(Received for publication, December 8, 1995; and in revised form, February 28, 1996)
From the
The actin-activated Mg-ATPase activity of Acanthamoeba myosin II minifilaments is dependent both on
Mg
concentration and on the state of phosphorylation
of three serine sites at the C-terminal end of the heavy chains.
Previous electric birefringence experiments on minifilaments showed a
large dependence of signal amplitude on the phosphorylation state and
Mg
concentration, consistent with large changes in
filament flexibility. These observations suggested that minifilament
stiffness was important for function. We now report that the binding of
nucleotides to dephosphorylated minifilaments at Mg
concentrations needed for optimal activity increases the
flexibility by about 10-fold, as inferred from the birefringence signal
amplitude increase. An increase in flexibility with nucleotide binding
is not observed for dephosphorylated minifilaments at lower
Mg
concentrations or for phosphorylated minifilaments
at any Mg
concentration examined. The relaxation
times for minifilament rotations that are sensitive to the conformation
myosin heads are also observed to depend on phosphorylation,
Mg
concentration, and nucleotide binding. These
latter experiments indicate that the actin-activated
Mg
-ATPase activity of Acanthamoeba myosin II
correlates with both changes in myosin head conformation and the
ability of minifilaments to cycle between stiff and flexible
conformations coupled to nucleotide binding and release.
The heavy chain component of the multiple members of the myosin
superfamily share a highly homologous head (but with sufficient
sequence differences to allow their classification into 10 or 11
families) connected to highly variable tails(1) . The ATP- and
actin-binding sites, actin-activated Mg-ATPase
activity, and in vitro motility activity all reside in the
conserved head domain. The tail is generally thought to determine the
supramolecular organization of the myosins and their associations with
specific cell structures and organelles. For example, the two
N-terminal heads of type II myosin heavy chains are attached to a long
rodlike C-terminal tail that self-associates into an
-helical
coiled-coil and subsequently assemble into a bipolar filament (the
heavy chains of other myosin classes either remain monomeric or form
dimers but do not form filaments).
The heavy chains of most class II
myosins can be proteolytically cleaved to light meromyosin (LMM), ()the C-terminal end of the
-helical, coiled-coil rod,
and heavy meromyosin (HMM), the N-terminal portion of the
-helical, coiled-coil rod with its two attached globular
heads(2) . LMM retains the filament-forming properties of
native myosin II, and HMM retains all of the catalytic activity and in vitro motility activity. HMM can be further proteolyzed to
two monomers of subfragment 1 (S1), a single globular head with an
-helical C-terminal tail, and subfragment 2 (S2), the
-helical, coiled-coil portion of HMM. The individual S1 fragments
retain most of the properties of HMM.
Class II myosins contain two
pairs of light chains, essential light chains (ELC) and regulatory
light chains (RLC), with one of each pair attached to the helical tail
of each S1 domain(3) . Some (molluscan) native type II myosins
are activated by direct binding of Ca to the ELC and
others (smooth muscle, vertebrate nonmuscle, and Dictyostelium) by phosphorylation of the RLC (4) . The
light chains stabilize the helical tail of S1(3, 5) ,
and removal of the light chains or the helical tail of S1 greatly
reduces the ability of S1 to move actin filaments in an in vitro motility assay(6) , but neither the light chains nor the
helical tail of S1 are necessary for maximal actin-activated
Mg
-ATPase activity of S1. These data are consistent
with the most recent structural model of the contractile cycle derived
from x-ray crystallography of S1; conformational changes in the
globular portion of S1 resulting from the binding and hydrolysis of ATP
are transmitted through the ELC and generate a rotational movement of
the helical tail of S1(7) . This model was supported by recent
experiments that demonstrate tilting of the light chain region of the
myosin head during muscle contraction(8) .
It seems likely,
however, that the head and tail domains of class II myosins are not as
functionally independent as the foregoing brief summary might imply.
For example, the myosin superfamily can be grouped into essentially the
same classes based on the overall structure of their tails(1) ,
suggesting co-evolution of the head and tail domains that would
presumably result from their functional interactions. Moreover, the
actin-activated Mg-ATPase activities of S1 from both
ELC-regulated and RLC-regulated muscle myosins are unregulated (whereas
HMM is regulated), suggesting that at least a portion of the tails is
necessary for the appropriate coupling of S1 head and light chain
conformations(4) .
Acanthamoeba myosin II provides
a striking example of the functional interaction of the head and tail
regions of a myosin II; both the actin-activated
Mg-ATPase activity and the in vitro motility
activity of minifilaments of Acanthamoeba myosin II are
inactivated by phosphorylation of 3 serine residues in a short (29
amino acids), nonhelical tail piece at the C-terminal tip of the
-helical, coiled-coil rod(9, 10) . Extensive
experimental data (11, 12, 13) show that the
activity of each molecule in the filament depends not on its own
phosphorylation state but on the level of phosphorylation of the
filament as a whole. This led to the hypothesis that the state of
phosphorylation at the tip of the tail affects the conformation of the
hinge region at the HMM-LMM junction in the tails of adjacent molecules
in the minifilament and, thereby, the interactions with F-actin of the
S1 heads at the ends of the HMM arms. Electric birefringence studies on
chymotrysin-treated Acanthamoeba myosin II parallel dimers (14) indeed indicated that the tip of the tail of one monomer
was in close proximity to the hinge region of the other.
Further
electric birefringence experiments showed that the signals from
minifilaments are fundamentally different from the monomer and parallel
dimer signals; they were interpreted as being due to a coupling of
internal motions, bending or flexing, and alignment in an electric
field(15) . Furthermore, these studies indicated that the
stiffness of Acanthamoeba myosin II minifilaments is
correlated with their actin-activated Mg-ATPase
activity. Filaments of dephosphorylated myosin II stiffen dramatically
between 1 and 4 mM Mg
. The optimal
concentration for both catalytic and in vitro motility
activities for dephosphorylated minifilaments occurs at about 4-5
mM Mg
. In contrast, filaments of
phosphorylated myosin, which are inactive at all Mg
concentrations, remain flexible even at 4 mM Mg
.
Recent experiments (16) showing
that binding of ATP to Acanthamoeba myosin II enhances the
rate of papain cleavage of the heavy chain of monomers in the region
corresponding to that which interacts with the ELC in chicken myosin II (3) are consistent with the proposal (7) that ATP
induces conformational changes in S1. However, ATP also promotes papain
cleavage within the C-terminal tail of phosphorylated
minifilaments(16) ; this was not anticipated. We now
investigate the effects of nucleotides on the flexibility of
phosphorylated and dephosphorylated minifilaments. The electric
birefringence signal amplitude of dephosphorylated minifilaments
indicates that an order of magnitude increase in flexibility
accompanies nucleotide binding at 4-5 mM Mg but not at lower Mg
concentrations.
Phosphorylated minifilaments remain flexible at all Mg
concentrations between 1 and 5 mM in the presence of
nucleotide. In addition, over the same range of experimental
conditions, differences in either the structure or orientation of S1
heads can be inferred from differences in relaxation times of the
spinning rotational motions. These results demonstrate that a complex
interplay among C-terminal phosphorylation state, Mg
concentration, and nucleotide binding determines minifilament
flexibility and conformation and, ultimately, regulates enzymatic
activity.
Myosin II stock solutions used in the
electric birefringence experiments were extensively dialyzed against
2.5 mM imidazole (pH 7.0), 5 mM KCl, 1 mM DTT, and 50% sucrose. Final protein concentrations were 1-2
mg/ml. Samples used in electric birefringence experiments were first
adjusted to 1 mM KCl, 2 mM imidazole (pH 7.4), 1
mM DTT, and 5% sucrose. The MgCl was added in
aliquots, with mixing, to obtain the final Mg
concentration. Although no difference in signal was observed for
adding nucleotide before Mg
, nucleotide was typically
added last.
where is the angle between polarizers relative to the
crossed orientation (typically 1 °C), I
is
the output voltage from the photodiode and amplifier at the angle
(typically 1000 mV), and
I is the change in output
voltage due to the birefringence of the oriented sample.
The
relaxation times of the two birefringence components were extracted
from the decay data by several methods. First, the relaxation time of
the slow, negative birefringence component was determined directly from
the slope of the semilog plot of signal intensity versus time
at long times (>150 µs after the end of the pulse) as
illustrated in Fig. 5. The relaxation time of the fast, positive
birefringence component was then determined from the slopes of semilog
plots of signal versus time after subtracting the contribution
of the slow component, as shown in Fig. 6. The entire decay
curve was also fit to a double exponential function, and best fitting
values for and
were
determined simultaneously. Finally, the birefringence decay was also
analyzed using the Fortran program Contin(23, 24) ,
which gives a spectrum of relaxation times from an inverse Laplace
transform of the data. The average relaxation times determined by the
three methods did not differ significantly. The slowly decaying
component can be well fit by a single exponential. We are less certain
that the positive birefringence component has only a single relaxation
time. We report fast decay times as average values,
<
>, to emphasize this point.
Figure 5:
The
effect of Mg and nucleotide on the decay kinetics of
the slow, negative birefringence component of dephosphorylated
minifilaments. Semilog plots of the optical signal, normalized for
field strength and protein concentration, versus time, with t = 0 set at the end of the applied pulse, are shown
for minifilaments in 3, 4.5, and 4.5 mM Mg
with 5 µM AMPPNP. The dashed lines are the
best linear fits to the data. The arrows indicate the ordinate
appropriate for each decay curve. The signal amplitudes for
minifilaments in 3 mM Mg
(left
ordinate) and in 4.5 mM Mg
with 5
µM AMPPNP (right ordinate) were about equal; the
signal intensity of minifilaments in 4.5 mM Mg
was about a factor of 10 smaller (left ordinate). Other
solution conditions were as described in the legend to Fig. 2.
Figure 6:
Effects of Mg and
nucleotide on the decay kinetics of the fast, positive birefringence
component for dephosphorylated minifilaments. Semilog plots for the
birefringence decay of the fast component signal are shown for 3 mM Mg
and for 4.5 mM Mg
with 5 µM ADP to illustrate the large changes in
relaxation time that accompany nucleotide binding. The total signal
amplitudes are closely similar for the two conditions shown. The
contribution of the slow, negative birefringence component was removed
by subtracting the single exponential determined from the best linear
fits to the long time data as shown in Fig. 5. The dashed
lines are the best linear fits to the data. Other solution
conditions were as described in the legend to Fig. 2.
Figure 2:
The effect of added ATP on the electric
birefringence signal amplitude for dephosphorylated minifilaments in 5
mM Mg at 20 °C. The optical signal is
shown normalized for field strength and protein concentration,
/cE
, as in Fig. 1. The start and end
of the 140-µs long electric field pulse are indicated by the arrows. The increase in signal amplitude was about 10-fold
with ATP binding. In addition to the Mg
, the samples
also contained 1 mM KCl, 2 mM imidazole (pH 7.4), 1
mM DTT, 5% sucrose, and 40-50 µg/ml protein. The
added ATP concentration was 250
µM.
Figure 1:
A typical electric birefringence signal
for minifilaments of dephosphorylated Acanthamoeba myosin II.
Minifilaments at 40 µg/ml were in 2 mM Mg, 1 mM KCl, 2 mM imidazole
(pH 7.4), 1 mM DTT, and 5% sucrose at 20 °C. The optical
signal, shown normalized for field strength and myosin concentration
and plotted as a function of time, is due to the orientation of
minifilaments in an applied electric field, E, with the long
filament axis aligned perpendicularly to the field. The square wave
electric field pulse starts at 180 µs (relative to the start of
data acquisition) and ends at 320 µs, as indicated by the arrows. The optical signal of both the build-up (with field
on) and the decay (after field is removed) is the sum of at least two
components, one with positive birefringence that relaxes much faster
than the component with negative birefringence that dominates the
signal amplitude. The fast, positive birefringence component is due to
a spinning rotation about the minifilament long axis, illustrated in
the lower left hand cartoon of a tetrameric, bipolar filament
with two HMM arms (with S1 globular heads depicted as spheres at the
ends of the S2 rods) extending in opposite directions from each end of
a filament comprising four LMM segments. The more slowly relaxing,
negative birefringence component is due to an end-over-end tumbling
rotation of the minifilament, illustrated in the lower right.
Each component is characterized by an amplitude, A, and a
relaxation time,
. The overall signal amplitude, A
, and the ratio of the birefringence change
at the maximum in the rise,
A
, to the
birefringence change at the minimum in the decay,
A
, are predicted to depend on the
flexibility of the minifilaments.
As previously
noted, a major experimental limitation was a field-induced,
time-dependent aggregation of minifilaments that occurred at
Mg concentrations above about 4-5 mM.
High protein concentrations, extended signal averaging, long field
pulses, or high field strengths led first to a slowly decaying tail in
the birefringence signal and eventually to visibly clouded solutions.
This aggregation was less of a problem at pH 7.4 used in these
experiments than at pH 7.0, which we used previously (15) . The
decrease in signal amplitude for dephosphorylated minifilaments,
however, requires higher Mg
concentrations at pH 7.4
than at 7.0. The previously reported signal amplitudes at 4 mM Mg
at pH 7.0 require 4.5-5 mM at
pH 7.4, depending on the specific myosin preparation. To avoid
aggregation but also to achieve maximal signal amplitude and signal to
noise ratio standard conditions for the experiments reported here were
a protein concentration of about 40 µg/ml, a field strength E
1.2 kV/cm, and a voltage pulse length of 140 µs. Optical
signals from a single sample were averaged over 32 pulses.
Occasionally, much longer pulse lengths (
800 µs) were used to
ensure that more slowly relaxing birefringence components were not
present. Signal amplitudes were also measured as a function of field
strength, E, up to about 2 kV/cm to verify that
scales
with E
. Signal amplitudes were also observed to
vary linearly with protein concentration up to about 100 µg/ml.
Compared with this complicated set of signals from minifilaments, the electric birefringence signals from Acanthamoeba myosin II monomers and parallel dimers (14) are straightforward and provide a basis for understanding the minifilament signals. A large permanent dipole from the distribution of charged amino acids (25) extends over the HMM region of Acanthamoeba myosin II. The alignment of this permanent dipole in an electric field results in positive birefringence signals for monomers and parallel dimers, indicating that molecules are orienting with their long axes parallel to the field. Qualitatively similar positive birefringence signals are seen for skeletal muscle myosin HMM and S1 heads(26, 27) . Unperturbed minifilaments, however, have no net dipole moment due to the symmetry of the structure. A dipole moment can still be induced if the HMM arms can bend or flex in the direction of the applied field(28) . An internal flexibility of synthetic muscle myosin filaments was observed previously using spectroscopically labeled S1 heads(29, 30, 31) .
The dipole moment
resulting from the bending or flexing motions of the HMM can couple
through the applied field to the two fundamental rotational modes of
the minifilament illustrated in Fig. 1: (i) the spinning
rotation about the long axis and (ii) the end-over-end tumbling of the
long axis, which correspond to the fast and slow relaxation kinetics of
the positive and negative birefringence components, respectively. For
average orientations of the HMM arms that are predominately parallel to
the minifilament long axis and for small bending perturbations (low
field strengths), the electric field induced dipole moment is predicted
to be perpendicular to the long axis. The negative birefringence of the
slowly relaxing component results from this unusual perpendicular
orientation of the dipole and optical axes. The relaxation time of this
component, , is dependent on the distribution of
mass along the minifilament axis. In practice, it is most sensitive to
the number of monomers in the minifilament and the spacings between
them. The positive birefringence signal is due to a component of the
optical anisotropy that is perpendicular to the long axis (parallel to
the dipole axis). We previously estimated, for example, that a 20 °
angle between the HMM axis and the minifilament axis is sufficient to
account for the observed ratio of component amplitudes, A
/A
. The S2 rods of HMM
are expected to contribute more substantially to the optical signal
than the globular S1 heads(26) . The decay time of the spinning
rotation that relaxes this positive birefringence component,
<
>, depends on the distribution of all mass
perpendicular to the long axis.
The magnitude of the total
birefringence signal is a measure of the bending force constant or the
stiffness resisting internal HMM motions. Additionally, because the
internal motions that create the net dipole moment couple with the
rotations of the minifilament only in the rise part of the
birefringence curve the ratio
-A
/
A
(see Fig. 1) is also expected to vary with the kinetics of
the bending or flexing and thus is also connected to the stiffness.
The direction and magnitude of the
effect of ATP on signal amplitude varied with the Mg concentration and with myosin phosphorylation, as shown in Fig. 3. At 1 and 2 mM Mg
, the total
signal amplitude of dephosphorylated minifilaments decreased somewhat
with added ATP (filled symbols), whereas above about 3 mM Mg
, ATP increased the total signal amplitude.
This reversal in the effect of ATP reflects the 20-25-fold
decrease in signal amplitude between 1 and 5 mM Mg
in the absence of ATP but comparatively constant amplitude in its
presence. In contrast, the birefringence signal amplitude of
phosphorylated minifilaments was not affected either by the
Mg
concentration, as observed
previously(15) , or by the presence of ATP (Fig. 3, open symbols).
Figure 3:
The effect of ATP on the electric
birefringence signal amplitude as a function of Mg
concentration. The total birefringence signal amplitude,
-A
, normalized by applied field strength
and protein concentration, after a 140-µs electric field pulse is
shown for both phosphorylated (open symbols) and
dephosphorylated (solid symbols) minifilaments. The ATP
concentrations were: 0 (circles), 50 (squares), and
250 µM (triangles). Other solution conditions
were as described in the legend to Fig. 2.
The nucleotide concentration dependence of
the amplitude of the birefringence signal for dephosphorylated myosin
II minifilaments at 4.5 mM Mg is shown in Fig. 4for ATP, ADP, and the nonhydrolyzable ATP analogue AMPPNP.
The protein concentration in these experiments was
0.1 µM in myosin II monomers or
0.2 µM in S1 heads. The
lowest concentration of AMPPNP used, 0.4 µM, was
sufficient to achieve essentially the maximal increase in signal
amplitude (Fig. 4, squares). Even when the protein
concentration was increased to
0.35 µM S1, an AMPPNP
concentration of 0.4 µM was still sufficient to obtain the
maximal effect, i.e., even when the concentration of heads and
AMPPNP were approximately equal (data not shown). Similarly, the
addition of an approximately equimolar concentration of AMPPNP was
sufficient to obtain the maximum decrease in the amplitude of the
birefringence signal at 2 mM Mg
(data not
shown). These data indicate that the K
for AMPPNP
binding is smaller than 0.2-0.4 µM, which is
consistent with the K
of
0.09 µM that was estimated by differential scanning calorimetry. (
)
Figure 4:
The dependence of the electric
birefringence signal amplitude for dephosphorylated minifilaments at
4.5 mM Mg on nucleotide concentration. The
field strength and protein concentration normalized optical signal
amplitude at 20 °C is plotted as a function of nucleotide
concentration for ATP (
), ADP (
), and AMPPNP (
). The inset shows signal amplitudes for nucleotide concentrations up
to 50 µM. Other solution conditions were as described in
the legend to Fig. 2. The 40-50-µg/ml protein
concentration corresponds to
0.2 µM S1.
In contrast to the stoichiometric effect of AMPPNP, the
increase in birefringence signal amplitude with added ATP or ADP was
more consistent with a titration of binding sites between 0 and 4
µM nucleotide (Fig. 4). No further increase in
signal amplitude was observed between 5 and 50 µM ADP or
ATP (Fig. 4, inset). Consistent with this behavior,
increasing the myosin concentration from 0.2 to
0.35
µM S1 significantly decreased the observed signal
amplitude at 0.4 µM ADP and to a somewhat greater extent
also at 0.4 µM ATP, but it had a much smaller effect at
ADP and ATP concentrations higher than 1 µM (data not
shown).
From the birefringence data, the K for
ADP is
1-2 µM, a not unreasonable value because
ADP would be expected to bind to myosin more weakly than AMPPNP. In
contrast to expectations, however, ATP also appears to bind to the
myosin II minifilaments more weakly than AMPPNP. But, because the
actin-independent Mg
-ATPase activity of
dephosphorylated myosin II under the experimental conditions used for
the electric birefringence measurements is
5-10
10
s
, most of the ATP (for
initial concentrations up to
4 µM) was likely
hydrolyzed to ADP during the course of a typical 30-60-min
experiment. Thus, the high K
estimated for ATP
binding might actually have been for ADP binding instead.
In contrast, the relaxation time of the fast,
positive birefringence component was markedly dependent on both
Mg and nucleotide. Fig. 6illustrates the
3.5-fold difference in the average fast component relaxation
times, <
>, for dephosphorylated minifilaments
in 4.5 mM Mg
with 5 µM ADP (45
µs) and in 3 mM Mg
without added
nucleotide (12 µs). This difference in relaxation times was
observed even though the total signal amplitudes for these two
experimental conditions were comparable. Average fast component
relaxation times for other conditions of nucleotide and Mg
concentration are summarized in Table 1for
dephosphorylated and phosphorylated minifilaments. The increase in the
fast relaxation time for dephosphorylated minifilaments depended on the
nucleotide species bound, ranging from 40-45 µs with bound
ADP to 25-30 µs with AMPPNP and 20-25 µs with ATP.
In contrast, the average fast relaxation time for phosphorylated
minifilaments actually decreased somewhat with added ATP from
12-13 to 8-9 µs.
The absence of a significant change in the relaxation time of the
slow component, , upon the addition of nucleotides
to dephosphorylated minifilaments further indicates that nucleotide
binding does not cause a large change in the overall structure of the
minifilament. The basic structure, the number of monomers, the repeat
distance between monomers, and the length of the central bare zone, is
not significantly different in the presence or the absence of
nucleotide. In particular, there is no evidence for aggregation,
consistent with analytical ultracentrifugation data(16) .
Therefore, the order of magnitude of increase in total signal amplitude
with nucleotide binding at 4-5 mM Mg
most probably results from a substantial increase in the
flexibility of minifilaments. The decrease of
-
A
/
A
at 4-5 mM Mg
from 0.4 to 0.2 (Table 1) with added nucleotide is also consistent with an
increase in flexibility (28) . This ratio will depend on
stiffness through the kinetics of the bending motion and a value of
0.2 is characteristic of other conditions that give approximately
the same signal amplitude.
Because nucleotide binding can either
increase or decrease the signal amplitude, depending on the
Mg concentration, the effects of nucleotides and
Mg
are neither independent nor simply additive.
Rather, both likely regulate the flexibility of either the same or
tightly coupled sites. We previously suggested that the bending site
was at the HMM-LMM junction rather than at the S1-S2 junction or an
elastic flexing of the S2 rod itself based on the estimated bending
kinetics extracted from
-
A
/
A
.
This has been confirmed by more recent experiments on Acanthamoeba myosin II rods and rods in which the bend at the HMM-LMM junction
has been removed or modified by amino acid substitution. The results
demonstrate that the S1 heads are not necessary to obtain full signal
amplitude but that the ``native'' HMM-LMM bend is
essential(32) . The electric birefringence signal of native Acanthamoeba myosin II minifilaments almost certainly arises
from bending at the HMM-LMM junction. Preliminary data (
)also show that nucleotides have no effect on the electric
birefringence of the rods, as expected if nucleotides interact only
with S1 heads.
The bending force spring constant of monomers in the
minifilament can be estimated (28) from the magnitude of the
electric birefringence signal, assuming that the bending is Gaussian
and planar and occurs at the HMM-LMM junction and that the S1 heads
make little contribution to the birefringence compared with the
coiled-coil -helices of the rest of the molecule. For a bending
spring constant
in units of energy/radian
and thermal
energy kT, reduced spring constants, a
(=
/2kT), for dephosphorylated minifilaments
at 4-5 mM Mg
with and without bound
nucleotide are estimated as
15 and
300, respectively. These
values correspond to root mean square angular fluctuations,
&cjs3484;<
>, of about 10 and 2 °,
respectively.
The decay of the
positive birefringence component is through the spinning rotation of
the long axis (cf. Fig. 1) and therefore is sensitive
to the distribution of mass perpendicular to the filament axis, in
particular to the angle between S2 rods and the minifilament axis, the
angle between S1 and S2, the angle between S1 heads, and S1
conformation. These structural parameters are not as yet well enough
characterized to justify hydrodynamic calculations. Given the
comparative insensitivity of A/A
to experimental
conditions (Table 1), however, the changes in
<
> likely reflect changes either in S1
conformation or in the orientation of S1 relative to the minifilament
axis, rather than in S2. A change in isolated S1 head conformation with
nucleotide binding has been observed
previously(33, 34, 35) .
The substantial differences in
<> seen with nucleotide binding and myosin
phosphorylation are perhaps even more closely linked to enzymatic
activity, given the dependence on nucleotide species, i.e.,
actin-activated Mg
-ATPase activity may also correlate
with nucleotide-dependent conformational changes in S1 or at the S1-S2
junction. Because there is no direct correlation between
<
> and A
, the postulated
nucleotide-dependent changes in S1 structure (and enzymatic activity)
are not simply or directly coupled to changes in filament flexibility.
The nucleotide-dependent conformational changes in S1 inferred from the
electric birefringence are entirely consistent with the current model
for the cross-bridge cycle but nucleotide-dependent changes in the
conformation of the HMM-LMM junction introduce a novel concept that is
likely not to be specific to Acanthamoeba myosin II.