, 268AA mutation in the α-subunit, was constructed using KKT 214 and KKT 215. pET-3d/CPα(R256A,K260A)β(R225A), encoding a R256A,K260A mutation in the α-subunit and an R225A mutation in the β-subunit, was constructed with primers KKT 257 and KKT 258 using pET-3d/CPα(R256A,K260A)β as a template plasmid.

Proteins

CP was expressed in bacteria and purified as described (16) with minor modifications for mutants. Purified proteins were stored at −20 °C in 10 mm Tris-HCl, pH 7.5, 50 mm KCl, 1 mm MgCl2, 1 mm EGTA, 0.5 mm DTT, and 50% glycerol. Muscle actin was purified and labeled with pyrene as described (17). Spectrin-F-actin seeds were prepared from human erythrocytes as described (19). Actin capping assays were performed as described (16).

Rabbit skeletal muscle myosin (Cytoskeleton, Inc., Denver, CO) (10 μm) was dialyzed against 10 mm imidazole, pH 7, 0.5 m KCl, 10 mm EDTA for 2 h. Following a 1-h incubation with 1 mm N-ethylmaleimide on ice, 1 mm DTT was added for 1 h on ice. N-ethylmaleimide-myosin was dialyzed overnight against 50% glycerol in 10 mm imidazole, pH 7,0.5 m KCl, 10 mm EDTA,1 mm DTT and stored at −20 °C.

Fitting of Actin Polymerization Reactions

Binding constants for CP with actin and lipids were determined by least-squares fitting of full time course data using Berkeley Madonna 8.3 as described (17) with the following kinetic mechanism. In these reactions, A is actin monomer, Nb is free barbed end, CP is capping protein, and L is lipid. In this simple scheme, the capped barbed end, CPNb, can neither add nor lose actin subunits; the complex of CP with lipid, CPL, cannot interact with a barbed end.

A+Nbkk+ANb
REACTION 1
ANb+CPkcapk+capCPNb
REACTION 2
CP+Lkk+CPL
REACTION 3

For Reaction 1, k+ was 11.6 μm−1 s−1 and k_ was 1.4 s−1 (20) The rate constants for capping in Reaction 2 were determined by fitting the experimental data for seeded actin assembly in the presence of CP. A range of CP concentrations was used, generating a family of curves, and they were fit together. The rate constants for CP binding to lipid in Reaction 3 were determined by fitting a set of curves produced by addition of PIP2 at various concentrations.

Tryptophan Quenching Assay for CP/Lipid Interaction

The equilibrium dissociation constant, Kd, for the binding of CP to phospholipid was determined by measuring the quenching of intrinsic tryptophan fluorescence as described (21) with minor modifications. Briefly, fluorescence emission spectra (300–400 nm) were collected using a PTI Quantmaster spectrofluorometer (Photon Technology International, Birmingham, NJ) with excitation at 292 nm. The mixture of CP and phospholipid was incubated for 5 min before was fluorescence measured. The maximum fluorescence intensity (ΔF) was plotted against the total concentration of PIP2, and Kd was determined by least-squares fitting as described (21).

TIRF Microscopy

To image the polymerization of individual actin filaments, we used TIRF microscopy essentially as described (22). The system included an inverted microscope (IX-81, Olympus America, Center Valley, PA), an electron-multiplication back-thinned frame transfer charge-coupled device video camera (Model C9100-12, Hamamatsu Photonics, Bridgewater, NJ) with a 60× 1.45 numerical aperture PlanApo oil objective. SlideBook software (Intelligent Imaging Innovations, Denver, CO) operated the system and collected the images. Frames of 1500-ms duration were collected every 2–3 min. Measurements of filament length versus time were obtained.

To construct the microscope flow chamber, No. 0 sapphire coverslips were sonicated in a water bath sonicator for 45 min in 2% (v/v) VersaClean detergent in hot tap water, rinsed in hot tap water, and sonicated for 30 min in hot tap water. Coverslips were rinsed in distilled water, incubated for 3 h in 1 m KOH at 42 °C, rinsed in deionized distilled water, and incubated overnight in 1 m HCl at 42 °C. They were cooled to room temperature, rinsed in deionized distilled water, sonicated for 30 min in deionized distilled water, rinsed twice in 5 mm EDTA, and sonicated for 30 min in EDTA. They were rinsed in 70% ethanol, sonicated for 30 min in 70% ethanol, rinsed in absolute ethanol, sonicated for 30 min in absolute ethanol, and rinsed and stored in absolute ethanol.

Flow cell chambers were prepared as described (23). A clean sapphire coverslip was removed from ethanol and dried. Parafilm strips were stretched to approximately three times their length and placed across the long axis of the coverslip. A conventional glass slide was placed on and perpendicular to the parafilm strips. Pressure was applied and the chamber was flamed briefly to seal it. Solutions were flowed into the chamber via capillary action.

For actin polymerization experiments, unlabeled and Alexa Fluor 488-labeled rabbit muscle actin (Invitrogen) were dialyzed overnight against G buffer and then centrifuged at 100,000 × g for 2 h. The upper two-thirds of the supernatant was taken, and the actin concentration was measured by absorbance at 290 nm. Ca-ATP-actin was converted to Mg ATP-actin by incubation with a 1/10 volume of 10 mm EGTA/0.2 mm MgCl2 at 23 °C for 2 min. N-ethylmaleimide-inactivated myosin at 0.1 μm in high salt Tris-buffered saline (HS-TBS; 50 mm Tris-Cl, pH 7.6,600 mm NaCl) was flowed into the chamber and incubated for 1 min at room temperature. The chamber was washed with 1% bovine serum albumin in HS-TBS followed by 1% bovine serum albumin in low salt Tris-buffered saline (50 mm Tris-HCl, pH 7.6,50 mm NaCl).

To polymerize actin filaments, Mg-ATP-actin with 10% Alexa-labeled actin was mixed 1:1 with 2× polymerization buffer (100 mm KCl, 0.2 mm MgCl2, 2 mm EGTA, 20 mm imidazole, pH 7.0, 100 mm DTT, 0.4 mm ATP, 30 mm glucose, 2% methylcellulose, 40 μg/ml catalase, 200 μg/ml glucose oxidase) giving a final actin concentration of 2 μm. 0.25 nm spectrin-F actin seeds were added immediately. After 10 min at room temperature, 10 nm CP α1β1 was added. After another 10 min, 10 μl of this mixture was loaded into the flow chamber. The actin filaments were monitored for several min to document the absence of growth, and then various concentrations of PIP2 were added along with 2 μm Mg-ATP actin (30% labeled) in polymerization buffer. Actin filament growth was monitored. The rate of filament growth was measured, as was the percentage of filaments growing over time. The rate of incorporation was converted to subunits/s assuming 370 subunits/μm (24).

Molecular Simulations

The starting point for our computational work was a molecular dynamics simulation of CP (Protein Data Bank code 1IZN) performed using NAMD (25). We used the CHARMM27 force field, TIP3P waters, particle mesh Ewald, Berendsen temperature, and pressure coupling (NPT ensemble) and 2-fs time steps. The heating was performed in 50,000 steps with α-carbon constraints, equilibrated at 300,000 without constraints, followed by a 5-ns production trajectory. Five representative structures of CP were extracted from the simulation at 800-ps intervals. Because the macromolecule is held rigid in molecular docking simulations, using multiple protein snapshots allowed us to capture some degree of side chain and backbone flexibility, resulting in significantly improved docking results (26). The molecular docking was carried out using AutoDock 3.0 (27). As PIP2 is somewhat cumbersome to deal with as a full molecule, we used a modified form where the alkyl tails were truncated, leaving just the inositol and diacylglycerol groups (supplemental Fig. 1). This truncated version of PIP2 was minimized using the Tripos force field, and Gasteiger-Marsili charges were assigned, all using Sybyl 6.8 (Tripos Inc., St. Louis, MO). Docking was performed using the Lamarckian genetic algorithm in AutoDock. Initial docking runs using the entire CP structure showed a preference for the region around the base of the α-tentacle. Based on these results, we constructed 0.2-Å spaced grids centered on the α-tentacle. 20 docking runs were completed for each of the five CP structures resulting in a total of 100 predictions. These results were clustered with a 3-Å cutoff, resulting in single, top-predicted structure that was used in subsequent analysis.

RESULTS

Structural Features of PIP2 Important for Interaction with CP

We asked what features of PIP2 were important for its interaction with CP. To measure these interactions, we tested the ability of PIP2 and related compounds to inhibit CP in an actin polymerization capping assay and to quench the intrinsic tryptophan fluorescence of CP. In previous studies with micelles, the degree of phosphorylation of the inositol group was found to be important, as was the anionic character of the head group in general (28).

We reasoned that the micellar structure of PIP2 in solution might be important for its ability to bind CP, because of the clustering of anionic head groups on the micelle surface. This hypothesis is supported by previous observations that dilution of PIP2 with Triton X-100, which forms small micelles, abolishes the ability of PIP2 to inhibit CP but that dilution with liposome-forming lipids does not (28). To test this idea further, we used synthetic versions of PIP2 with shorter acyl chains. The diC4, diC8, and diC16 synthetic forms of PIP2 have 4, 8, and 16 carbons per acyl chain, respectively, whereas PIP2 purified from natural sources has one C18 and one C20 chain. Purified PIP2 at concentrations up to 5 μm completely inhibited the capping activity of 4 nmCP (Fig. 1A). The curves were fit well by a simple model of CP binding to PIP2 (Fig. 1A, gray lines), with a Kd of 0.33 ± 0.04 μm. diC4-PIP2 showed no effect on the actin capping activity of CP (Fig. 1B), and diC8-PIP2 inhibited CP partially, with an apparent Kd of 68 μm (Fig. 1C). These concentrations are below the expected CMC (critical micellar concentration) values for diC4- and diC8-PIP2 (29, 30). diC16-PIP2 inhibited CP completely with a Kd of ~ 0.3 μm (Fig. 1D), comparable with the value for PIP2, which is less than the expected CMC value.

FIGURE 1
Assay of PIP2 acyl chain variants for the ability to inhibit the capping activity of CP

To test physical binding, we used quenching of intrinsic tryptophan fluorescence (10). PIP2 quenched the intrinsic tryptophan fluorescence of CP, with no change in the maximum emission wavelength. The effect was saturable at high PIP2 concentration (Fig. 2A). A simple 1:1 binding model fit the data well (Fig. 2A, gray lines), with a Kd of 5.2 ± 0.2 μm, similar to a previous value for mouse CP in this type of assay (10). diC4-PIP2 and diC8-PIP2 had little effect, whereas diC16-PIP2 produced saturable quenching with a Kd of 7 μm (Fig. 2B), close to the value for purified PIP2. These result support the notion that the ability of PIP2 to assemble in micelles is important for binding and inhibition of CP, presumably because of multimerization of the anionic head groups.

FIGURE 2
PIP2 interaction with CP by quenching of intrinsic tryptophan fluorescence

Next, we tested the head group alone, by using IP3 (inositol-1,4,5 triphosphate). In an actin polymerization seeded growth assay, IP3 at 100 μm had no effect on the capping activity of 4 nm CP (data not shown). In a tryptophan fluorescence titration, addition of IP3 to concentrations >100 μm had no effect on the fluorescence of CP (Fig. 2B). As part of this experiment, we tested diacylglycerol, to represent the lipid portion of PIP2. Again, no effect was seen (Fig. 2B). Thus, neither the head group nor the lipid backbone of the PIP2 molecule is sufficient to bind CP, even at high concentrations.

Computational Docking Analysis of PIP2/CP Interaction

To predict potential sites of interaction between PIP2 and CP, we used a computational molecular docking approach. We began with a collection of CP structures produced by molecular dynamics simulation. The use of multiple structures for CP allowed us to sample different side chain and backbone conformations, and because we performed flexible docking where the rotatable bonds in PIP2 were allowed to move freely, this approach captured some degree of flexibility for both molecules. The molecular docking studies resulted in a clear single prediction for the structure of the complex of PIP2 and CP. As one may have anticipated based on the anionic character of PIP2, we saw interaction primarily with a group of basic residues on the surface of CP. This group included Lys-256 and Arg-260 of the α-subunit and Arg-225 of the β-subunit (Fig. 3, A and B). Given the residues and charged groups involved, the interactions are predominantly electrostatic in nature, with the addition of several hydrogen bonds (Fig. 3B). The inositol ring sits in a pocket formed by these residues, between the two subunits, and the location and orientation of these basic side chains suggests some degree of specificity for PI(4,5)P2 versus other phosphoinositides, as observed previously (28).

FIGURE 3
Predictions for the binding site of PIP2 on CP

In terms of the primary amino acid sequence, Lys-256 and Arg-260 are two residues in a conserved cluster of multiple basic residues near the C terminus of the CP α-subunit (Fig. 3C). Among these conserved basic residues, the CP crystal structure shows that Lys-256, Arg-260, Arg-266, and Lys-268 are on the surface and exposed to solvent (Fig. 3A). In contrast, Arg-259 is conserved but buried, where it forms an ionic bond with the conserved acidic residue Glu-221 of the β-subunit. Residue Arg-225 of CP β is also highly conserved in sequence alignments and exposed to solvent in the crystal structure. The C-terminal region of the α-subunit has been implicated in binding to actin (17), so the presence of a PIP2 micelle at this site would be expected to provide an effective steric block to actin binding, as seen for villin, for example (31).

Mutation of the Conserved Basic Residues

To test for involvement of this basic patch in binding PIP2, we introduced point and truncation mutations into the region. First, we tested binding with tryptophan quenching as shown in Fig. 2 and Table 1. For the residues implicated by the computational docking analysis, the single mutants CP α1(K256A)β1, CP α1(R260A)β1 and CP α1β1(R225A) bound PIP2 normally. The double mutant CP α1(R256A,K260A) β1 bound more weakly, with a Kd of 12 μm, compared with 5 μm for wild-type CP. The triple mutant CP α1(R256A,K260A)β1(R225A) had a higher Kd of 19 μm.

TABLE 1
Binding constants for the interaction of CP mutants with PIP2 from tryptophan fluorescence quenching

The α-subunit double mutant KR266,268AA bound to PIP2 with normal affinity (Table 1). Truncation of 28 residues from the C terminus of the α-subunit removes the final helix, including Arg-260, Lys-266, and Arg-268, and leaves Leu-258 as the C-terminal residue. This mutant is known to bind actin poorly (17). The affinity of the CP α Δ28 truncated protein for PIP2 was decreased, with a Kd of 17 μm (Fig. 2, Table 1).

The C-terminal region of the β-subunit of CP also appears to function as a binding site for actin, independently of the α C terminus (17). Truncation of the β-subunit by 34 residues had no effect on the ability of CP to bind PIP2 in the tryptophan quenching assay (Fig. 2, Table 1).

We next tested all of these CP mutants for interaction with PIP2 based on inhibition of capping activity in actin polymerization assays. As part of this analysis, we first determined the affinity of the CP mutants for the F-actin barbed end, i.e.Kcap.” These values are required in order to calculate the binding constant for the CP/PIP2 interaction. Wild-type CP (α1β1) inhibited actin polymerization at barbed ends of actin filaments created by spectrin-actin seeds in a dose-dependent manner (Fig. 4A, black lines). Kinetic modeling gave good fits (Fig. 4A, gray lines), with a Kcap of 0.1 ± 0.05 nm (Table 2), similar to previous results (17). The α-subunit single and double mutations K256A,R260A, K266A,R268A, and R256A,K260A produced small to moderate increases in Kcap, with values of 0.18, 0.36, 0.4, and 1.3 nm, respectively (Fig. 4, Table 2). The β-subunit single mutant R225A also showed a moderate increase in Kcap to 1.2 nm. The triple mutant CPα(R256A,K260A)β(R225A) capped actin weakly (Fig. 4F), with a Kcap of 45 nm, which was mainly because of an increase in the off-rate constant (Table 2). The α-subunit truncation mutant CPα(Δ28)β capped actin filaments weakly, with a Kcap of 1570 nm, and the β-subunit truncation mutant CPαβ(Δ34) capped actin with a Kcap of 10.5 nm (Table 2), consistent with previous studies (17).

FIGURE 4
Actin capping assays with CP and mutants
TABLE 2
Binding constants for the interaction of CP with actin

With values for Kcap in hand, we then repeated the assays with increasing concentrations of PIP2 and used kinetic modeling to fit the curves and determine Kd values for the interaction of the CP mutants with PIP2 (Fig. 5, Table 3). The α-subunit single and double mutations K256A, R260A, K266A,R268A, and R256A,K260A produced small to moderate increases in the Kd, with values of 0.31, 0.36, 0.41 and 3.9 μm, respectively (Table 3). The β-subunit single mutant R225A showed no increase in Kd. The triple mutant CPα(R256A,K260A)β(R225A) gave a ~60-fold increase in Kd to 21 μm (Fig. 5F and Table 3). For the α-subunit truncation mutant CPα(CΔ28)β, the Kd was 42 μm, ~130-fold greater than the value for wild-type CP (Fig. 5A, Table 3). The β-subunit truncation mutant CPαβ(Δ34) had a Kd of 1.6 μm (Fig. 5B, gray lines, and Table 3).

FIGURE 5
PIP2 binding affinities of CP mutants
TABLE 3
Binding constants for the interaction of CP with PIP2 based on seeded actin polymerization assays

Nitrate Ions in the Crystal Structure

Nitrate ions can occupy phosphate-binding sites, and CP was crystallized in the presence of nitrate (14). The crystal structure has two nitrate ions in contact with the surface of the β-subunit. One interacts with Lys-95, and the other interacts with the helix dipole of helix 5. We mutated Lys-95 to alanine to make CP αβ(K95A). We found that this mutant capped actin with a normal affinity (Table 2), and its affinity for PIP2 was the same as that of wild-type CP in both the actin capping and tryptophan fluorescence assays (Tables 1 and and33).

Direct Observation of Uncapping by PIP2

One prediction of the wobble model is that an inhibitor of capping that does not bind to the CP β-subunit tentacle, about which the wobbling is proposed to occur, should be able to uncap. Indeed, previous studies showed that addition of PIP2 to capped actin filaments in a polymerization reaction leads to an increase in the polymerization rate consistent with complete and rapid uncapping (11). Here, we tested uncapping with a more direct assay in which individual actin filaments are visualized by TIRF microscopy. In these experiments, movies reveal actin filaments growing from their ends. When CP was added, the growth stopped. Subsequent addition of PIP2 caused the ends to return to the growing state (Fig. 6A and supplemental Movie 1). The mean rate of growth was 3.4 ± 0.3 subunits/s before capping and 2.8 ± 0.4 subunits/s after uncapping, in the presence of 2.0 μm monomeric actin. The effect of PIP2 was nearly complete and relatively rapid, which is comparable with previous results with assays in solution (11). The fraction of ends converted from the capped to the growing state increased with PIP2 concentration (Fig. 6B). The concentration of PIP2 producing a half-maximal effect in this assay was ~100 μm, substantially higher than those seen with the tryptophan fluorescence and actin capping assays above. This may reflect sequestration of PIP2 by the relatively large surface of bovine serum albumin-coated glass in this assay or perhaps a decreased accessibility of the actin filaments because of tethering at the surface.

FIGURE 6
Uncapping of capped actin filaments by PIP2

DISCUSSION

In this study we performed a structure/function analysis to identify regions of CP necessary for interaction with PIP2. The analysis was directed by predictions from computational modeling of CP/PIP2 interactions, and the computational results were consistent with the functional and physical assays of the interaction. In addition, direct visualization of actin filament growth by TIRF microscopy provided new direct evidence of uncapping. Together, the results support the wobble model for the interaction of CP with the barbed end of the actin filament.

Nature of the Interaction between CP and PIP2

We sought evidence of what regions of CP were needed for its interaction with PIP2 by site-directed mutagenesis, changing single amino acids and truncating the C termini of the two subunits of the heterodimer. In previous studies, clusters of basic residues on the surface of proteins were implicated in binding PIP2. For example, crystal structures of an AP180 protein complexed with phosphatidylinositols reveal a surface-exposed binding site composed of three lysine residues and one histidine, which contact the phosphate groups (32). Sequence analysis of related proteins, combined with mutagenesis, identified a consensus binding sequence of KX9KX(K/R)(H/Y). PIP2 binds and inhibits a number of actin-binding proteins, including profilin and the gelsolin family, and a number of studies identify (K/R)X3–5(K/R)X(K/R)(K/R) as a consensus binding site (31, 33, 34).

In addition, we know that CP from every organism tested, including yeast, plants and mammals, is inhibited by PIP2 (10). Therefore, we focused on conserved clusters of basic residues on the surface of the protein. The largest and most distinct cluster is in and around the C-terminal region of the α-subunit, including its connection to the body of the protein. A recent study of phosphoinositides interacting with Arabidopsis CP also noted the existence of this conserved basic region and suggested that it might be important (10). Indeed, we found that a cluster of three conserved basic residues on the surface, near the α C terminus, were important for binding PIP2. These include Lys-256 and Arg-260 of the α-subunit and Arg-225 of the β-subunit. In contrast, two nearby basic residues, also conserved and on the surface, were not important.

Computational docking studies provide complementary evidence for the primary role of these three residues. Of note, the docking studies were done largely in advance of the mutagenesis studies, and they helped us to narrow and define our list of potential mutants. A recent computational study of CP interacting with lipids, including ones other than polyphosphoinositides, raises the possibility that portions of CP may become partially buried in the lipid bilayer (35). One of several regions identified as potentially important in that study was the 215–232 region of the β-subunit, which includes Arg-225, one of the three residues identified as important in our results here. The notion of lipid binding to the interior of CP may be supported by an early study in which, by gel filtration analysis, micelles of PIP2 appeared to partially dissociate the α- and β-subunits (28).

Implications for How CP Binds Actin

As part of our analysis, we tested the ability of a number of new point mutants of CP to cap actin filaments. Previous studies, largely with truncations, had implicated the C-terminal region of the α-subunit as important, and these new point mutations help to define the residues important for binding actin more closely. Of note, a cluster of three conserved basic residues on the surface, near the α C terminus, were found to be important, including Lys-256 and Arg-260 of the α-subunit and Arg-225 of the β-subunit. Two other basic conserved surface residues located nearby, Arg-266 and Lys-268 of the α-subunit, were found to be much less important. These results do not show that these residues bind actin directly. This will require further studies, including structural ones.A recent cryo-electron microscopy study indicates that Arg-260, Arg-266, and Lys-268 are among a cluster of basic of residues making close contact with nearby Lys-256 (15).

Implications for the Wobble Hypothesis

As originally suggested by the analysis of the crystal structure of CP (14), we currently hypothesize that CP has two independent sites for binding to the barbed end of the actin filament. One includes the C-terminal “tentacle” of the β-subunit, so-named because it is surrounded by solvent in the crystal structure and is highly mobile in molecular dynamics simulations (6, 14). The other involves the C-terminal region of the α-subunit. However, in this case, the region does not appear to be mobile, based on S100 inhibitor and molecular dynamics studies (6, 14). CP bound to the barbed end of the actin filament appears to utilize both sites (17). We hypothesize that they can dissociate independently, which predicts that CP bound only by its β-tentacle should be relatively mobile and “wobble” on the barbed end of the filament. One prediction of the model is that a molecule that binds to CP at the second actin-binding site, near the α C terminus, should be able to bind to the wobble state and thereby promote dissociation and thus “uncapping.”

Our results here with PIP2 confirm this prediction. First, the point mutations near the α C terminus show that this region is involved in binding actin and PIP2. Second, PIP2 can induce uncapping, as visualized here in real time with TIRF microscopy of polymerizing actin filaments. A complementary prediction and confirmation of the model was provided by our recent results with V-1/myotrophin, which binds to CP and inhibits actin binding but does so by interacting with the β-tentacle (6). In this case, the model predicts that uncapping will not occur, and that was the result.

Implications for Actin Assembly in Cells

CP is an essential element of the dendritic nucleation model, and consistent with that view, loss of CP leads to loss of lamellipodia in cultured cells (3). In this model, barbed ends are created near the plasmamembrane and grow for a time before being capped by CP. One simple hypothesis based on our results is that PIP2 in the plasma membrane, acting as a second messenger, will inhibit capping and thus promote actin polymerization at the membrane. Indeed, overexpression of phosphatidylinositol 5-kinase can lead to rocketing of membrane vesicles in the cell (36, 37).

A more interesting and speculative hypothesis is that PIP2 or other molecules that can cause uncapping, such as the protein CARMIL (9), might be responsible for promoting disassembly of actin filaments in zones of depolymerization or for providing a source of free barbed ends. The latter has been suggested by studies of platelet activation, where explosive actin polymerization occurs (12).

Supplementary Material

Supplemental Material can be found at: http://www.jbc.org/cgi/content/full/M609850200/DC1

Footnotes

*This work was supported in part by National Institutes of Health Grants GM38542 (to J. A. C.) and GM067246 (to D. S.).

SThe on-line version of this article (available at http://www.jbc.org) contains supplemental Tables I and II, Fig. 1, and Movie 1.

5The abbreviations used are: CP, capping protein; PIP2, phosphatidylinositol 4,5-bisphosphate; TIRF, total internal reflection fluorescence; DTT,dithiothreitol.

References

1. Wear MA, Cooper JA. Trends Biochem Sci. 2004;8:418–428. [PubMed]
2. Pollard TD, Borisy GG. Cell. 2003;4:453–465. [PubMed]
3. Mejillano MR, Kojima S, Applewhite DA, Gertler FB, Svitkina TM, Borisy GG. Cell. 2004;3:363–373. [PubMed]
4. Loisel TP, Boujemaa R, Pantaloni D, Carlier MF. Nature. 1999;6753:613–616. [PubMed]
5. Kim K, Galletta BJ, Schmidt KO, Chang FS, Blumer KJ, Cooper JA. Mol Biol Cell. 2006;17:1354–1363. [PMC free article] [PubMed]
6. Bhattacharya N, Ghosh S, Sept D, Cooper JA. J Biol Chem. 2006;281:31021–31030. [PMC free article] [PubMed]
7. Bruck S, Huber TB, Ingham RJ, Kim K, Niederstrasser H, Allen PM, Pawson T, Cooper JA, Shaw AS. J Biol Chem. 2006;281:19196–19203. [PMC free article] [PubMed]
8. Canton DA, Olsten ME, Niederstrasser H, Cooper JA, Litchfield DW. J Biol Chem. 2006;281:36347–36359. [PMC free article] [PubMed]
9. Yang C, Pring M, Wear MA, Huang M, Cooper JA, Svitkina TM, Zigmond SH. Dev Cell. 2005;2:209–221. [PMC free article] [PubMed]
10. Huang S, Gao L, Blanchoin L, Staiger CJ. Mol Biol Cell. 2006;17:1946–1958. [PMC free article] [PubMed]
11. Schafer DA, Jennings PB, Cooper JA. J Cell Biol. 1996;1:169–179. [PMC free article] [PubMed]
12. Barkalow K, Witke W, Kwiatkowski DJ, Hartwig JH. J Cell Biol. 1996;2:389–399. [PMC free article] [PubMed]
13. Eddy RJ, Han J, Condeelis JS. J Cell Biol. 1997;139:1243–1253. [PMC free article] [PubMed]
14. Yamashita A, Maéda K, Maéda Y. EMBO J. 2003;7:1529–1538. [PMC free article] [PubMed]
15. Narita A, Takeda S, Yamashita A, Maeda Y. EMBO J. 2006;25:5626–5633. [PMC free article] [PubMed]
16. Kim K, Yamashita A, Wear MA, Maeda Y, Cooper JA. J Cell Biol. 2004;4:567–580. [PMC free article] [PubMed]
17. Wear MA, Yamashita A, Kim K, Maéda Y, Cooper JA. Curr Biol. 2003;13:1531–1537. [PubMed]
18. Soeno Y, Abe H, Kimura S, Maruyama K, Obinata T. J Muscle Res Cell Motil. 1998;6:639–646. [PubMed]
19. DiNubile MJ, Cassimeris L, Joyce M, Zigmond SH. Mol Biol Cell. 1995;12:1659–1671. [PMC free article] [PubMed]
20. Pollard TD. J Cell Biol. 1986;103:2747–2754. [PMC free article] [PubMed]
21. Wear MA, Cooper JA. J Biol Chem. 2004;279:14382–14390. [PMC free article] [PubMed]
22. Kuhn JR, Pollard TD. Biophys J. 2005;88:1387–1402. [PMC free article] [PubMed]
23. Kron SJ, Spudich JA. Proc Natl Acad Sci U S A. 1986;83:6272–6276. [PMC free article] [PubMed]
24. Huxley HE, Brown W. J Mol Biol. 1967;30:383–434. [PubMed]
25. Kale L, Skeel R, Bhandarkar M, Brunner R, Gursoy A, Krawetz N, Phillips J, Shinozaki A, Varadarajan K, Schulten K. J Comput Phys. 1999;151:283–312.
26. Mitra A, Sept D. Biochemistry. 2004;43:13955–13962. [PubMed]
27. Morris GM, Goodsell DS, Halliday RS, Huey R, Hart WE, Belew RK, Olson AJ. J Comput Chem. 1998;19:1639–1662.
28. Heiss SG, Cooper JA. Biochemistry. 1991;30:8753–8758. [PubMed]
29. Campbell RB, Liu F, Ross AH. J Biol Chem. 2003;278:33617–33620. [PubMed]
30. Rebecchi MJ, Eberhardt R, Delaney T, Ali S, Bittman R. J Biol Chem. 1993;268:1735–1741. [PubMed]
31. Kumar N, Zhao P, Tomar A, Galea CA, Khurana S. J Biol Chem. 2004;279:3096–3110. [PubMed]
32. Ford MG, Pearse BM, Higgins MK, Vallis Y, Owen DJ, Gibson A, Hopkins CR, Evans PR, McMahon HT. Science. 2001;291:1051–1105. [PubMed]
33. Yu F-X, Sun H-Q, Janmey PA, Yin HL. J Biol Chem. 1992;267:14616–14621. [PubMed]
34. Janmey PA, Xian W, Flanagan LA. Chem Phys Lipids. 1999;101:93–107. [PubMed]
35. Smith J, Diez G, Klemm AH, Schewkunow V, Goldmann WH. Theor Biol Med Model. 2006;3:30. [PMC free article] [PubMed]
36. Rozelle AL, Machesky LM, Yamamoto M, Driessens MH, Insall RH, Roth MG, Luby-Phelps K, Marriott G, Hall A, Yin HL. Curr Biol. 2000;6:311–320. [PubMed]
37. Schafer DA, Welch MD, Machesky LM, Bridgman PC, Meyer SM, Cooper JA. J Cell Biol. 1998;7:1919–1930. [PMC free article] [PubMed]
38. Humphrey W, Dalke A, Schulten K. J Mol Graph. 1996;14:33–8. 27–28. [PubMed]
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