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Copyright © 2006 The Biophysical Society. All rights reserved.
Biophysical Journal, Volume 90, Issue 3, 985-992, 1 February 2006

doi:10.1529/biophysj.105.063081

Muscle and Contractility

Thymosin β₄ Induces a Conformational Change in Actin Monomers

Irina V. Dedova^*, Olga P. Nikolaeva^†, Daniel Safer^‡, Enrique M. De La Cruz^§, ,   and Cris G. dos Remedios^*

^* 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
^§ Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, Connecticut

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.

The actin cytoskeleton plays a central role in the basic functions of eukaryotic cells such as maintenance of cell shape, cytoplasmic organization, cell movement, and cell division. This is achieved through a dynamic equilibrium between polymeric and monomeric actin. Many actin-binding proteins (ABP) that regulate the monomer-to-polymer transition by binding monomers, filaments or both, have been identified 1. Thymosin β₄ (tβ₄) is a small (M_r<5kDa) 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. Tβ₄ forms a 1:1 complex with monomeric actin and can bind to F-actin at high concentrations 3,4. β-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 tβ₄ binding surfaces on the actin monomer are unknown and several models have been proposed 4,5,6,7. 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 tβ₄. It has been proposed that the C-terminal region of tβ₄ 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 tβ₄ binding affinity 12. Subtilisin cleavage of actin in the DNase I-binding loop causes a twofold decrease in the affinity of tβ₄13, consistent with SD2 contributing to the stability and strength of the actin-tβ₄ interaction. Image reconstruction of actin filaments chemically crosslinked to tβ₄4 suggests tβ₄ binds SD2 and SD4 as well as the gap between SD1 and SD2 5. Binding of tβ₄ 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 tβ₄ folding upon binding could account for the change in heat capacity 5, and recent evidence favors this interpretation 13. It was also proposed that tβ₄ 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 tβ₄ changes the actin monomer conformation 5 and evaluate the contributions of actin SD2 to tβ₄ binding. Our results are consistent with tβ₄ binding to SD1, SD2 and SD3, causing changes in the spatial orientation of actin monomer subdomains.

5-(((2-Iodoacetyl)amino)ethyl)aminonaphthalene-1-sulfonic acid (IAEDANS), 1,N⁶-ethenoadenosine-5′-triphosphate (ɛATP), 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 (5mM imidazole, pH 7.0, 0.2mM ATP, 0.1mM CaCl₂, 0.5mM DTT, and 1mM NaN₃). Tβ₄ 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 0.63cm⁻¹ (0.1%, 290nm), 1.1cm⁻¹ (0.1%, 280nm), and 0.98cm⁻¹ (0.1%, 280nm), 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), ɛ-ATP 17. The following molar extinction coefficients were used for probes bound to actin: DC, 4640M⁻¹cm⁻¹ at 330nm and DDPM, 3050M⁻¹cm⁻¹ at 440nm 14; IAEDANS, 6100M⁻¹cm⁻¹ at 336nm 15; and FITC, 74,500M⁻¹cm⁻¹ at 493nm 18. The concentration of actin in labeled samples was determined by the DC Protein Assay (λ=795nm, 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 ɛATP as described earlier 17. In brief, nonlabeled or FITC-labeled G-actin (24–48μM) was treated with 0.1vol.mes of Dowex-1 on ice for 10min to remove free ATP. Dowex-1 was removed by filtration and the solution was supplemented with 0.5mM ɛATP, 0.1mM CaCl₂, and 1mM Tris/HCl (pH 8.0) and equilibrated on ice overnight. This procedure was repeated again to ensure a complete exchange of bound ATP with ɛATP. Free nucleotide was removed with Dowex-1 immediately before fluorescence experiments. Because the actin concentrations used are much higher than the K_d 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 (F_DA) and absence (F_D) of the acceptor (A) according to Eq. 1: E=(1−F_DA/F_D)/α, where α 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: R_o 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, κ² is the orientation factor, Q_D is the quantum yield of the D on actin in the absence of the A, and J is the overlap integral given in M⁻¹cm⁻¹nm⁴. R_o distances of actin alone were taken as 49.6Å for IAEDANS/FITC 20 and 47.4Å for ɛATP/FITC 17. These values were corrected for changes in the donor quantum yield 21 due to tβ₄ binding. The R_o 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 κ²=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), ɛATP (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, 0.2–0.4mgml⁻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 F_DA×C_i/F_D×C′_i (Eq. 4), where C_i and C′_i are the concentrations of D in D and D/A samples, respectively. Acrylamide quenching of DC-actin fluorescence (λ_ex=332nm, λ_em=512nm) was performed at 22°C in a thermostated cuvette as described 14. The apparent Stern-Volmer constant (K_SV) was obtained from the slope of a plot of F_o/F versus [acrylamide] by fitting to Eq. 5 as F_o/F=1+K_SV [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 (62.5mM Tris-Cl, pH 6.8, 10% glycerol, and 0.1% bromophenol blue) and separated by native PAGE gel electrophoresis at 70–110V for 90–120min 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 1K min⁻¹. Transition temperatures (T_m) were determined from the peak of thermal transition, and the apparent enthalpies (ΔH°_app) were calculated from the integrated areas of the denaturation peaks.

A donor probe, DC, at Gln-41 and an acceptor, DDPM, at Cys-374 (Fig. 1), located on opposite sides of the small domain of actin were used to measure FRET along the SD1 and SD2 axis. Figure 2A shows corrected and averaged (n=5) fluorescence emission spectra of DC-actin (curve 1) and DDPM-DC-actin (curve 2). A calculated value of 30.3Å for R₀ of the DC/DDPM pair in actin alone (J=9.8×10¹⁵M⁻¹cm⁻¹ nm⁴) and a FRET efficiency of 0.268±0.03 correspond to a distance of 35.8±1.0Å between these probes. Note that the emission maximum of DC is shifted by 15nm to the longer wavelength in the presence of DDPM, suggestive of a possible allosteric effect caused by a modification of Cys-374 consistent with reports on the allosteric relationship between SD1 and SD2 25.

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Figure 1

Position of the donor/acceptor probe pairs used for FRET measurements in actin. Gln-41, Lys-61, Cys-374, and ATP are shown as space-filled structures (26). The numerals refer to the four subdomains.

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Figure 2

Effect of tβ₄ on the distance between Gln-41 and Cys-374 of actin. Corrected and averaged fluorescence spectra (λ_ex=332nm and λ_em=525nm) of DC- and DC/DDPM-actin in the presence and absence of tβ₄. (A) DC- and DC/DDPM-actin (2–5μM) before (solid curves 1 and 2, respectively) and after addition of a 10-fold molar excess of tβ₄ (dashed curves 1′ and 2′, respectively); and DC/NEM-actin in the presence and absence of tβ₄ (dash-dot curves 3 and 3′, respectively). (B) (Left) A Coomassie-stained 12% native PAGE gel of DC/DDPM-actin (lane 1) and its binary complex with tβ₄ (lane 2). (Right) The same gel viewed under UV light. (Symbols: A, monomeric actin; T, tβ₄; and AT, actin/tβ₄ complex.) (C) Stern-Volmer quenching plots: open squares, DC-actin monomers; solid squares, DC-actin filaments; circles, the actin monomer/tβ₄ complex; and triangles, the DNase I/actin monomer complex. All samples were in G-buffer except DC-actin filaments that additionally contained 50mM KCl and 2mM MgCl₂. The temperature was constant (22°C).

Bound tβ₄ decreases the intensity of DC-actin by 12±2% (dashed curve 1′, n=5) in agreement with a previous report 8. However, in the presence of DDPM this decrease is abolished (dashed curve 2′). The resulting FRET efficiency calculated using R₀=29.4Å (corrected for changes in the Q_D) is 0.182±0.03, which corresponds to a distance of 37.8±1.2Å between DC and DDPM in the actin-tβ₄ complex. Thus, tβ₄ increases the distance between the probes attached to Cys-374 and Gln-41 by 2Å (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=31.05Å (recalculated for changes in the Q_D 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±0.9Å. The addition of tβ₄ 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±1.0Å between DC and DDPM in the presence of tβ₄. Thus, tβ₄ increases the measured distance by ∼2Å. Native PAGE gels (Figure 2B) demonstrate that concentrations of tβ₄ used in FRET experiments are sufficient to saturate labeled actin as expected from the equilibrium binding affinities 5. The tβ₄-induced increase in the Gln-41/Cys-374 distance is accompanied by a reduction in the solvent accessibility of Gln-41 (Figure 2C). Acrylamide quenches the fluorescence emission of DC in the G-actin/tβ₄ complex (curve GT, K_SV=1.0±0.3, n=6) less efficiently than in G-actin alone (curve G, K_SV=2.4±0.1, n=6). A similar effect is observed in F-actin (curve F, K_SV=1.3±0.1, n=5) and in the actin-DNase I complex (curve GD, K_SV=1.2±0.1, n=6) where Gln-41 is protected from solvent because it forms part of the binding interface 26.

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±1.1Å, in agreement with a previous report 20. The fluorescence emission spectrum of IAEDANS-actin (Figure 3A, solid curve 1) demonstrates that tβ₄ binding (dashed curve 1′) generates a 39±3% (n=10) decrease in fluorescence intensity at 475nm and an 11-nm red shift of the emission maximum wavelength 5. However, in the presence of FITC (solid curve 2), tβ₄ causes only a 27±4% decrease (λ=475nm, 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 tβ₄ is bound. The ratios of corrected fluorescence intensities (F_DA to F_D at λ=475nm) of the IAEDANS-actin/tβ₄ (dashed curve 1′) and IAEDANS/FITC-actin/tβ₄ complexes (dashed curve 2′) yields a FRET efficiency of 0.59±0.04 (n=9). Q_D of the D significantly decreases in the presence of tβ₄ (0.48 in actin alone compared to ∼0.3 in the actin/tβ₄ complex, n=6). This reduces R₀ from 49.6Å (in actin alone) to 46.5Å (in the actin/tβ₄ complex), yielding a distance between IAEDANS and FITC in the actin/tβ₄ complex of 43.8±0.9Å, a mean reduction of only 0.9Å (p=0.018).

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Figure 3

Effect of tβ₄ on the distance between Lys-61 and Cys-374 of actin. (A) Corrected and averaged fluorescence spectra (λ_ex=365nm and λ_em=520nm) of IAEDANS-actin and IAEDANS/FITC-actin (1.5–3μM) in the presence (dashed curves 1′ and 2′) and absence (solid curves 1 and 2) of 10-fold molar excess of tβ₄. Conditions: G-buffer, 22°C. Spectra were normalized to a 100% acceptor labeling ratio. (Inset) The emission spectrum of FITC-actin excited at 520nm (solid curve) and at 360nm (dashed curve). (B) Effect of DNase I on the corrected and averaged fluorescence spectra of IAEDANS- and IAEDANS/FITC-actin (1.5–3μM; solid curves 1 and 2, respectively) in the presence (dashed curves 1 and 2) and absence (dotted curves 1 and 2) of a 10-fold molar excess of tβ₄. Curve 3 shows an emission spectrum of G-buffer. (C) The reverse order of ABP addition, i.e., tβ₄ (dashed curves) followed by the DNase I (dotted curves). (D) A 10% Native PAGE gel. (Lane 1) IAEDANS/FITC-actin (3μM) alone; (lane 2) 3μM actin+20μM tβ₄; (lanes 3–5) 1.5, 3, and 6μM DNase I was added to the actin/tβ₄ complex; (lane 6) 3μM actin+6μM DNase I; (lane 7) 20μM tβ₄+actin/DNase I complex (shown in lane 6); and (lane 8) 20μM tβ₄+6μM DNase I.

Therefore, although the binding of tβ₄ significantly decreases the FRET efficiency, the effect on the distance between these probes is not substantial. Figure 3A (inset) shows the emission spectrum of actin labeled with acceptor-only (solid curve), excited at 520nm, and the same sample excited at 360nm (dashed curve). From these data it is clear there can be no contribution of FITC fluorescence at 475nm where the FRET is determined. DNase I binding does not alter the emission spectra of IAEDANS- or IAEDANS/FITC-actin (Figure 3B) nor does it affect the FRET efficiency between these sites 27. DNase I inhibits the tβ₄-induced spectroscopic changes (Fig. 3) because it prevents tβ₄ from binding to actin as assayed by native gel electrophoresis (Figure 3D), consistent with common or partially overlapping binding sites 8.

FRET efficiencies between ɛATP and FITC were calculated from comparison of normalized fluorescence intensities of ɛATP/FITC- and ɛATP-actin in the absence and presence of tβ₄. The distance between ɛATP and FITC attached to Lys-61 of actin (in the absence of tβ₄ or DNase I) is 38.4±0.8Å, the R₀ is 47.4Å, and the FRET efficiency is 0.78±0.02 17. Binding of tβ₄ results in a 14±3% (n=6) increase in the ɛATP-actin fluorescence (Fig. 4), which is almost completely eliminated when tβ₄ binds to ɛATP/FITC-actin, indicating that tβ₄ binding increases the FRET efficiency between bound ɛATP 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 2mM ATP, which results in separation of the D- and A-probes and allows normalization of the reference fluorescence with respect to the ɛATP concentration 17. Unlabeled ATP readily competes with ɛATP bound to actin. The comparison of fluorescence intensities of digested ɛ-ATP/FITC- and ɛATP-actin is used for final normalization of FRET efficiency (see Methods). In the presence of tβ₄, the normalized ratio of the fluorescence intensities of ɛATP/FITC-actin to that of ɛATP-actin is 0.16±0.02, i.e., a FRET efficiency of 0.84±0.02. Using an R₀ value of 48.1Å (corrected for the tβ₄-induced increased Q_D), the distance between the probes in the actin/tβ₄ complex is 36.5±1.0Å. Thus, tβ₄ closes the gap between the FITC probe on Lys-61 and ɛATP by 1.9Å (p<0.000).

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Figure 4

Effect of tβ₄ on the distance between bound ɛATP and Lys-61 of actin. Corrected and averaged fluorescence spectra (λ_ex=315nm and λ_em=415nm) of ɛATP- and ɛATP/FITC-actin in the absence (solid curves) and presence of 10-fold molar excess tβ₄ (dashed curves). The actin concentration was 1.5μM in G-buffer containing 1mM Tris/HCl pH 8.0 and 0.1mM CaCl₂. The temperature was 10°C. Spectra were normalized to a labeling ratio of FITC to actin equal to one.

Binding of tβ₄ to G-actin results in thermal stabilization of actin monomers (Fig. 5). The thermal transition of actin monomers has a single maximum at 59.9°C, and an apparent calorimetric enthalpy (ΔH°_app) of 550±20kJ/mol (n=5). Tβ₄ 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 tβ₄ occurs at 61.7°C, and the ΔH°_app increases to 620±25kJ/mol (n=5). The width at the half-height of the thermal denaturation peak, ΔT_0.5, is equal to 6.5°C for the actin/tβ₄ complex but it is 7.5°C for actin alone, suggesting a more cooperative thermal transition with tβ₄ binding. The thermal denaturation of actin in the presence and absence of tβ₄ 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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Figure 5

Effect of tβ₄ on the thermal unfolding of actin. Representative DSC scans of G-actin (A) and actin-tβ₄ (AT). The trace of tβ₄ alone is not shown because it superimposes the buffer baseline. Conditions: 24μM actin and 30μM tβ₄ in G-buffer. Heating rate is 1Kmin⁻¹.

Cofilin, tβ₄, and their binary complexes with actin can be successfully resolved by native polyacrylamide gel electrophoresis 22,23, which has been employed previously to show an increased affinity of cofilin for actin in the presence of DNase I 23. The actin/tβ₄ complex migrates as a distinct band ahead of the actin monomers with free tβ₄ appearing below the actin/tβ₄ band (Figure 6A). 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 tβ₄ from the actin/tβ₄ complex. Cofilin and tβ₄ do not bind each other (lane 1, Figure 6A). 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 tβ₄ competitions are independent of the order of addition (Figure 6B). However, tβ₄ does not compete with cofilin for actin binding in the presence of DNase I (Figure 6C), and the cofilin-DNase I-actin ternary complex 23 is stable even after the addition of a 15-fold molar excess tβ₄ (Figure 6C). However, the addition of the same excess tβ₄ to the actin/cofilin complex in the absence of DNase I leads to the dissociation of the actin/cofilin complex (Figure 6D). Importantly, under experimental conditions, binding of tβ₄ to actin is completely abolished in the presence of DNase I, irrespective of the presence of cofilin (Figure 3DE). We interpret these results to mean that accessibility to the DNase I binding loop is necessary for tβ₄ to compete with cofilin for actin binding, presumably because SD2 forms part of the tβ₄ binding site on actin and contributes to the high affinity.

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Figure 6

Effect of tβ₄ on cofilin-actin and cofilin-actin-DNase I complexes. (Panel A) A representative 10% Native PAGE gel shows the effects of progressive addition of cofilin (C) to the preformed actin-tβ₄ complex (AT). (Lane 1) A mixture of 14μM C and 10μM tβ₄ (T); (Lane 2) 2μM actin alone (A); (Lanes 3–5) a mixture of 2μM A and 4, 6, and 10μM T, respectively; (Lanes 6–9) titration of the preformed AT complex (2μM A and 10μM T) with 2, 6, 10, and 14μM C, respectively; and (Lane 10) a mixture of 2μM A and 14μM C. (Panel B) A representative 10% Native PAGE gel shows the reverse order of ABP addition, i.e., progressive addition of T to the preformed AC complex. (Lane 1) A alone (3μM); (Lane 2) C alone (9μM); (Lane 3) a mixture of 3μM A and 9μM C; (Lanes 4 and 5) addition of 30 and 45μM T, respectively, to the preformed AC complex (3μM A and 9μM C); and (Lane 6) a mixture of 3μM A and 45μM T. (Panel C) A representative 10% Native PAGE gel shows C (12μM), A (3μM), DNase I (D, 5μM), and T (45μM) alone (lanes 1, 2, 3, and 10, respectively). The binary complexes of A with C, D, and T are shown (lanes 4, 5, and 9, respectively). (Lane 6) The ternary complex formed by A, C, and D (A/C/D=1:1.6:4) in the absence of T. (Lane 7) The addition of a 15-fold molar excess T to the preformed ternary ACD complex (shown in lane 6). (Lane 8) The addition of D to preformed AT (1:15) complex in the presence of a fourfold unbound C (this sample before addition of D is shown in lane 12, Gel D). (Panel D) A representative 10% Native PAGE gel shows the AC (1:4) complex (lane 11), and the addition of a 15-fold molar excess T to it (lane 12). (Panel E) This 10% Native PAGE gel shows the AT (1:15) complex (lane 13); the addition of a 1.6-fold molar excess D to it (lane 14); and the control AD (1:1.6) binary complex (lane 15). Gels were run simultaneously.

This investigation was initiated to determine if tβ₄ binding induces a conformational change in actin and to evaluate models of tβ₄ binding to G-actin. Tβ₄ binding induces significant changes in the average conformation of actin monomers as measured by FRET. The distance between probes along one side of the monomer axis (Cys-374 to Gln-41) is increased, whereas the distance down the center (Cys-374 to Lys-61) decreases slightly. There is a significant reduction in the accessibility of the probe attached at Gln-41 when tβ₄ binds, suggesting it forms part of the actin-tβ₄ binding interface 5. The observation that tβ₄ 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 tβ₄ binding. We conclude that tβ₄ 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 tβ₄ enhances the fluorescence of bound ɛATP (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 tβ₄ binding affinity 5,28. When ATP bound to actin is exchanged for ADP, the DNase I binding loop adopts a folded α-helical conformation 11,12,29. Tβ₄ has a much higher affinity for ATP-actin than ADP-actin monomers 5,28, suggesting that the conformation of SD2 dictates how tightly tβ₄ binds actin 12 and that the higher affinity of tβ₄ for ATP-actin results from stabilizing contacts made with SD2 5,8.

The detailed three-dimensional structure of the actin-tβ₄ complex has not been determined experimentally. However, two alternatives have been proposed for tβ₄ binding to actin monomers, based on structures of actin bound to tβ₄ homologs 6,7. Hertzog et al. 6 reported the structure of actin bound to domain 1 of ciboulot, which shares 30% sequence identity with tβ₄, while Irobi et al. 7 reported the structure of actin bound to a construct consisting of tβ₄ 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 tβ₄ binds SD1 and SD3 of actin, and show the central region of tβ₄ (residues 15–29) in an extended conformation overlying the nucleotide binding cleft. In both models, the C-terminus of tβ₄ 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 tβ₄ binding directly to SD2 as we proposed 5,8, while Irobi et al. 7 showed residues 30–40 of tβ₄ in an α-helix positioned between actin SD2 and SD4 on the pointed end; the three C-terminal residues of tβ₄ are unresolved in this structure. This is consistent with our crosslinking studies 5,8, which showed that the N-terminus of tβ₄ 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-tβ₄ based on them, are likely to differ from the actual structure of actin-tβ₄.

Ciboulot and tβ₄ have opposite effects on actin tryptophan fluorescence and on the fluorescence of AEDANS-labeled actin. In addition, while tβ₄ acts as a pure sequestering protein, ciboulot promotes monomer addition to barbed ends 31. Ciboulot and tβ₄ 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 tβ₄ 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 tβ₄. Such differences in the binding of the C-terminal segment may contribute to the observed differences in activity. The design of the gelsolin-tβ₄ construct was similarly based on limited sequence homology, in this case between tβ₄ 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 tβ₄32, the gelsolin domain must provide most of the binding energy for the gelsolin-tβ₄ construct. Thus the position of the gelsolin binding site on actin may constrain the binding of the tβ₄ segment of the construct, so as to favor a binding mode that is less favorable for full-length tβ₄.

The conformation of actin SD2 and its contact with tβ₄ 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 tβ₄ 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 tβ₄ in an earlier study 13, rather than indicating a lack of contact, may indicate that tβ₄ binding favors a protease-susceptible conformation of SD2 consistent with conformational rearrangement of this region of actin. The position of tβ₄ 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-tβ₄ complex 8 showed that although tβ₄ becomes more structured when bound to actin, it retains substantial segmental mobility. The high mobility in the bound state suggests that tβ₄ 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-tβ₄ fusion protein and the position of the gelsolin binding site on actin may favor a binding mode that is less favorable for full-length tβ₄. A dynamic tβ₄ C-terminus could account for the apparently conflicting biochemical solution studies demonstrating that tβ₄ 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 tβ₄ 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 tβ₄ differs from cofilin, which also inhibits nucleotide exchange but binds preferably to ADP-actin monomers 1. In addition, the competition between cofilin and tβ₄ 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 tβ₄ for ATP- and ADP-actin, and allows tβ₄ to maintain a large reservoir of unpolymerized ATP-actin monomers.