©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Kinetics of the Interaction of Myosin Subfragment-1 with G-Actin
EFFECT OF NUCLEOTIDES AND DNaseI (*)

(Received for publication, October 19, 1994; and in revised form, December 19, 1994)

Laurent Blanchoin Stéphane Fievez Franck Travers (§) Marie-France Carlier (¶) Dominique Pantaloni

From the Laboratoire d'Enzymologie du CNRS, 91198 Gif-sur-Yvette Cedex, France

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
Appendix
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The kinetics of interaction of monomeric pyrenyllabeled G-actin with myosin subfragment-1 (S(1) (A(1)) and S(1)(A(2)) isomers) has been examined in the stopped-flow at low ionic strength. The data confirm the previously reported existence of binary GS and ternary G(2)S complexes. The increase in pyrenyl-actin fluorescence which monitors the G-actin-S1 interactions is linked to the isomerization of these complexes following rapid equilibrium binding steps. The rates of isomerization are 200 s for GS and 50 s for G(2)S at 4 °C and in the absence of ATP. DNaseI and S(1) bind G-actin essentially in a mutually exclusive fashion. Both GS and G(2)S are dissociated by MgATP and MgADP. The kinetics and mechanism of ATP-induced dissociation of G(2)S are quantitatively close to the ATP-induced dissociation of F-actin-S(1), which indicates the G(2)S is a good model for the F-actin-S(1) interface. GS and G(2)S display different kinetic behaviors in response to nucleotides, GS being less efficiently dissociated than G(2)S by MgATP. This result suggests that different mechanical properties of the cross-bridge might correlate with different orientations of the myosin head and different actin/myosin binding ratios.


INTRODUCTION

Kinetic studies of the contractile cycle of the actomyosin ATPase have shown that the energy of ATP hydrolysis is transduced into movement via changes in affinity of myosin for F-actin, modulated by the nucleotide bound to myosin(1, 2, 3, 4, 5) . A complete understanding of the mechanism of transduction of chemical energy into mechanical energy necessitates a description of the structural changes in the actomyosin interface in connection with the biochemical steps. The recent determinations of the three-dimensional structures of actin(6) , of the myosin head(7) , and the combination of these crystallographic data with the low resolution electron microscopy structure of the decorated filament (8) have led to a structural model for the F-actin-myosin subfragment-1 rigor complex(9) .

An alternative approach to the functional changes in the actomyosin interface consists in characterizing the small size complexes formed between the myosin head and monomeric actin. These complexes, which are precursors in the myosin subfragment-1 (S(1))-induced polymerization of G-actin into decorated filaments, are helpful to define the minimum actin-S(1) contractile unit. It was found (10) that S(1) forms binary (GS) and ternary (G(2)S) tight complexes with G-actin at low ionic strength and in the absence of ATP. Both nucleotide effects on the stability of GS and G(2)S and cross-linking studies (11) showed that the G-actin-S(1) interface is very similar to the F-actin-S(1) interface in the rigor state. The second step in the polymerization process of G-actin induced by S(1) appears to be the condensation of G(2)S units in small oligomers (12) , which emphasizes the functional relevance of the interaction of the myosin head with two actin molecules. The interaction of the myosin head with two actin subunits, adjacent along the long pitch helix of the filament, is also an essential feature of the structural model of the actin-myosin rigor complex(9) , which may have important implications in the mechanism of force generation. The present paper is a kinetic analysis of the interaction of monomeric pyrenyl-G-actin with the two myosin subfragment-1 isoforms carrying either the A(1) or the A(2) light chain. The pyrene probe has previously been used to characterize the interaction of S(1) with F-actin (13, 14, 15) and G-actin(10) . The kinetics of G-actin interaction with S(1)(A(1)) and S(1)(A(2)) presented here confirm the existence of GS and G(2)S complexes with both S(1) isoforms and show that G(2)S is more easily dissociated by MgATP than GS, which may have implications in muscle contraction.


MATERIALS AND METHODS

Proteins

Actin was purified from rabbit muscle acetone powder (16, 17) and isolated as CaATP-G-actin through Sephadex G-200 chromatography (18) at 4 °C in G buffer (5 mM Tris-Cl pH 7.8, 0.2 mM dithiothreitol, 0.1 mM CaCl, 0.2 mM ATP, 0.01% NaN(3)). Actin was fluorescently labeled on Cys using pyrene iodoacetamide(19) . The labeling stoichiometry was routinely 0.80-1.0. The 1:1 complex ATP-G actin without free ATP was prepared by treatment of CaATP-G-actin solutions in G buffer with Dowex 1-X8 equilibrated in G(0) buffer (i.e. G buffer containing no ATP) (20) . Myosin was purified from rabbit back and leg muscles as described (21) up to the ammonium sulfate fractionation step (included) and stored at 6-10 mg/ml at -20 °C in 75 mM potassium phosphate buffer, pH 7.5, containing 1 mM EDTA and 50% glycerol. Chymotryptic myosin subfragment-1 (22) was resolved in S(1)(A(1)) and S(1)(A(2)) isomers by sulfopropyl bisacrylamide cation-exchange chromatography (12) , stored on ice at 50-80 µM in 10 mM MOPS, (^1)pH 7.0, 20 mM NaCl, 1 mM dithiothreitol. Both actin and S(1) were used within 2 weeks following purification. Just prior to the experiments, S(1) was rapidly equilibrated in G(0) buffer by 1-h dialysis using Spectra Por 2 membranes.

Bovine pancreatic DNaseI grade II was purchased from Boehringer and purified by hydroxylapatite chromatography(23) . The concentration of DNaseI was determined by titration of an aliquot of the purified preparation by G-actin using the DNase inhibition assay(24, 25) . The DNA substrate was bought from Sigma.

Actin and S(1) concentrations were determined spectrophotometrically as described(12) .

Stopped-flow Experiments

The time course of reaction of S(1)(A(1)) or S(1)(A(2)) with pyrenyl-labeled G-actin was monitored by fluorescence using a stopped-flow (DX.17 MV, Applied Photophysics), with the excitation monochromator set at 366 nm and a KV 380 Schott filter placed on the emission beam. Experiments were carried out in G(0) buffer at 4 °C, to avoid the formation of actin-S(1) oligomers(12) . The signal was analyzed using a software attached to the instrument. The dead time of the apparatus was 0.8 ms.

Dissociation of G-actin S(1) complexes by nucleotides was carried out as follows. One of the drive syringes contained either calcium nucleotide (prepared as an equimolar CaCl(2)-nucleotide mixture diluted to the desired concentrations in buffer G(0)), or magnesium nucleotide (prepared in the same way but diluted in buffer G(0) containing 100 µM MgCl(2) instead of 100 µM CaCl(2)). The other drive syringe contained the G-actin-S(1) complex rapidly preformed at 0 °C immediately before being brought in the syringe. For experiments with magnesium nucleotide, G-actin-S(1) was in buffer G(0) containing only 10 µM CaCl(2) so that after mixing the nucleotide was 90% magnesium nucleotide.

Data Analysis and Simulation of Kinetics

Examination of both the amplitudes and the kinetics of the change in pyrenyl-actin fluorescence upon interaction with S(1), in series of experiments in which the concentrations of either pyrenyl-G-actin or S(1) were varied, led us to propose the minimum kinetic model able to account for the data. Values of equilibrium dissociation constants and intrinsic fluorescence of the different actin and actin-S(1) complexes were derived from the actin and S(1) concentration dependences of the amplitudes and observed rate constants as described under ``Appendix.'' Time courses of the change in pyrene fluorescence under various experimental conditions were simulated according to this proposed model, using the KINSIM softwares kindly provided to us by Drs. Carl Frieden and Thomas Pollard. Values of yet undetermined rate constants were then adjusted to obtain the best fit of simulated kinetic curves to experimental stopped-flow traces.


RESULTS

Kinetics of the Interaction of Pyrenyl-G-actin with S(1)(A(1)) or S(1)(A(2))

In a first series of experiments, pyrenyl G-actin at a constant concentration (2 µM) was rapidly mixed with S(1)(A(1)) or S(1)(A(2)) at different concentrations in the range 0-20 µM. The resulting increase in pyrene fluorescence fitted a single exponential process at all S(1) concentrations. Fig. 1a shows typical time courses of the reaction at three different concentrations of S(1)(A(1)) and the exponential best fit.


Figure 1: Rapid kinetics of complex formation between pyrenyl G-actin and myosin subfragment 1. a, pyrenyl G-actin-ATP 1:1 complex (2 µM) in G(0) buffer was mixed at 4 °C with S(1)(A(1)) at the following concentrations (in µM, top to bottom): 4, 1, 0.2. The fluorescence of 2 µM pyrenyl-G-actin is normalized to 1. b, S(1)(A(1)) (0.4 µM) was mixed with pyrenyl-G-actin at the following concentrations in µM (top to bottom): 2, 0.6, 0.2. The fluorescence of pyrenyl G-actin is subtracted. Noisy curves are experimental traces (average of four consecutive shots); solid lines represent the monoexponential best fit; dashed lines represent the simulated time courses using KINSIM and the fluorescence, equilibrium and rate parameters given in Table 1.





In a second series of experiments, the concentration of S(1) was kept constant (0.4 µM), and the concentration of pyrenyl-G-actin was varied. In the whole range of actin concentrations (0-2 µM), the time courses of the increase in pyrene fluorescence were again well fitted by monoexponentials (Fig. 1b).

The actin and S(1) concentration dependences of the amplitudes and k were analyzed as follows. When the concentration of pyrenyl-G-actin was constant, the amplitude of the fluorescence change showed a saturation behavior upon increasing S(1)(A(1)) or S(1)(A(2)) concentration. As illustrated in Fig. 2, and in agreement with previous static fluorescence titration curves(12) , all data points fell above the stoichiometric line corresponding to the formation of a 1:1 GS complex of infinitely high affinity and were close below the stoichiometric line corresponding to a 2:1 G(2)S complex of infinitely high affinity. Similar data were obtained with S(1)(A(1)) and S(1)(A(2)), S(1)(A(1)) displaying an apparently slightly higher affinity than S(1)(A(2)). The amplitude of the fluorescence change was maximum at 2-3 µM S(1) and did not show any further increase with up to 20 µM S(1). The maximum increase in pyrenyl G-actin fluorescence at saturation by S(1) was 2.1-fold in satisfactory agreement with the 3.3-fold enhancement observed (10) in equilibrium measurements performed with higher spectral resolution of excitation and emission wavelengths.


Figure 2: Fluorescence titration curves of pyrenyl G-actin by S(1) (stopped-flow amplitude data). Pyrenyl G-actin (2 µM) was mixed with S(1)(A(1)) () or S(1)(A(2)) (box) at the indicated concentrations under the conditions described in Fig. 1a. Thin lines represent the theoretical titration curves of infinitely high affinity G(2)S (left) and GS (right) complexes, respectively. Thick lines are calculated curves within using values of parameters in Table 1.



When the concentration of S(1) was constant the dependence of the amplitude of the fluorescence change on G-actin concentration, shown in Fig. 3, was consistent with the formation of a tight G(2)S complex when S(1) was saturated by G-actin. Note that interpretation of the fluorescence titration curve of S(1) by G-actin within the formation of a single 1:1 GS complex would imply that the affinity is low (K(D) 0.3 µM), and this interpretation would be totally inconsistent with the titration of G-actin by S(1) shown in Fig. 2. Solid curves in Fig. 2and Fig. 3are calculated within , (see ``Appendix''), and values of parameters shown in Table 1.


Figure 3: Fluorescence titration curves of S(1) by pyrenyl G-actin (stopped-flow amplitude data). Pyrenyl G-actin at the indicated concentrations was rapidly mixed with 0.4 µM S(1)(A(1)) () or S(1)(A(2)) (box), under conditions described in Fig. 1B. The thin lines represent the theoretical titration curves corresponding to infinitely high affinity GS (left) and G(2)S (right) complexes, respectively. Thick lines are calculated curves within using values of parameters given in Table 1.



Analysis of the rate constants showed the following. When G-actin was constant, the observed first-order rate constant for the fluorescence increase varied with S(1) in a sigmoïdal fashion, as shown in Fig. 4, between two limits of 50 s at low S(1) to 200 s at saturating S(1) (20-30 µM). On the other hand, as shown in Fig. 5, when S(1) was constant, the observed first-order rate constant decreased upon increasing G-actin from 100 s to a lower limit of 55-60 s at high G-actin. The fact that the rates reach a limit indicates that the change in fluorescence monitors a conformation change following the formation of rapid equilibrium complexes. However, the complex formed when G-actin is saturated by S(1) (k = 200 s) is not identical to the one obtained when S(1) is saturated by G-actin (k = 55 s). Hence the kinetic data confirm the interpretation of previous (10) and present amplitude data and lead to the conclusion that the rate constant of 200 s represents the sum of the rate constants for isomerization of a 1:1 GS complex, while the rate constant of 55-60 s represents the sum of the rate constants for isomerization of a 2:1 G(2)S complex. The sigmoidal appearance of the change in kversus S(1) (Fig. 4) reflects the fact that, as illustrated in Fig. 10a under ``Appendix,'' the formation of G(2)S is the predominant reaction (k = 55 s) at low S(1), and the formation of GS* (k = 200 s) becomes increasingly predominant upon increasing S(1). The converse argument holds for the decrease in k upon increasing G-actin at constant S(1) (compare Fig. 5and 10b).


Figure 4: S(1) concentration dependence of the first-order rate constant for interaction of pyrenyl G-actin with S(1). The first-order rate constant for the change in fluorescence observed upon reaction of pyrenyl G-actin (squares, 2 µM; circles, 1 µM) with S(1)(A(1)) (, bullet) or S(1)(A(2)) (box, circle) is plotted as a function of [S(1)]. Closed triangles represent the monoexponential best fit of simulated time courses using KINSIM and the parameter values given in Table 1. Inset, expanded view of the data at low [S(1)].




Figure 5: G-actin concentration dependence of the first-order rate constant for interaction of pyrenyl G-actin with S(1). Pyrenyl-G-actin at the indicated concentrations was reacted with 0.4 µM S(1)(A(1)) (, bullet) or S(1)(A(2)) (box, circle). Squares and circles refer to two independent experiments. Large squares represent the rate constant of the exponential best fit of simulated curves (e.g.dashed lines in Fig. 1a) using KINSIM and values of parameters in Table 1.




Figure 10: Distribution of the different fluorescent and non-fluorescent G-actin-S(1) complexes in a range of G-actin and S(1) concentrations. a, molar fraction of actin in different species at 2 µM total actin and different concentrations of S(1)(A(1)). Dashed line, free G-actin; , GS; bullet, GS (fluorescent); box, G(2)S; circle, G(2)S (fluorescent). b, molar fraction of S(1)(A(1)) in different species at 0.4 µM total S(1) and different G-actin concentrations. Dashed line, free S(1)(A(1)); other symbols as in panela.



The above results lead us to propose the simple following model () for binding G-actin to S(1).

In the above scheme, GS and G(2)S are rapid equilibrium complexes. The fluorescence change is associated to the isomerization steps leading to GS and G(2)S. This scheme therefore represents an extension of the one previously proposed (10) in which no isomerization of GS and G(2)S was included. The details of the procedure used to derive the analytical expression of the fluorescence change at equilibrium within , to fit the values of the equilibrium dissociation constants to the data, to derive the rate constants, and simulate the kinetic curves are all given under ``Appendix.''

Fig. 1, a and b, show the superimposed experimental and calculated time courses at a constant G-actin (resp. S(1)) concentration and varying S(1) (resp. G-actin) in a 10-20-fold concentration range. The simulated time courses were consistent with monoexponentials. The calculated rate constants varied with either S(1) or G-actin in a manner quantitatively consistent with experimental data (large symbols in Fig. 4and Fig. 5), which confirmed the validity of .

In all experiments presented in Fig. 1Fig. 2Fig. 3Fig. 4Fig. 5, G-actin has CaATP as bound nucleotide. Since under physiological conditions actin has MgATP as bound nucleotide, supplementary experiments were carried out in which CaATP-pyrenyl-G-actin 1:1 complex at a concentration leq4 µM was rapidly converted into MgATP-G-actin by addition of 15 µM MgCl(2) and 0.2 mM EGTA 3 min before being rapidly mixed with S(1). The amplitude of the pyrene fluorescence change upon addition of S(1) gave titration curves (data not shown) superimposable with those obtained in Fig. 2and Fig. 3(controls with CaATP-G-actin from the same preparation were run in parallel in the same experiment). The rate constants for the isomerizations, k + k and k + k, were 80 s for GS GS* and 30 s for G(2)S G(2)S transitions, respectively, i.e. about twice lower than the corresponding values obtained with calcium G-actin. Hence all conclusions drawn here with calcium-actin also apply to the physiological magnesium-actin species. For technical reasons the work was done with calcium-actin which is more stable and does not give rise to oligomers with concomitant hydrolysis of actin-bound ATP.

Displacement of Pyrenyl G-actin from G-actin-S(1) Complexes by an Excess of Unlabeled G-actin (Chase Experiment)

To test the proposed model for the mechanism of interaction of G-actin with S(1), a chase experiment was designed in which pyrenyl actin was chased off the G-actin-S(1) complexes by unlabeled actin. Preformed complexes were rapidly mixed with unlabeled G-actin at different concentrations. A monoexponential decrease in pyrenyl fluorescence was observed. The amplitude of the change increased with G-actin, total dissociation of the fluorescent complex being observed at saturation by G-actin (data not shown). The rate constant of the exchange process decreased upon increasing G-actin from a value of 140 s at low G-actin to a limit value of 60 s at saturating G-actin levels in a fashion similar to the change in k upon increasing pyrenyl G-actin shown in Fig. 5. The interpretation of these data within is as follows: the exchange of unlabeled for labeled G-actin in the non-fluorescent actin-S(1) complexes is extremely rapid, and the fluorescence change is rate-limited by the isomerization processes, as in the association reaction of S(1) with pyrenyl-G-actin, and at saturation by G-actin the observed rate constant in the chase experiment again represents k + k = 60 s.

Dissociation of G-actin-S(1) Complexes by MgATP, CaATP, and MgADP

Previous static measurements (12) have shown that the GS and G(2)S complexes were dissociated by ADP and, more efficiently, by ATP. The kinetics of dissociation were monitored by the decrease in pyrene fluorescence upon rapid mixing of ATP or ADP with preformed G-actin-S(1) complexes.

The decrease in fluorescence was a first-order process at all nucleotide concentrations. The amplitudes of the fluorescence decrease showed a saturation behavior as a function of nucleotide concentration, and the rate constants increased linearly with nucleotide concentration up to at least 400 µM, i.e. above the range shown in the figures.

The minimum kinetic scheme accounting for this process is the following.

where n = 1 or 2, and N represents the nucleotide (MgATP or CaATP or MgADP). K and K(D) are rapid equilibrium dissociation constants and k and k the isomerization rate constants corresponding to the conformational change, which is monitored by the change in fluorescence and through which the affinity of S(1) for G-actin decreases.

1) The dissociation of G(2) S by nucleotides was first examined. Pyrenyl G-actin (3 µM) and S(1)A(1) (0.5 µM) were premixed and reacted with nucleotide at different concentrations. The results are shown in Fig. 6. Given a total concentration of 0.5 µM ATP-binding sites, the amplitude data (Fig. 6a) showed that the equilibrium dissociation constant for MgATP, K(T)k/k was too low (leq0.1 µM) to be measured. In other words, with MgATP, the value of K(D) is very high, and the reaction is quasi irreversible. From the slope of the linear increase of k with MgATP (Fig. 6b), a value of 1.6 µM s was derived for k+/K(T). The value of k, as derived from the ordinate origin, was leq0.2 s. Only a lower estimate of 400 µM can be proposed for K(T).


Figure 6: ATP and ADP induced dissociation of (G-actin)(2)-S(1) ternary complex. Preformed G(2)S complex (3 µM pyrenyl-G-actin, 0.5 µM S(1)(A(1)) was rapidly mixed with either MgATP (box) or MgADP (circle) or ATP-Ca () at the indicated concentrations. The decrease in pyrenyl fluorescence was consistent with the complete dissociation of S(1) from G-actin at saturation by all nucleotides. Panel a, amplitude data. Panel b, observed dissociation rate constant. Inset, expanded view of the data at low [ATP] or [ADP].



When the dissociating nucleotide was CaATP or MgADP, the values found for k(T) times k/k were 2.8 and 1 µM, respectively. The slopes of kversus CaATP or MgADP were similar to those obtained with MgATP, however, the values of k were definitely higher (2 s), indicating that the reaction is reversible and that the value of K(D) must be lower than with MgATP.

2) The dissociation of GS by nucleotides was investigated in a similar fashion, and qualitatively similar data were obtained, pointing out to the same mechanism for nucleotide-induced dissociation of GS. However, a large quantitative difference between GS and G(2)S was observed regarding the efficiency of MgATP to promote dissociation of S(1) from G-actin. In the experiment shown in Fig. 7, the GS complex was preformed and rapidly mixed with different amounts of MgATP. Essentially GS was present, with S(1)/G ratios of 5 and 16, respectively. The decrease in fluorescence linked to dissociation of GS varied with MgATP in an identical fashion at the two S(n)/G ratios, consistent with a much lower affinity of MgATP for GS than for G(2)S. In this experiment, MgATP binds to GS and free S(1). The fact that identical amplitude patterns were observed although the concentration of S(1) was different in the two assays (hence some competition between GS and S(1) for binding ATP was expected), suggests that the binding of MgATP to GS is completed before its binding to S(1). The analysis of the rate constants, gave k = 0.4 ± 0.1 s and k/K(T) = 0.08 ± .01 µM s (Fig. 7, inset), a value 20-fold lower than the one (1.6 µM s, Fig. 6b) obtained for G(2)S. Using these two numbers, a value of 5 ± 1.8 µM was derived for K`(T). This estimate is consistent with the amplitude data which can be well fitted using an equilibrium dissociation constant K`(T) of 3 µM, a value again 30-fold higher than the one found for the binding of MgATP to G(2)S. An alternative model implying dissociation of GS as a consequence of the initial binding of MgATP to free S(1) has not been considered here.


Figure 7: MgATP dissociates S(1) more efficiently from the G(2)S than from the GS complex. The preformed GS complex was rapidly mixed with MgATP at the indicated concentrations. The amplitude of the fluorescence decrease linked to the dissociation of S(1)-MgATP is plotted versus the concentration of MgATP. bullet, , 1 µM pyrenyl-G-actin, 5 µM S(1)(A(1)); circle, 1 µM pyrenyl-G-actin, 5 µM S(1)(A(2)); , 0.5 µM pyrenyl-G-actin, 8 µM S(1)(A(1)) (normalized to the same up triangleF(0), i.e. to 1 µM G-actin). Continuous lines are calculated within , and with k/K = 0.08 µM s, k = 0.4 s, values coming from the k data () shown in the inset. The data obtained for dissociation of G(2)S (box) coming from Fig. 6a are replotted, for comparison, in the main frame and in the inset.



In conclusion both amplitude and kinetic data indicate that MgATP is much more efficient to promote dissociation of G(2)S than to promote dissociation of GS.

A comparison of these data, with the ATP-induced dissociation of F-actin-S(1)(26) is interesting. At 0.5 °C and in the presence of 0.1 M KCl, values of K`(T) < 0.01 µM, and k/K(T) = 0.6 µM s have been reported, which compare well with the data obtained here for the dissociation of G(2)S by MgATP, but differ significantly from the data obtained for the dissociation of GS.

All equilibrium and rate parameters for nucleotide-induced dissociation of GS and G(2)S are summarized in Table 2.



DNaseI Promotes the Dissociation of G-actin

DNaseI forms a very tight 1:1 complex with G-actin, with an equilibrium dissociation constant of 0.5-2 nM(24, 25, 26, 27, 28) . DNaseI interacts with the exposed loops of subdomains 2 and 4 of actin, bridging the ATP binding cleft at the top of the pointed end of the molecule(6) , a region spatially distinct from the main myosin-binding site(9) . Therefore, the formation of a ternary complex DNaseI-actin-S(1) is theoretically possible and indeed has been reported(29, 30, 31) . One report showed that the inhibition of DNaseI activity by G-actin was not relieved by S(1)(30) , which led the authors to conclude that the affinity of DNaseI for G-actin was unaffected by S(1); the fluorescence titrations of pyrenyl G-actin by S(1) in the presence of DNaseI indicated that the affinity of S(1) for G-actin was decreased at most 4-5-fold by DNaseI(30, 31) , which allowed the formation of a ternary complex in a micromolar range of concentration of all three proteins. Whether the fluorescence of pyrenyl actin in the ternary complex DNase-actin-S(1) is the same as or lower than in the G-actin-S(1) complex does not appear clearly from available data.

The binding of S(1) to pyrenyl G-actin in the presence of DNaseI was investigated in the stopped-flow as follows. A first experiment showed that upon mixing DNaseI with the preformed fluorescent pyrenyl-G-actin-S(1) complex (0.5 µM G-actin, 3 µM S(1)(A(2))), a decrease in pyrene fluorescence was observed. At saturation by DNaseI, the final fluorescence level was identical to that of G-actin. At all concentrations of DNaseI, the decrease in fluorescence was a first-order process whose rate constant k increased with the concentration of DNaseI and reached a higher limit of 60 s at saturation by DNaseI. The DNaseI dependences of the amplitudes as well as of the k were both described by the same hyperbolic binding isotherm.

In another experiment shown in Fig. 8, pyrenyl G-actin in the presence of different amounts of DNaseI was mixed with S(1)(A(2)) at a constant concentration. Three series of experiments were performed concentrations of S(1)(A(2)). The extent of fluorescence change reflecting binding of S(1) decreased upon increasing DNaseI concentration. The process was first-order with a rate constant of 200 s at all DNaseI concentrations. The data were tentatively analyzed within a general square model described below () within which binary DNaseIG-actin (DG) and G-actin-S(1) (GS) complexes and a ternary DNaseI-G-actin-S(1) complex (DGS) can be formed, with equilibrium dissociation K(D) and K(S) in the binary complexes, K`(D) and K`(S) in the ternary complexes, with K(D)K`(S) = K(S)K`(D) (detailed balance). It was assumed from experimental evidence that the fluorescence of pyrenyl G-actin was enhanced only in the GS complex.


Figure 8: Mutual exclusion binding of DNaseI and S(1) to G-actin. Pyrenyl-G-actin (1 µM) in the presence of the indicated concentrations of DNaseI was rapidly mixed with S(1)(A(1)) at 3 µM (circle, box), 8 µM (bullet), and 15 µM (). The increase in pyrenyl fluorescence was monitored. Solid lines are calculated within the general square model () using the following values of equilibrium parameters: K = 3 nM; K = 63 nM; K` = 5 µM; K` = 105 µM. Inset, concentration of DNaseI, [DNaseI], at which 50% of the maximum fluorescent change was observed, as a function of S(1)(A(1)) concentration.



The amplitude data in Fig. 8were analyzed within , using K(D) = 3 nM, and K(S) = 63 nM. The data could be fitted by only if K`(D)/K(D) (= K`(s)/K(s)) was higher than 10^3, which essentially reduces to a mutual exclusion binding behavior of DNaseI and S(1) to G-actin, the ternary complex DGS being negligible in solution in the range of concentrations of S(1) and DNaseI investigated. Accordingly, the concentration of DNaseI causing half-dissociation of G-actin-S(1) varies linearly with S(1) (Fig. 8, inset) consistent with the apparent competitive binding of DNaseI and S(1) to G-actin. Kinetic data indicate that S(1) and DNase dissociate from the ternary complex at rates 60 s and >200 s, respectively.

If S(1) and DNaseI bind G-actin in competition with each other as indicated by the above result, the inhibition of DNaseI activity by G-actin should be relieved by S(1). Surprisingly, the opposite result was obtained, and we confirmed the data obtained by Chen et al. ((30) , Fig. 5) when we performed an identical experiment. However, a control experiment displayed in Fig. 9demonstrated that the presence of 40 µg/ml DNA (the substrate concentration used in the DNase activity assay) very potently inhibits the interaction between G-actin and S(1), presumably due to the binding of the polyanionic DNA to the lysines of S(1) which are involved in the G-actin-S(1) contact. In conclusion, the result obtained by Chen et al.(30) , and confirmed here, is explained by the failure of S(1) to bind to G-actin in the DNA-containing DNase activity test and should not be taken as a proof that S(1) does not significantly inhibit DNaseI binding to G-actin. The DNase activity assay simply appears inappropriate to examine the effect of S(1) on DNaseI binding to G-actin.


Figure 9: The interaction between G-actin and S(1) is inhibited by DNA. Pyrenyl-G-actin (2 µM) was mixed with S(1)(A(2)) at the indicated concentrations, in the absence (bullet) or in the presence () of 40 µg/ml DNA. Note that the affinity of S(1) for G-actin decreases by 3 orders of magnitude (K 3 µM) in the presence of 40 µg/ml DNA.




DISCUSSION

The kinetic results presented here bring new information about the mechanism of interaction of the myosin head (myosin subfragment-1 S(1)(A(1)) and S(1)(A(2)) isomers) with monomeric actin using the fluorescence of pyrenyl-actin as a probe. In agreement with previous reports (10, 12, 32) but at variance with others(31, 33) , S(1)(A(1)) and S(1)(A(2)) can both interact with two molecules of monomeric actin, hence binary (GS) and ternary (G(2)S) complexes exist in solution. Kinetic analysis of the interaction of pyrenyl-G-actin with S(1) further shows that the increase in pyrene fluorescence which monitors the formation of GS and G(2)S is linked to an isomerization of these complexes following the rapid bimolecular reversible reactions of G-actin with S(1). Protein-protein interactions are tightened about one order of magnitude by this isomerization, hence the fluorescent GS and G(2)S* complexes are the major G-actin-S(1) complexes at equilibrium. The isomerization rate constants of GS and G(2)S are appreciably different (200 s and 50 s at 4 °C) which, added to the analysis of the fluorescence titration curves of pyrenyl G-actin by S(1) and of S(1) by pyrenyl G-actin (stopped-flow amplitude data) leaves no doubt about the existence of the two types of G-actin-S(1) complexes. One should note that although a recent report (31) states that G-actin interacts with S(1) with a 1:1 stoichiometry, the data shown ( Fig. 5in Ref.31) cannot be interpreted within a 1:1 binding scheme.

The kinetics of S(1)(A(1)) and S(1)(A(2)) association to G-actin were practically identical, especially regarding the isomerization reactions. Hence the quantitative difference in ability of the two isomers to promote actin polymerization (10, 12, 33) is linked to the different stabilities of short oligomers, resulting from the condensation of the G(2)S complex, which are kinetic intermediates in the formation of arrowhead-decorated filaments(12) . Since hydrophobic actin-actin contacts, plausibly corresponding to the lateral bonds along the genetic helix of the filament, are involved in oligomers, the results suggest that the 40 amino acid extension of light chain A(1) somehow stabilizes actin-actin bonds along the genetic helix, in decorated F-actin-S(1)(A(1)) filaments, rather than longitudinal actin-actin bonds.

The kinetic scheme which is presented here is the simplest which can account for the data. This does not eliminate the possibility that other more complex schemes, in particular containing a larger number of kinetic steps connecting the different complexes, may be relevant. The simple scheme, however, provides an easy comparison of the characteristics of the interaction of the myosin head with monomeric actin and with filamentous actin. The changes in pyrenyl F-actin fluorescence upon binding S(1) have also shown evidence for a similar two-step binding mechanism, the change in fluorescence being linked to the second isomerization step(14, 15) . The rate constant for isomerization of pyrenyl-F-actin-S(1) was 200 s at 20 °C and 20 s at 6 °C and low ionic strength(15) . This latter value is close to the isomerization rate constant found here for G(2)S, which supports the view that the G(2)S complex may be a good model of the F-actin-S(1) interface. The possibility that S(1) interacts with two actin subunits in the filament, adjacent along the long pitch helix, in a geometry similar to that of the G(2)S complex, has been raised by the image reconstruction of the decorated filament (8) and is an intrinsic feature of the current atomic model of the F-actin-myosin complex(9) , supported by available biochemical cross-linking studies(34, 35, 36) . Within this model the two F-actin subunits, called actin 1 and actin 2, which are part of the F-actin-S(1) rigor complex, interact longitudinally, the barbed end (subdomains 1 and 3) of actin 1 being in contact with the pointed end (subdomains 2 and 4) of actin 2. The myosin head interacts mainly with the N- and C-terminal regions of actin 1 (in subdomain 1), and makes a secondary contact with the top of subdomain-1 of actin 2. In this ternary complex, S(1) bridges subdomain 2 of the actin 2 subunit. In support of this model, changes in subdomain 2 of actin such as covalent modification of lysine 61 (25, 37, 38) or subtilisin cleavage between Met and Gly(39) all affect S(1) binding to F-actin. Similarly, limited proteolytic digestion studies (30, 40) showed that S(1) binding to G-actin induces changes in subdomain 2 (loop 38-69) of G-actin, which supports a structure of the G(2)S complex similar to the F-actin-S(1) complex.

In a recent report (41) equilibrium and kinetic data of S(1) binding to F-actin were interpreted in terms of two different rigor complexes, actin-S(1) and (actin)(2)-S(1), depending on the F-actin/S(1) ratio. More work is needed, to correlate the possible different F-actin-S(1) complexes to the GS and G(2)S complexes studied in the present work. An important difference between G-actin-S(1) and F-actin-S(1) interaction is that while G-actin monomers are dispersed in solution, actin subunits interact with each other in the filament, so that at all F-actin/S(1) ratios, the two adjacent actin subunits which may form the regular (actin)(2)-myosin interface are preassociated to interact with S(1) in a ternary rigor complex. Hence we suggest that the G(2)S structure is a good model of the F-actin-S(1) interface.

Both GS and G(2)S complexes are dissociated by ATP and ADP with relative efficiencies that compare well with F-actin-S(1) records. Kinetic data show that the mechanism of the ATP or ADP-induced dissociation involves binding of nucleotide to the G-actin-S(1) complexes followed by an isomerization of the (G-actin)-S(1)-nucleotide complexes which kinetically limits the dissociation of S(1)-nucleotide from G-actin. Again the change in pyrene fluorescence monitors the isomerization step rather than the dissociation of S(1). All these features are qualitatively similar to the mechanism of ATP-induced dissociation of S(1) from F-actin (42, 43) at low ionic strength, which emphasizes that the G-actin-S(1) interaction is functional in terms of regulation by nucleotides.

One of the interests of the G-actin-S(1) complexes resides in the opportunity to study the regulation of the interaction of the myosin head with one actin molecule or two actin molecules separately, since experimental conditions can be designed under which essentially either GS or G(2)S exist in solution. The different behavior of GS and G(2)S in response to ATP and ADP is striking. Both the high affinity derived from the amplitude data and the dependence of k on ATP concentration show that G(2)S is more similar to F-actin-S(1) in solution than GS, which supports the view of the ternary G(2)S complex as the minimum actomyosin unit. On the other hand, the fact that the myosin head can interact with either one or two actin subunits with different mechanical properties (i.e. different use of ATP) should be considered in cross-bridge function. Whether a change in the actin/S(1) binding stoichiometry is involved in the cross-bridge ATPase cycle and in force development requires further investigation. Nevertheless, recent measurements of the mechanics of single myosin (44) molecules have shown that the time during which myosin interacts with the actin filament displays different ATP concentration dependences at low load and high load and varies with the load when ATP is not limiting (see (45) for a recent review). Our data, showing a different ATP dependence for GS and G(2)S, suggest that the different actin-myosin interaction times may correlate with different orientations of the myosin head, for instance the myosin head would interact predominantly with 2 actin subunits at high load (high ATP affinity) and with 1 actin subunit at low load (low ATP affinity). These possible different orientations would be compatible with the noticed (46) large variability in the measurements of displacement and forces of single myosin molecules along the actin filament. In solution studies of F-actin-S(1) interaction, it has also been reported (47) that the binding of ATP to F-actin-S(1) varied with the actin/S(1) molar ratio.

The interaction of G-actin with S(1) appears potently inhibited by DNaseI. At variance with previous reports (29, 30, 31) the present results can be quantitatively accounted for by a simple mutual exclusion binding scheme of S(1) and DNaseI to G-actin, the affinity of either S(1) or DNaseI for G-actin being reduced about 10^3-fold in the potential ternary complex. In other words, in the presence of micromolar amounts of G-actin and DNaseI, a concentration as high as 100 µM S(1) would be necessary to observe a ternary DNaseI-G-actin-S(1) complex. Our conclusion is consistent with the inhibition by DNaseI of the S(1)-induced polymerization of G-actin, as well as with previous results indicating competition between DNaseI and S(1) for binding to actin (37, 38, 39, 48, 49) , and with data showing a decrease in binding strength of DNase I by at least one order of magnitude to the covalent MBS actin-S(1) complex(29) .

In conclusion, the interaction of G-actin with the myosin head may be a useful system to investigate the change in actin-myosin interaction during the ATPase cycle, and the G(2)S complex is a good model of the F-actin-S(1) interface.


Appendix

Kinetic Model for Interaction of S(1) with G-actin

According to , the observed fluorescence is described by the following equation:

In , f(0) represents the intrinsic pyrene fluorescence in G-actin, GS, and G(2)S complexes, while f(1) and f(2) represent the intrinsic pyrene fluorescences of actin in GS* and G(2)S*, respectively. As discussed previously(12) , f(2) represents the average fluorescence of the two actins in G(2)S*, but the specific fluorescences of each actin molecule in G(2)S*, f`(2) and f``(2), are unknown, and f(2) = 1/2(f`(2) + f``(2)).

The amplitude DeltaF of the fluorescence change is the difference between the fluorescences observed at equilibrium (t) and at time 0:

where

Quantitative analysis of the amplitude data ( Fig. 2and Fig. 3) is complex since a total of six parameters (four equilibrium dissociation constants K(1), K(2), K(3), and K(4) and two fluorescence parameters f(1) and f(2)) are involved. Theoretical curves describing the amplitude of the fluorescence change at different total G-actin and S(1) concentrations were generated by computer using and and the following mass conservation equations:

The concentration of free G-actin was the solution of the quadratic as a function of [S]. A series of incremented values of [S] were generated, from which the corresponding values of [G], [S], and the concentrations of all G-actin-S(1) complexes were calculated. To simplify the search for equilibrium parameter values, useful hints were derived from visual inspection of the data obtained under extreme conditions where only binary or ternary complexes exist, as follows. In Fig. 4, the change in k with [S(1)] in a range of high values (geq4 µM) of [S(1)] reflects the formation of GS and GS* complexes essentially, in which only the constants K(1) and K(3) are involved. Examination of the data suggests that K(1) is of the order of 1 µM. The value of k reached at saturation by S(1) represents k + k = 210 ± 20s. The shape of the curve kversus [S(1)] above 5 µM S(1) imposed a range of possible values of k between 5 and 30 s, corresponding to a range of K(3) of 6-40. Note that if K(1) was much larger than 1 µM, the value of K(3) 0.1 would impose K(1)K(3) 0.1 µM, a value too high to be compatible with the dependences of the amplitudes on S(1) and G-actin, which impose K(1)K(3) < 0.1 µM. Similarly, the dependence of k on G-actin (Fig. 5) in a range of high concentrations, and the shape of the titration curves (amplitude data, Fig. 2and Fig. 3) required that the value of K(2) be at most 0.1 µM, and K(4) between 3 and 6. With these constraints the best fit to the titration curves (amplitude data, Fig. 2and Fig. 3) was sought varying K(3), K(4), f(1), and f(2). In parallel, the time courses of the change in fluorescence were simulated, in a large range of G-actin and S(1) concentrations, using KINSIM, searching for the best fit to experimental traces by refining the values of k, k, k, k keeping with the imposed constraints. The resulting best values of equilibrium and rate parameters are given in Table 1. From these values, it appears that a large increase in affinity of G-actin for S(1) is linked to the isomerization steps k and k, and the fluorescent complexes GS* and G(2)S* are largely predominant over their non-fluorescent counterparts. Setting f(0) = 1 by convention, a value of f(1) = f(2) = 1.6 ± 0.1 was found to fit all data adequately. In a search for the best fit to the data, it was found that fluctuations of more than 20% in the values of each of the parameters taken separately yielded sets of theoretical curves that could not fit the set of amplitude data adequately even upon varying the values of other parameters in a compensatory fashion. In other words, the two complementary experimental saturation curves in Fig. 2and Fig. 3together impose serious constraints in the choice of equilibrium parameters, and we trust that the proposed set of values is reasonably robust. The solid lines in Fig. 2and Fig. 3are theoretical curves calculated within using the values of K(1), K(2), K(3), K(4), f(1), and f(2) given in Table 1for the two isomers of S(1).

Finally the distribution of the different actin-S(1) complexes present in solution in a broad range of actin and S(1) concentrations, calculated within and consistent with all data shown, is displayed in Fig. 10.


FOOTNOTES

*
This work was supported in part by the Association pour la Recherche contre le Cancer (ARC), the Ligue Nationale Française contre le Cancer, and the Association Française contre les Myopathies (AFM). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Present address: INSERM U128, CNRS, 34033 Montpellier, France.

To whom correspondence should be addressed. Tel.: 33 1 69 82 34 65; Fax: 33 1 69 82 31 29; carlier{at}pegase.enzy.cnrs-gif.fr.

(^1)
The abbreviation used is: MOPS, 4-morpholinepropanesulfonic acid.


ACKNOWLEDGEMENTS

We thank Dominique Didry for technical help in the preparation of actin and myosin subfragment-1.


REFERENCES

  1. Lymn, R. W., and Taylor, E. W. (1971) Biochemistry 10, 4617-4624 [Medline] [Order article via Infotrieve]
  2. Geeves, M. A., Goody, R. S., and Gutfreund, H. (1984) J. Mus. Res. Cell Motil. 5, 351-361 [Medline] [Order article via Infotrieve]
  3. Eisenberg, E., and Hill, T. L. (1985) Science 227, 999-1006 [Medline] [Order article via Infotrieve]
  4. Hibberd, M. G., and Trentham, D. R. (1986) Annu. Rev. Biophys. Biophys. Chem. 15, 119-161 [CrossRef][Medline] [Order article via Infotrieve]
  5. Goldman, Y. E. (1987) Annu. Rev. Physiol. 49, 637-654 [CrossRef][Medline] [Order article via Infotrieve]
  6. Kabsch, W., Mannherz, H. C., Suck, D., Pai, E., and Holmes, K. C. (1990) Nature 347, 37-44 [CrossRef][Medline] [Order article via Infotrieve]
  7. Rayment, I., Rypniewski, W. R., Schmidt-Base, K., Smith, R., Tomchick, D. R., Benning, M. M., Winkelmann, D. A., Weisenberg, G., and Holden, H. M. (1993) Science 261, 50-57 [Medline] [Order article via Infotrieve]
  8. Milligan, R. A., Whittaker, M., and Safer, D. (1990) Nature 348, 217-221 [CrossRef][Medline] [Order article via Infotrieve]
  9. Rayment, I., Holden, H. M., Whittaker, M., Yohn, C. B., Lorenz, M., Holmes, K. C., and Milligan, R. A. (1993) Science 261, 58-65 [Medline] [Order article via Infotrieve]
  10. Valentin-Ranc, C., Combeau, C., Carlier, M.-F., and Pantaloni, D. (1991) J. Biol. Chem. 266, 17872-17879 [Abstract/Free Full Text]
  11. Combeau, C., Didry, D., and Carlier, M.-F. (1992) J. Biol. Chem. 267, 14038-14046 [Abstract/Free Full Text]
  12. Valentin-Ranc, C., and Carlier, M.-F. (1992) J. Biol. Chem. 267, 21543-21550 [Abstract/Free Full Text]
  13. Coates, J. A., Criddle, A. H., and Geeves, M. A. (1985) Biochem. J. 232, 351-356 [Medline] [Order article via Infotrieve]
  14. Geeves, M. A., Jeffries, T. E., and Millar, N. C. (1986) Biochemistry 25, 8454-8458 [Medline] [Order article via Infotrieve]
  15. Taylor, E. W. (1991) J. Biol. Chem. 266, 294-302 [Abstract/Free Full Text]
  16. Spudich, J. A., and Watt, S. (1971) J. Biol. Chem. 246, 4866-4871 [Abstract/Free Full Text]
  17. Eisenberg, E., and Kielley, W. W. (1974) J. Biol. Chem. 249, 4742-4748 [Abstract/Free Full Text]
  18. McLean-Fletcher, S., and Pollard, T. D. (1980) Biochem. Biophys. Res. Commun. 96, 18-27 [Medline] [Order article via Infotrieve]
  19. Kouyama, T., and Mihashi, T. (1981) Eur. J. Biochem. 114, 33-38 [Abstract]
  20. Mockrin, S., and Korn, E. D. (1980) Biochemistry 19, 5358-5363
  21. Offer, G., Moos, C., and Starr, R. (1973) J. Mol. Biol. 74, 653-676 [Medline] [Order article via Infotrieve]
  22. Weeds, A., and Pope, B. (1977) J. Mol. Biol. 111, 129-157 [Medline] [Order article via Infotrieve]
  23. Price, P. A., Liu, T. Y., Stein, W. H., and Moore, S. (1969) J. Biol. Chem. 244, 917-923 [Medline] [Order article via Infotrieve]
  24. Blikstad, I., Markey, F., Carlsson, L., Persson, T., and Lindberg, U. (1978) Cell 15, 935-943 [Medline] [Order article via Infotrieve]
  25. Combeau, C., and Carlier, M.-F. (1992) Biochemistry 31, 300-309 [Medline] [Order article via Infotrieve]
  26. Criddle, A. H., Geeves, M. A., and Jeffries, T. (1985) Biochem. J. 232, 343-349 [Medline] [Order article via Infotrieve]
  27. Mannherz, H. G., Goody, R. S., Konrad, M., and Nowak, E. (1980) Eur. J. Biochem. 104, 367-379 [Abstract]
  28. Pinder, J. C., and Gratzer, W. B. (1982) Biochemistry 21, 4886-4890 [Medline] [Order article via Infotrieve]
  29. Bettache, N., Bertrand, R., and Kassab, R. (1990) Biochemistry 29, 9085-9091 [Medline] [Order article via Infotrieve]
  30. Chen, T., Haigentz, M., Jr., and Reisler, E. (1992) Biochemistry 31, 2941-2946 [Medline] [Order article via Infotrieve]
  31. Lheureux, K., Forné, T., and Chaussepied, P. (1993) Biochemistry 32, 10005-10014 [Medline] [Order article via Infotrieve]
  32. Arata, T. (1991) J. Biochem. (Tokyo) 109, 335-340 [Abstract]
  33. Chen, T., and Reisler, E. (1991) Biochemistry 30, 4546-4552 [Medline] [Order article via Infotrieve]
  34. Mornet, D., Bertrand, R., Pantel, P., Audemard, E., and Kassab, R. (1981) Nature 292, 301-306 [Medline] [Order article via Infotrieve]
  35. Méjean, C., Boyer, M., Labbe, J.-P., Morlier, L., Benyamin, Y., and Roustan, C. (1987) Biochem. J. 244, 571-577 [Medline] [Order article via Infotrieve]
  36. Labbe, J., Méjean, C., Benyamin, Y., and Roustan, C. (1990) Biochem. J. 271, 407-413 [Medline] [Order article via Infotrieve]
  37. Miki, M., Barden, J. A., DosRemedios, C. G., Phillip, L., and Hambly, B. D. (1987) Eur. J. Biochem. 165, 125-130 [Abstract]
  38. Barden, J. A., and Phillip, L. (1990) Biochemistry 29, 1348-1354 [Medline] [Order article via Infotrieve]
  39. Schwyter, D., Phillips, M., and Reisler, E. (1989) Biochemistry 28, 5889-5895 [Medline] [Order article via Infotrieve]
  40. Fievez, S., and Carlier, M.-F. (1993) FEBS Lett. 316, 186-190 [CrossRef][Medline] [Order article via Infotrieve]
  41. Andreev, O. A., Andreeva, A. L., Markin, V. S., and Borejdo, J. (1993) Biochemistry 32, 12046-12053 [Medline] [Order article via Infotrieve]
  42. Millar, N. C., and Geeves, M. A. (1983) FEBS Lett. 160, 141-148 [CrossRef][Medline] [Order article via Infotrieve]
  43. Geeves, M. A., Jeffries, T. E., and Millar, N. C. (1986) Biochemistry 25, 8454-8458 [Medline] [Order article via Infotrieve]
  44. Finer, J. T., Simmons, R. M., and Spudich, J. A. (1994) Nature 368, 113-119 [CrossRef][Medline] [Order article via Infotrieve]
  45. Jiang, M. Y., and Sheetz, M. P. (1994) BioEssays 16, 531-532 [Medline] [Order article via Infotrieve]
  46. Howard, J. (1994) Nature 368, 98-99 [Medline] [Order article via Infotrieve]
  47. Tesi, C., Travers, F., and Barman, T. (1990) Biochemistry 29, 1846-1852 [Medline] [Order article via Infotrieve]
  48. Hitchcock, S. E., Carlsson, L., and Lindberg, U. (1976) Cell 7, 531-542 [Medline] [Order article via Infotrieve]
  49. Mannherz, H. G., Barrington Leigh, J., Leberman, R., and Pfrang, H. (1975) FEBS Lett. 60, 34-38 [CrossRef][Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.