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* Muscle Research Unit, Institute for Biomedical Research, University of Sydney, New South Wales, Australia; Belozersky Institute of Physico-Chemical Biology, Lomonosov Moscow State University, Moscow, Russia; Pennsylvania Muscle Institute and Department of Physiology, University of Pennsylvania, School of Medicine, Philadelphia, Pennsylvania; and Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, Connecticut
Correspondence: Address reprint requests to Enrique M. De La Cruz, Yale University, Dept. of Molecular Biophysics and Biochemistry, PO Box 208114, New Haven, CT 06520-8114. Tel.: 203-432-5424; Fax 203-432-1296; E-mail: enrique.delacruz{at}yale.edu.
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ABSTRACT |
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ATP) and Lys-61 by 1.9 Å, whereas the distance between Cys-374 and Lys-61 is minimally affected. Distance determinations are consistent with tß4 binding being coupled to a rotation of subdomain 2. By differential scanning calorimetry, tß4 binding increases the cooperativity of ATP-actin monomer denaturation, consistent with conformational rearrangements in the tß4-actin complex. Changes in fluorescence resonance energy transfer are accompanied by marked reduction in solvent accessibility of the probe at Gln-41, suggesting it forms part of the binding interface. Tß4 and cofilin compete for actin binding. Tß4 concentrations that dissociate cofilin from actin do not dissociate the cofilin-DNase I-actin ternary complex, consistent with the DNase binding loop contributing to high-affinity tß4-binding. Our results favor a model where thymosin binding changes the average orientation of actin subdomain 2. The tß4-induced conformational change presumably accounts for the reduced rate of amide hydrogen exchange from actin monomers and may contribute to nucleotide-dependent, high affinity binding. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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). Thymosin ß4 (tß4) is a small (Mr < 5 kDa) member of a highly conserved family of actin monomer-sequestering proteins present in actively motile cells, developing chick brain, and embryonic skeletal muscle (1,2). Tß4 forms a 1:1 complex with monomeric actin and can bind to F-actin at high concentrations (3,4). ß-thymosins are the principal in vivo regulators of unpolymerized actin, and are essential for maintaining the cytoplasmic pool of actin monomers required for rapid filament elongation (reviewed in (1)).
The exact tß4 binding surfaces on the actin monomer are unknown and several models have been proposed (4–7). There is a general agreement that subdomains 1 and 3 (SD1 and SD3) of actin form part of the interface, similar to gelsolin and profilin, accounting for competitive binding (8,9). Binding to SD1 and SD3 is expected to restrict propeller-type opening and closing motion of the two major domains (10). However, there is disagreement over the role of SD2 of actin in binding tß4. It has been proposed that the C-terminal region of tß4 binds directly to the DNase I-binding loop (5) because it can be chemically crosslinked to residues of the DNase I-binding loop in SD2 (5,8). The nucleotide-dependence of SD2 conformation (11) could therefore account for the nucleotide-dependence of tß4 binding affinity (12). Subtilisin cleavage of actin in the DNase I-binding loop causes a twofold decrease in the affinity of tß4 (13), consistent with SD2 contributing to the stability and strength of the actin-tß4 interaction. Image reconstruction of actin filaments chemically crosslinked to tß4 (4) suggests tß4 binds SD2 and SD4 as well as the gap between SD1 and SD2 (5). Binding of tß4 to actin monomers is coupled to a change in heat capacity, dissociation of bound waters, a reduction in amide hydrogen exchange (5), and inhibition of nucleotide exchange (3). It was hypothesized that tß4 folding upon binding could account for the change in heat capacity (5), and recent evidence favors this interpretation (13). It was also proposed that tß4 binding inhibits the movement and separation of SD2 and SD4, closing the nucleotide-binding cleft and reducing the rates of nucleotide and amide hydrogen exchange (5). In this report, we used fluorescence spectroscopy to test the hypothesis that tß4 changes the actin monomer conformation (5) and evaluate the contributions of actin SD2 to tß4 binding. Our results are consistent with tß4 binding to SD1, SD2 and SD3, causing changes in the spatial orientation of actin monomer subdomains.
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MATERIALS AND METHODS |
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ATP), dansyl cadaverine (DC), and fluorescein-5-isothiocyanate (FITC) were purchased from Molecular Probes (Eugene, OR). N-(4-(dimethylamino)-3,5-dinitrophenyl)-maleimide (DDPM) and N-ethylmaleimide (NEM) were from Sigma-Aldrich (St. Louis, MO). Rabbit skeletal muscle actin was prepared in G buffer (5 mM imidazole, pH 7.0, 0.2 mM ATP, 0.1 mM CaCl2, 0.5 mM DTT, and 1 mM NaN3). Tß4 was isolated from bovine spleen (8). Bovine pancreatic DNase I (DPRF grade) was from Worthington Biochemicals (Lakewood, NJ). Cofilin was prepared as a recombinant protein using a cDNA sequence derived from chick embryo, kindly supplied by Dr. Takashi Obinata (Chiba, Japan). Concentrations of actin, DNase I, and cofilin were determined using extinction coefficients of 0.63 cm–1 (0.1%, 290 nm), 1.1 cm–1 (0.1%, 280 nm), and 0.98 cm–1 (0.1%, 280 nm), respectively. Actin was labeled with the following fluorescent probes as described: a), DC (14); b), DC-actin further modified with DDPM (14); c), IAEDANS (15); d), NEM or FITC (16); and e), -ATP (17). The following molar extinction coefficients were used for probes bound to actin: DC, 4640 M–1 cm–1 at 330 nm and DDPM, 3050 M–1 cm–1 at 440 nm (14); IAEDANS, 6100 M–1 cm–1 at 336 nm (15); and FITC, 74,500 M–1 cm–1 at 493 nm (18). The concentration of actin in labeled samples was determined by the DC Protein Assay (
= 795 nm, Bio-Rad Laboratories, Hercules, CA) using unlabeled actin as the standard. The typical labeling ratios were: 0.63–0.9 for DC (n = 10); 0.5–0.85 for DDPM (n = 7); 0.8–0.98 for IAEDANS (n = 10); and 0.85–1.0 for FITC (n = 9). Bound ATP was exchanged with ATP as described earlier (17). In brief, nonlabeled or FITC-labeled G-actin (24–48 µM) was treated with 0.1 volumes of Dowex-1 on ice for 10 min to remove free ATP. Dowex-1 was removed by filtration and the solution was supplemented with 0.5 mM ATP, 0.1 mM CaCl2, and 1 mM Tris/HCl (pH 8.0) and equilibrated on ice overnight. This procedure was repeated again to ensure a complete exchange of bound ATP with ATP. Free nucleotide was removed with Dowex-1 immediately before fluorescence experiments. Because the actin concentrations used are much higher than the Kd for nucleotide binding (19), bound nucleotide will not dissociate to any appreciable extent during the time-course of the experiments.
Fluorescence measurements were carried out in G buffer at 10°C in a temperature-controlled cuvette using an SLM 48000 S Multiple Frequency Lifetime Spectrofluorometer (SLM Aminco, Foster City, CA) operating on a xenon arc lamp essentially as described (14). All experiments were repeated 5–10 times and results were presented as means ± SE. Fluorescence resonance energy transfer (FRET) efficiency (E) was determined from the intensities of the donor (D) in the presence (FDA) and absence (FD) of the acceptor (A) according to Eq. 1: E = (1 – FDA/FD)/
, where is a degree of labeling with the acceptor in the double-labeled actin. Intensities were normalized to the D-concentration after background subtraction. Energy transfer efficiency is related to distance separating D- and A-probes according to Eq. 2: 
Ro is the Förster critical distance between the D and A (when E = 0.5) defined by Eq. 3: 
where n is the refractive index of the medium, 
2 is the orientation factor, QD is the quantum yield of the D on actin in the absence of the A, and J is the overlap integral given in M–1 cm–1 nm4. Ro distances of actin alone were taken as 49.6 Å for IAEDANS/FITC (20) and 47.4 Å for ATP/FITC (17). These values were corrected for changes in the donor quantum yield (21) due to tß4 binding. The Ro for DC/DDPM was obtained experimentally using fluorescence emission spectrum of the DC-actin in the presence of nonfluorescent DDPM and the absorption spectrum of the DDPM-actin, taking 2 = 2/3. FRET measurements between the following positions were done using the following donor-acceptor pairs as described: a), DC (Gln-41) and DDPM (Cys-374) (14); b), IAEDANS (Cys-374) and FITC (Lys-61) (17); and c), ATP (nucleotide-binding cleft) and FITC (Lys-61) (17). Intensities of the probe in the D-only and D/A samples were normalized to the fluorophore concentration by overnight tryptic digestion, as described (17). In brief, 0.2–0.4 mg ml–1 of trypsin in G-buffer was added into the D and D/A sample solutions after FRET measurements. After digestion, the amino-acid residues with D- and A-probes were separated, abolishing FRET. Thus, fluorescence intensities were directly proportional to the concentration of the D-probe in solution. The ratio of these fluorescence intensities was taken to be proportional to FDA x Ci/FD x C'i (Eq. 4), where Ci and C'i are the concentrations of D in D and D/A samples, respectively. Acrylamide quenching of DC-actin fluorescence (ex = 332 nm, em = 512 nm) was performed at 22°C in a thermostated cuvette as described (14). The apparent Stern-Volmer constant (KSV) was obtained from the slope of a plot of Fo/F versus [acrylamide] by fitting to Eq. 5 as Fo/F = 1 + KSV [Q], where Q is the quencher, i.e., acrylamide.
SDS PAGE and native 10% PAGE gels were done following standard protocols (22). After FRET measurements, labeled actin and its complexes were mixed with equal volumes of native sample buffer (62.5 mM Tris-Cl, pH 6.8, 10% glycerol, and 0.1% bromophenol blue) and separated by native PAGE gel electrophoresis at 70–110 V for 90–120 min in an ice bath (23). Differential scanning calorimetry (DSC) was performed on a DASM-4M differential scanning microcalorimeter (Institute for Biological Instrumentation, Pushchino, Russia) as described (24). The scanning rate was 1 K min–1. Transition temperatures (Tm) were determined from the peak of thermal transition, and the apparent enthalpies (
H°app) were calculated from the integrated areas of the denaturation peaks.
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RESULTS |
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). However, in the presence of DDPM this decrease is abolished (dashed curve 2'). The resulting FRET efficiency calculated using R0 = 29.4 Å (corrected for changes in the QD) is 0.182 ± 0.03, which corresponds to a distance of 37.8 ± 1.2 Å between DC and DDPM in the actin-tß4 complex. Thus, tß4 increases the distance between the probes attached to Cys-374 and Gln-41 by 2 Å (p < 0.000). Since DDPM modification of Cys-374 induces an allosteric effect on the actin monomer conformation, we modified DC-actin with NEM at Cys-374, and repeated the FRET experiments using DC/NEM-actin as a donor-only sample. The NEM probe has no absorbance in the overlap region of the donor emission, and therefore, there is no FRET between these two probes. Fluorescence intensity of DC slightly increases in the presence of NEM (dash-dot curve 3, n = 5). However, it has no significant effect on the final distance. Using R0 = 31.05 Å (recalculated for changes in the QD of DC in the presence of NEM), and the efficiency of 0.307 ± 0.03 (n = 5), the calculated distance between the probes attached to Gln-41 and Cys-374 is 35.5 ± 0.9 Å. The addition of tß4 to DC/NEM-actin (dash-dot curve 3') results in a decrease in fluorescence intensity by 13 ± 4%, i.e., similar to NEM-unmodified DC-actin. The calculated efficiency of 0.191 ± 0.02 (n = 5) yields a distance of 37.4 ± 1.0 Å between DC and DDPM in the presence of tß4. Thus, tß4 increases the measured distance by 
2 Å. Native PAGE gels (Fig. 2 B) demonstrate that concentrations of tß4 used in FRET experiments are sufficient to saturate labeled actin as expected from the equilibrium binding affinities (5). The tß4-induced increase in the Gln-41/Cys-374 distance is accompanied by a reduction in the solvent accessibility of Gln-41 (Fig. 2 C). Acrylamide quenches the fluorescence emission of DC in the G-actin/tß4 complex (curve GT, KSV = 1.0 ± 0.3, n = 6) less efficiently than in G-actin alone (curve G, KSV = 2.4 ± 0.1, n = 6). A similar effect is observed in F-actin (curve F, KSV = 1.3 ± 0.1, n = 5) and in the actin-DNase I complex (curve GD, KSV = 1.2 ± 0.1, n = 6) where Gln-41 is protected from solvent because it forms part of the binding interface (26).
Effect of tß4 on the distance between Lys-61 and Cys-374 of actin
Actin labeled with an IAEDANS (donor) at Cys-374 and a FITC (acceptor) at Lys-61 (Fig. 1) has a FRET efficiency of 0.65 ± 0.04 corresponding to a distance of 44.7 ± 1.1 Å, in agreement with a previous report (20). The fluorescence emission spectrum of IAEDANS-actin (Fig. 3 A, solid curve 1) demonstrates that tß4 binding (dashed curve 1') generates a 39 ± 3% (n = 10) decrease in fluorescence intensity at 475 nm and an 11-nm red shift of the emission maximum wavelength (5). However, in the presence of FITC (solid curve 2), tß4 causes only a 27 ± 4% decrease ( = 475 nm, n = 9) of the D-fluorescence intensity (dashed curve 2'). The difference demonstrates that there is a reduction in FRET efficiency between the probes at Cys-374 and Lys-61 when tß4 is bound. The ratios of corrected fluorescence intensities (FDA to FD at = 475 nm) of the IAEDANS-actin/tß4 (dashed curve 1') and IAEDANS/FITC-actin/tß4 complexes (dashed curve 2') yields a FRET efficiency of 0.59 ± 0.04 (n = 9). QD of the D significantly decreases in the presence of tß4 (0.48 in actin alone compared to 0.3 in the actin/tß4 complex, n = 6). This reduces R0 from 49.6 Å (in actin alone) to 46.5 Å (in the actin/tß4 complex), yielding a distance between IAEDANS and FITC in the actin/tß4 complex of 43.8 ± 0.9 Å, a mean reduction of only 0.9 Å (p = 0.018).
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). DNase I inhibits the tß4-induced spectroscopic changes (Fig. 3) because it prevents tß4 from binding to actin as assayed by native gel electrophoresis (Fig. 3 D), consistent with common or partially overlapping binding sites (8).
Effect of tß4 on the distance between bound -ATP and Lys-61 of actin
FRET efficiencies between ATP and FITC were calculated from comparison of normalized fluorescence intensities of ATP/FITC- and ATP-actin in the absence and presence of tß4. The distance between ATP and FITC attached to Lys-61 of actin (in the absence of tß4 or DNase I) is 38.4 ± 0.8 Å, the R0 is 47.4 Å, and the FRET efficiency is 0.78 ± 0.02 (17). Binding of tß4 results in a 14 ± 3% (n = 6) increase in the ATP-actin fluorescence (Fig. 4), which is almost completely eliminated when tß4 binds to ATP/FITC-actin, indicating that tß4 binding increases the FRET efficiency between bound ATP and Lys-61. The precise value of the change in FRET efficiency was obtained by incubating samples with 0.4 mg/ml of trypsin in the presence of 2 mM ATP, which results in separation of the D- and A-probes and allows normalization of the reference fluorescence with respect to the ATP concentration (17). Unlabeled ATP readily competes with ATP bound to actin. The comparison of fluorescence intensities of digested -ATP/FITC- and ATP-actin is used for final normalization of FRET efficiency (see Methods). In the presence of tß4, the normalized ratio of the fluorescence intensities of ATP/FITC-actin to that of ATP-actin is 0.16 ± 0.02, i.e., a FRET efficiency of 0.84 ± 0.02. Using an R0 value of 48.1 Å (corrected for the tß4-induced increased QD), the distance between the probes in the actin/tß4 complex is 36.5 ± 1.0 Å. Thus, tß4 closes the gap between the FITC probe on Lys-61 and ATP by 1.9 Å (p < 0.000).
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H°app) of 550 ± 20 kJ/mol (n = 5). Tß4 alone has no thermal transition (i.e., it is undistinguishable from the buffer baseline, data not shown), consistent with a disordered solution structure in the absence of actin (13). The thermal transition of actin in the presence of tß4 occurs at 61.7°C, and the H°app increases to 620 ± 25 kJ/mol (n = 5). The width at the half-height of the thermal denaturation peak, T0.5, is equal to 6.5°C for the actin/tß4 complex but it is 7.5°C for actin alone, suggesting a more cooperative thermal transition with tß4 binding. The thermal denaturation of actin in the presence and absence of tß4 was irreversible (data not shown). It is important to clarify that for calorimetric enthalpies of thermal transitions to be thermodynamically significant, denaturation must be reversible. Since actin denaturation is an irreversible process, the measured calorimetric enthalpies are best considered to be apparent thermodynamic parameters reflecting the nonequilibrium denaturation of actin (24).
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,23), which has been employed previously to show an increased affinity of cofilin for actin in the presence of DNase I (23). The actin/tß4 complex migrates as a distinct band ahead of the actin monomers with free tß4 appearing below the actin/tß4 band (Fig. 6 A). Cofilin addition (molar ratios of A/T/C = 1:5:2) results in the appearance of a new band corresponding to the actin/cofilin complex. Excess cofilin (A/T/C molar ratios of 1:5:7) competes and displaces tß4 from the actin/tß4 complex. Cofilin and tß4 do not bind each other (lane 1, Fig. 6 A). The higher affinity of cofilin for actin monomers (1) accounts for the concentration-dependence of the competition. As expected for a reversible equilibrium reaction, cofilin and tß4 competitions are independent of the order of addition (Fig. 6 B). However, tß4 does not compete with cofilin for actin binding in the presence of DNase I (Fig. 6 C), and the cofilin-DNase I-actin ternary complex (23) is stable even after the addition of a 15-fold molar excess tß4 (Fig. 6 C). However, the addition of the same excess tß4 to the actin/cofilin complex in the absence of DNase I leads to the dissociation of the actin/cofilin complex (Fig. 6 D). Importantly, under experimental conditions, binding of tß4 to actin is completely abolished in the presence of DNase I, irrespective of the presence of cofilin (Fig. 3, D and E). We interpret these results to mean that accessibility to the DNase I binding loop is necessary for tß4 to compete with cofilin for actin binding, presumably because SD2 forms part of the tß4 binding site on actin and contributes to the high affinity.
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DISCUSSION |
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). The observation that tß4 competes with cofilin and dissociates the cofilin-actin complex but not the cofilin-actin-DNase I ternary complex (Fig. 6) favors a role for SD2 in high affinity tß4 binding. We conclude that tß4 binds SD2 and rotates the DNase I-binding loop toward the nucleotide-binding site and away from SD1 (Fig. 1), providing a plausible structural mechanism for how tß4 enhances the fluorescence of bound ATP (Fig. 4), changes the hydration (5), and reduces amide hydrogen exchange from actin (5). The nucleotide-dependence of SD2 conformation likely contributes to the nucleotide-dependence of the tß4 binding affinity (5,28). When ATP bound to actin is exchanged for ADP, the DNase I binding loop adopts a folded -helical conformation (11,12,29). Tß4 has a much higher affinity for ATP-actin than ADP-actin monomers (5,28), suggesting that the conformation of SD2 dictates how tightly tß4 binds actin (12) and that the higher affinity of tß4 for ATP-actin results from stabilizing contacts made with SD2 (5,8).
The detailed three-dimensional structure of the actin-tß4 complex has not been determined experimentally. However, two alternatives have been proposed for tß4 binding to actin monomers, based on structures of actin bound to tß4 homologs (6,7). Hertzog et al. (6) reported the structure of actin bound to domain 1 of ciboulot, which shares 30% sequence identity with tß4, while Irobi et al. (7) reported the structure of actin bound to a construct consisting of tß4 residues 21–43 fused to the C-terminus of gelsolin segment 1 (gelsolin residues 27–152). Both Hertzog's and Irobi's structures indicate that the N-terminal helix of tß4 binds SD1 and SD3 of actin, and show the central region of tß4 (residues 15–29) in an extended conformation overlying the nucleotide binding cleft. In both models, the C-terminus of tß4 contacts the pointed end of the actin monomer, although the two models differ in the site of contact. Hertzog et al. (6) favor the C-terminus of tß4 binding directly to SD2 as we proposed (5,8), while Irobi et al. (7) showed residues 30–40 of tß4 in an -helix positioned between actin SD2 and SD4 on the pointed end; the three C-terminal residues of tß4 are unresolved in this structure. This is consistent with our crosslinking studies (5,8), which showed that the N-terminus of tß4 contacts the barbed end of actin, while its C-terminus contacts the pointed end. However, some features of the crystallographic structures, and the models for actin-tß4 based on them, are likely to differ from the actual structure of actin-tß4.
Ciboulot and tß4 have opposite effects on actin tryptophan fluorescence and on the fluorescence of AEDANS-labeled actin. In addition, while tß4 acts as a pure sequestering protein, ciboulot promotes monomer addition to barbed ends (31). Ciboulot and tß4 show substantial sequence homology. However, nonhomologous substitutions occur at several residues, which have been shown by NMR to form close contacts with actin (13). In particular, polar residues T20 and N26 in tß4 are replaced by hydrophobic residues in ciboulot, and Domanski et al. (13) suggest that these substitutions result in differences between the binding sites of the C-terminal segments of ciboulot and tß4. Such differences in the binding of the C-terminal segment may contribute to the observed differences in activity. The design of the gelsolin-tß4 construct was similarly based on limited sequence homology, in this case between tß4 residues 17–23 (LKKTETQ) and gelsolin residues 149–155 (FKHVVPN) (7). Considering that gelsolin segment 1 binds actin with 100-fold higher affinity than even full-length tß4 (32), the gelsolin domain must provide most of the binding energy for the gelsolin-tß4 construct. Thus the position of the gelsolin binding site on actin may constrain the binding of the tß4 segment of the construct, so as to favor a binding mode that is less favorable for full-length tß4.
The conformation of actin SD2 and its contact with tß4 in the complex remain to be determined, since it was not fully resolved in either structure. Our results, particularly the reduced solvent exposure of Gln-41, strongly favor direct interaction of tß4 with SD2 of actin, and therefore support this feature of the Hertzog structure, in contrast to that of Irobi et al. The fact that limited proteolysis of actin on SD2 did not affect the binding of tß4 in an earlier study (13), rather than indicating a lack of contact, may indicate that tß4 binding favors a protease-susceptible conformation of SD2 consistent with conformational rearrangement of this region of actin. The position of tß4 residues 30–40, which Irobi et al. (7) observe in their crystal structure, may be influenced by several factors. Our original NMR and CD study of the actin-tß4 complex (8) showed that although tß4 becomes more structured when bound to actin, it retains substantial segmental mobility. The high mobility in the bound state suggests that tß4 may bind actin in multiple modes with comparable energies that exist in dynamic equilibrium as demonstrated for tropomyosin and actin filaments (30,33). The equilibrium between these modes would likely be dependent on experimental conditions, and thus be influenced by crystal packing forces, ionic composition, and osmolarity of the crystallization solvent. In addition, the structure of the gelsolin-tß4 fusion protein and the position of the gelsolin binding site on actin may favor a binding mode that is less favorable for full-length tß4. A dynamic tß4 C-terminus could account for the apparently conflicting biochemical solution studies demonstrating that tß4 can be chemically crosslinked to SDs 1, 2, and 3 (5,8) as well as the actin-bound nucleotide (34).
The principal biological effect of tß4 is to maintain a reservoir of unpolymerized ATP-actin monomers. Spontaneous polymerization is inhibited by blocking sites involved in intersubunit interactions (22). The FRET data indicate that stable binding to ATP-actin monomers is coupled to reorganization of actin SD2, which is likely to account for the specificity for ATP-actin over ADP-actin monomers. In this respect tß4 differs from cofilin, which also inhibits nucleotide exchange but binds preferably to ADP-actin monomers (1). In addition, the competition between cofilin and tß4 for binding actin may contribute to the balance between F- and G-actin in living cells by modulating dynamics of filament assembly/disassembly at the barbed and pointed ends (1). The ability to modulate the conformation of SD2 may account for the different affinities of tß4 for ATP- and ADP-actin, and allows tß4 to maintain a large reservoir of unpolymerized ATP-actin monomers.
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ACKNOWLEDGEMENTS |
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I.V.D. was supported by grants from the Australian Research Council and the National Health and Medical Research Council of Australia, The Young Australian Researcher 2000 Award from The Australian Academy of Science, and The James Kentley Memorial Scholarship from the University of Sydney, Australia. E.M.D.L.C. was supported by a Hellman Family Fellowship and grants from the American Heart Association (No. 0235203N) and the National Science Foundation (No. MCB-0216834).
Submitted on March 16, 2005; accepted for publication October 17, 2005.
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