From the Department of Biochemistry, University of
Leicester, Leicester LE1 7RH and
The Central Laser Facility,
Rutherford Appleton Laboratory, Chilton, Didcot, OX11 0QX, United
Kingdom, and the § Department of Biochemistry,
Eötvös University, Budapest H-1088, Hungary
Received for publication, December 4, 2000, and in revised form, February 16, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Steady-state and time-resolved
fluorescence measurements were performed on a Dictyostelium
discoideum myosin II motor domain construct retaining a single
tryptophan residue at position 501, located on the relay loop. Other
tryptophan residues were mutated to phenylalanine. The Trp-501
residue showed a large enhancement in fluorescence in the presence of
ATP and a small quench in the presence of ADP as a result of perturbing
both the ground and excited state processes. Fluorescence lifetime and
quantum yield measurements indicated that at least three microstates of
Trp-501 were present in all nucleotide states examined, and these could not be assigned to a particular gross conformation of the motor domain.
Enhancement in emission intensity was associated with a reduction of
the contribution from a statically quenched component and an increase
in a component with a 5-ns lifetime, with little change in the
contribution from a 1-ns lifetime component. Anisotropy measurements
indicated that the Trp-501 side chain was relatively immobile in all
nucleotide states, and the fluorescence was effectively depolarized by
rotation of the whole motor domain with a correlation time on 50-70
ns. Overall these data suggest that the backbone of the relay loop
remains structured throughout the myosin ATPase cycle but that the
Trp-501 side chain experiences a different weighting in local
environments provided by surrounding residues as the adjacent converter
domain rolls around the relay loop.
Spectroscopic signals from tryptophan residues have long been used
as an empirical tool for distinguishing different nucleotide states of
the myosin ATPase cycle (1-3). Interest in this topic has been
rekindled by the elucidation of crystal structures of the myosin motor
domain in which the location and environments of the tryptophan
residues can be compared in several different states (4-8). Recently,
site-directed mutagenesis has been used to define the contributions of
individual tryptophan residues to the observed spectroscopic signals
(9-12). These studies were preceded by and have been supplemented with
chemical approaches to isolate the spectroscopy-sensitive residues
(13-15). There is now unanimous agreement that the conserved
tryptophan residue located on the relay loop of the lower 50 kDa
domain (skeletal myosin Trp-510, smooth myosin Trp-512, and
Dictyostelium Trp-501) is uniquely associated with the
fluorescence enhancement observed during ATP hydrolysis. The hydrolysis
step itself appears coupled to the open The kinetic mechanism of ATP hydrolysis by all myosins II appears
similar. Although the absolute values of the rate constants of
equivalent steps may differ among species, their ratios are such
that a similar distribution of nucleotide states is obtained during the
steady state. In the case of the D. discoideum motor domain
the mechanism may be summarized as in Reaction 1
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
closed transition,
identified crystallographically (16). It appears that the open
closed transition is actually responsible for the structural changes in
the vicinity of the spectroscopy-sensitive tryptophan residue, but the
hydrolysis reaction serves to pull the equilibrium toward the
predominant steady-state intermediate in the Dictyostelium
discoideum motor domain, M*·ADP·Pi or the
equivalent M**·ADP·Pi intermediate in vertebrate
skeletal myosins (12). The origin of the smaller fluorescence
enhancement seen on nucleotide binding to skeletal and smooth myosin is
less clear and is species-specific. In smooth muscle myosin the
enhancement induced by ADP binding appears to emanate from Trp-512
(10), whereas the equivalent residue in D. discoideum,
Trp-501, responds with a fluorescence quench (12). The fluorescence
enhancement associated with ADP binding to skeletal myosin may have its
origins in the nonconserved Trp-131 residue (15). In this work we
address further properties of the relay loop tryptophan of D. discoideum myosin motor domain, Trp-501.
where
represents a quenched state, and * is an enhanced state
of Trp-501 (12). States in parentheses are present at near negligible
concentrations under most conditions, but their existence is implied by
kinetic arguments. Under the usual conditions of assay, the
M*·ADP·Pi state predominates during the steady-state hydrolysis of ATP (which is equivalent to the
M**·ADP·Pi state of vertebrate myosins in which
other tryptophan residues may contribute to the overall fluorescence
enhancement (17, 18)). By adding the Pi analog
AlF
·ADP complex, a stable M*·ADP·AlF4
complex can be made which has an enhanced fluorescent state and
provides a useful analog of the M*·ADP·Pi state for
spectroscopic studies (12). Care is required, however, in relating a
particular analog state to particular ATPase intermediate because in
many cases these complexes exist as a mixture of conformations.
Fluorescence quenching studies with acrylamide and iodide indicate that
on binding nucleotide, Trp-501 (or equivalent) becomes slightly less
accessible to solvent and is protected further after hydrolysis to give
the long lived M*·ADP·Pi species (10, 12, 15). The
crystal structures are generally in accord with the spectroscopic
results in that, when resolved, Trp-501 (or equivalent) points toward
the converter domain, but it is not internalized completely and has
limited exposure to solvent (5, 6). Interestingly, in the case of the
Dictyostelium crystal structures, the main chain bearing
Trp-501 is usually unresolved in nucleotide states corresponding to the
open state (M·ADP,
M·ATPS),1 but an
exception is the dinitrophenyl aminoethyl diphosphate BeFx
complex (pdb 1D1A (19)). One possible explanation of the origin of the
fluorescence enhancement associated with the open
closed transition
is that the relay loop goes from a disordered to ordered state and
internalizes the Trp-501 side chain during the ATPase cycle, so
protecting it from solvent (H2O) quenching. We have
therefore carried out time-resolved fluorescence measurements on a
mutant containing a single tryptophan residue at Trp-501 to test this idea.
Overall we find that the Trp-501 residue is static on the nanosecond
time scale in all states (nucleotide and nucleotide-free) of the
Dictyostelium motor domain. The anisotropy value is similar for all complexes and decays slowly with a time consistent with rotational motion of the whole motor domain (50-100 ns). However, the
fluorescence intensity lifetime profiles and quantum yields suggest
that in any one nucleotide state, the Trp-501 residue senses at least
two or three different local environments on a time scale >5 ns to
<20 µs. The fluorescence enhancement arises from a shift in the
relative populations of these states that comprise both static and
dynamically quenched components. Although accessibility to solvent
water may contribute to quenching, interactions with nearby charged and
polar groups are likely to be dominant factors. These results are
compatible with the proposal that the relay loop remains structured
throughout the ATPase cycle and maintains its contact with the
converter domain (20) but that the environment of the tryptophan
residue is affected by local rearrangements of the side chain and
surrounding residues.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials--
The cloning and preparation of a D. discoideum myosin II M761 motor domain containing a single
tryptophan residue at position 501 was carried out as described
previously (10) and is based on the expression vector developed by
Manstein et al. (21). In this construct, denoted W501+, the
nonresponsive tryptophan residues at positions 36, 432, and 584 were
mutagenized to phenylalanine. A tryptophan-less construct, termed W,
was also made as a control in which all tryptophan residues were
changed to phenylalanine. Nucleotides, NATA and human serum albumin
(which contains a single buried tryptophan) were obtained from Sigma
Chemical Co. (Poole, U. K.)
Steady-state Fluorescence Measurements-- Steady-state fluorescence spectra and anisotropy were measured with an SLM 48000 spectro-fluorometer (SLM Instruments, Urbana, IL) using a 5 mm path length cell (101.034-QS, Hellma, Westcliff-on-Sea, U. K.). For spectral measurements, tryptophan was excited with a Hg-Xe lamp at 297 nm with 1-nm slits to minimize photobleaching and to reduce inner filter effects arising from high nucleotide concentrations. Quantum yields were determined using the comparative method taking the value for tryptophan of 0.13 (22). Emission spectra were corrected using the calibration protocol supplied by SLM Instruments. Absorption spectra were measured using a Pye Unicam SP8-100 spectrophotometer.
Steady-state anisotropies were measured using the T format method with
Glan-Thomson calcite prism polarizers. Tryptophan was excited at 295 nm
with 2-nm slits, and the vertical (parallel) emission was detected
through a monochromator set at 335 nm with 16-nm slits. The horizontal
(perpendicular) emission was detected through WG320 and UG11 filters
(Comar Instruments, Cambridge, U. K.). The anisotropy values
(A) were calculated from the intensity values of the
vertical and horizontal measurements with correction for the detector
G value (A = (Iv
G·Ih)/(Iv+2·G·Ih).
The G factor was calculated from the ratio of intensities
(Iv/Ih) when the excitation light was horizontally polarized. The gain on the
photomultipliers was adjusted so that the G factor was close
to 1. However, we noted that the SLM automated polarization routine
(48000 DOS version 2.1) gave erroneous values for the anisotropy
because the background signal was not subtracted from intensities
used to define the G value. Although this error may be
negligible for strongly fluorescing samples, it was a significant
factor in our measurements. For this reason we computed the anisotropy
values independently from the measured intensity values. We also
checked the anisotropy using the L format, so avoiding the use of the
emission monochromator that makes a significant contribution to the
G factor correction.
Transient fluorescence and absorption measurements were made using Applied Photophysics SX18MV stopped-flow apparatus with a xenon light source at 295 nm (2-4-nm slit width). Fluorescence emission was selected with WG320 and UG11 filters. All reactions were studied in a buffer comprising 40 mM NaCl, 20 mM TES, 1 mM MgCl2 at pH 7.5 at 20 °C and the reagent concentrations stated refer to the reaction chamber.
Time-resolved Fluorescence Measurements--
Time-resolved
fluorescence measurements were carried out using the time-correlated
single-photon counting method at the Rutherford Appleton Laboratory,
Central Laser Facility. Samples (5-25 µM) were excited
at 296 nm, achieved with a frequency-doubled, cavity-dumped, mode-locked synchronously pumped dye laser (Spectra-Physics model 3500)
operating at 4 MHz with a pulse width of 10 ps. The fluorescence emission was collected via a WG335/UG11 filter combination (WG320 was
used for tyrosine emission) using a MCP-PMT photomultiplier tube
(Hamamatsu, R3809U) with response time of 85 ps. The WG335 filter was
used in preference to the WG320 filter in these studies to minimize the
potential scattering artifact and tyrosine contribution to the signal.
Time-resolved anisotropy was measured through a film polarizer (Halbo
Optics, DPUV-25) in the emission channel, and the vertical and
horizontally polarized light was collected sequentially. The total
emission intensity for lifetime measurements was calculated by summing
Iv + 2·Ih. A wedge
depolarizer was placed in front of the photomultiplier tube, and the
G factor of the detector was confirmed as unity by
excitation with horizontally polarized light. The data were collected
in 1024 channels (width 40 ps) using the Becker and Hickl GmbH (Berlin)
single time-correlated single photon counting pc card module (SPC-700).
The standard functions of the single photon counting setup, such as
time-to-amplitude converter and constant fraction
discriminator, were computer-controlled with software supplied
with the SPC-700 module.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Trp-501 Fluorescence Enhancement and Quantum Yield-- The addition of ATP to the D. discoideum W501+ motor domain causes a large (30-90%) enhancement in tryptophan emission intensity (12). The percentage enhancement observed is critically dependent on the wavelength and band width of the excitation source and emission detector. Samples were excited at 295-297 nm to minimize the contribution from tyrosine fluorescence and inner filter effects on the addition of high adenine nucleotide concentrations. To obtain sufficient signal with xenon light sources, 2-4-nm slit widths were required, and particular care was required in analysis because of the extreme sensitivity to these extraneous contributions when operating at the edge of the excitation spectrum. The properties of the emission detector are also crucial because the enhancement in fluorescence is accompanied by about a 9-nm blue shift. For time-resolved measurements cut-off filters were used in the emission channel to maximize photon throughput, but this approach underestimates the percentage change in fluorescence compared with that determined from the integrated intensity of the complete corrected emission spectrum. On the other hand, spectral purity is more important for determining quantum yields, so these measurements were made on instruments with monochromators.
From the integrated emission intensities ATP gave an 85% enhancement
in fluorescence, whereas ADP gave an 18% quench, in line with previous
measurements on the W501+ construct (12). Early spectrophotometric
studies on skeletal muscle myosin showed that there was a significant
change in the absorption spectra induced by nucleotides (1). We
reinvestigated this aspect with D. discoideum W501+
and found that at 297 nm there was a 13% increase in the absorption
coefficient on ATP addition, but there was no detectable change with
ADP at this wavelength. Thus, although there is a significant effect of
ATP on the ground state properties of Trp-501, the fluorescence
emission process is much more sensitive to the presence of nucleotide.
This result is clearly demonstrated in a stopped-flow experiment using
the same optics for the absorption and excitation process (Fig.
1). At wavelengths less than 290 nm the
absorption change on ATP addition was inverted in sign. However, under
these conditions the signal has a contribution from tyrosine residues,
which outnumber the tryptophan residues by 29:1.
|
Quantum yields were calculated from the spectrally resolved
emission data using free tryptophan as a standard (Table
I). The addition of
AlF
|
Trp-501 Fluorescence Lifetime--
Previous studies of the
tryptophan fluorescence lifetime of myosin were carried out using a
native protein fragment that contained 5 tryptophan residues and
revealed 3 classes of tryptophans with lifetimes of 0.72, 4.5, and 8.8 ns (23). However, for most proteins it has been shown that a single
tryptophan residue displays more than a single lifetime component and
that site-directed mutagenesis is required for an unambiguous
assignment of individual residues. In the case of the D. discoideum W501+ myosin motor domain we found that the lifetime
decay was well fitted by two components with values of the order of
1 and 5 ns. Interestingly on addition of nucleotides, these lifetimes
did not change significantly; rather, the fractional amplitudes changed
(Fig. 2 and Table I). This agrees with
the earlier studies (23) which showed that the major effect of ATP was
to increase the fractional contribution of the 4.5-ns component. The
change in mean lifetime is in the direction expected for the observed
emission intensity i.e. ATP and ADP·AlF4 cause
a relative increase in the 5-ns component and an increase in the
overall emission intensity, whereas ADP causes smaller changes in the
opposite direction. Thus the nucleotide-induced changes in emission
intensity contain a dynamically quenched component. However,
quantitative comparison shows that the change in lifetime is only a
partial explanation of the change in emission intensity. From the area
under the normalized decay curves (Table I), ATP induces a 1.18 increase in mean lifetime relative to the nucleotide-free state,
whereas ADP induces a reduction by 0.92. On the other hand, the
measured emission intensity from the total counts (i.e. area under the un-normalized decay curves) was increased by 1.38 by ATP and
reduced by 0.8 in the presence of ADP. The relative total photon counts
are less than the maximum changes from the integrated fluorescence
emission spectra (Table I) because light was collected from the red end
of the spectrum (WG335 nm cut-off filter; see above). However, these
values provide further evidence that the nucleotide-induced changes in
Trp-501 fluorescence emission have both static and dynamic components.
Thus, in a significant proportion (
60%; see below) of the
population Trp-501 forms a ground state complex with a quenching center
that changes the absorption characteristics and causes the excited
state to return to the ground state by a nonradiative mechanism.
|
To estimate the contribution of the statically quenched component in the various nucleotide states, the quantum yields were compared with that of free tryptophan (0.13). If it assumed that free tryptophan in aqueous solution is quenched exclusively by a dynamic mechanism, then the intrinsic (natural) lifetime of tryptophan is about 16 ns. On this basis the fractional contributions of the static, 1-, and 5-ns components can be evaluated for each nucleotide state of the Trp-501 motor domain (Table II). This analysis reveals that the enhancement in Trp-501 fluorescence on addition of ATP is caused through a decrease in the statically quenched component and an increase in the 5-ns lifetime component with little change in the 1-ns lifetime component. The subsequent net quench observed after turnover to ADP arises from an opposite shift in the fractions of statically quenched and 5-ns components, again with little change in the 1-ns component.
|
As a control sample, we measured the fluorescence lifetime of
human serum albumin under the same conditions and found two components
with 1 = 1.3 ns (a1 = 0.32) and
2 = 6.0 ns (a2 = 0.68), in fair
agreement with values of
1 = 1.42 ns
(a1 = 0.16) and
2 = 6.06 ns
(a2 = 0.84) determined by phase fluorometry
(24). We also measured the lifetime of NATA and found a two-component fit with
1 = 0.98 ns (a1 = 0.24)
and
2 = 4.7 ns (a2 = 0.76). Other
workers have reported satisfactory fits to a single component with
lifetime of 3.2 ns (22). Lifetime components of 5-ns in tryptophan
analogs have been ascribed to photoconversion of the indole ring during
the exposure to the excitation light. This does not appear to be a
contributory factor in our measurements of the D. discoideum
W501+ myosin motor domain, at least. When the ATP was hydrolyzed after
several hours of incubation, the emission intensity fell, and the
lifetime distribution returned to that characteristic of the ADP state
(i.e. a decrease in the 5-ns component).
As a control for the contribution of tyrosine fluorescence to the
signal, a tryptophan-less construct, W, was investigated using the
same instrumental settings. When fluorescence was detected through a
335-nm cut-off filter very little signal was obtained (see Ref. 12).
With a 320-nm cut-off filter sufficient counts were obtained to reveal
lifetimes of 0.4 ns (a1 = 0.72) and 3.4 ns
(a2 = 0.28). These data indicate that there is
negligible tyrosine contribution to the lifetimes resolved in the W501+
construct at wavelengths >335 nm, despite the presence of 29 tyrosine residues.
Trp-501 Steady-state and Time-resolved
Anisotropy--
Steady-state fluorescence anisotropy measurements
showed that there was no significant change in anisotropy of tryptophan in the D. discoideum W501+ construct on addition of ATP or
ADP (Table I). The values observed of 0.21 at 20 °C were close to the limiting value of 0.23 measured in the presence of 90% glycerol at
9 °C. In contrast, when the D. discoideum W501+ motor
domain was denatured with 6.6 M guanidinium hydrochloride,
the anisotropy fell to 0.1. These data suggest that the tryptophan is
relatively immobile in all native states of W501+, and reducing the
rotational rate of the protein by increased viscosity has only a
limited effect. The difference between the limiting anisotropy value
and the theoretical maximum value of 0.4 is indicative of a difference in the angle between the absorption and emission dipoles and/or small
amplitude, high frequency motions (t 1 ns) that lead
to partial depolarization. The value of 0.23 is close to that observed for immobilized tryptophan in model studies, but this measurement shows
a very steep dependence of excitation wavelength in the region of
290-300-nm excitation (22).
These steady-state studies were complemented with time-resolved
anisotropy measurements. As expected, a high limiting anisotropy value
was observed (around 0.2) with only a slight decay over 40 ns in all
records (Fig. 3). The slightly lower
limiting anisotropy (Ao) compared with the
steady-state measurements (Ass) could be a
result of the lower discrimination of the film polarizers compared with
the Glan-Thomson prisms. The first time point, coincident with the peak
of the laser pulse, showed a slightly higher anisotropy value
(0.24-0.27), but the values after 80 ps were not significantly different from that of the major decay profile. The first two time
points may indicate a very rapid but limited amplitude motion of
Trp-501; however, any breakthrough scattered light would also contribute to this signal, and we have not attempted to analyze it
further. We focus on the major slow phase in the anisotropy decay.
|
The short lifetime of tryptophan results in a limited ability to follow
the anisotropy decay beyond 10-20 ns. However, because a protein
solution is randomly oriented, there should be no residual anisotropy
at infinite time, and the data may be analyzed by force fitting it to
an exponential function that decays to zero. With such an analysis,
single rotational correlation times, , of the order of 50-70 ns,
were obtained for all nucleotide states of the D. discoideum
motor domain (Table I). We found no systematic difference in
for
the different nucleotide states in three independent runs. This time
scale is close to that expected for the whole protein molecule.
Although we cannot rule out that there is a second slower rotational
correlation time, the calculated limit from the whole protein motion
indicates that this would not be longer than about 150 ns. Whole
molecule relaxation times from electric birefringence of about 250 ns
have been reported for a myosin subfragment containing the complete
regulatory domain (25) which, when extended, is nearly 80% longer than
the D. discoideum domain motor domain used here. As a
control sample we measured the anisotropy decay of human serum albumin
(molecular weight 66,000) under the same conditions and found the
rotational correlation time,
, of the slow component to be 39.5 ns
at 20 °C, in good agreement with 38.6 ns determined by phase
fluorometry (24). Overall, the time-resolved anisotropy data
corroborate the steady-state values and demonstrate that the Trp-501
has little independent motion relative to the rotational correlation
time of the whole protein in all nucleotide states examined.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The photophysics of tryptophan in proteins is a complex topic, but
several key features have been elucidated over the last few decades
through the use of model compounds and single tryptophan-containing proteins (22, 26). The indole ring has two emission dipoles La and Lb
that are oriented at 90° to each other. The former is almost parallel
to the absorption dipole and is responsible for high positive
anisotropy values when immobilized. At an excitation wavelength of 295 nm and emission collection >330 nm, the Lb emission is unlikely to
make any significant contribution to the observed signal (22). However,
tryptophan emission remains complex because different rotamers bring
the indole ring into proximity with different groups whose quenching
efficiency varies. In free tryptophan, the interconversion of rotamers
is slower than the fluorescence emission process, leading to a
non-single exponential profile. Under aqueous conditions at pH 7, two
phases are resolved (with around 0.5 and 3.1 ns) with the shorter
lifetime attributed to quenching by the positively charged amino group
that is in closest proximity in one rotamer (27). This short lifetime
component is lacking in the NATA derivative. Tryptophan residues within proteins are less susceptible to quenching from their own uncharged
-amide and
-carbonyl groups, but they may have additional
interactions with side chains brought into close proximity within the
tertiary structure. From the quantum yield of around 0.13 for free
tryptophan and lifetimes of 0.5 and 3.1 ns in aqueous solvents, the
intrinsic lifetime of tryptophan is calculated to be about 16 ns,
assuming that the quenching mechanism is exclusively dynamic in nature. For tryptophan residues in proteins, lifetimes typically range from 0.1 to 10 ns (28), demonstrating that even when protected from solvent,
interactions with other side chains or chromophores provide an
efficient route for relaxation from the excited state. Groups that have
been suggested to contribute to quenching within proteins are the side
chains of lysine, tyrosine, glutamine, asparagine, glutamic acid,
aspartic acid, cysteine, and histidine (26) as well as the backbone
carbonyl groups (29). In general, multiphasic or stretched exponential
kinetics are to be expected for the lifetimes of single tryptophan
residues in proteins, and this is born out in practice. Single
exponential kinetics are only observed if the local environment is
homogeneous or nearby groups are in very rapid motion (subnanosecond)
to average out any inhomogeneity. Even in protein crystals where a
tryptophan residue is clearly resolved and appears to reside in a
unique orientation, fluorescence lifetime measurements of the
crystalline state show multiple phases (27). These results are not
incompatible because the existence of multiple orientations may go
undetected in fitting the electron density. Resolutions in excess of
1.5 Å are generally required to resolve multiple orientations of side
chains in protein crystals.
With this background, it is not surprising that the fluorescence
lifetime of Trp-501 in the D. discoideum myosin motor domain deviates from single exponential kinetics. More unexpected is the
finding that changes induced by nucleotide appear to be associated with
a relative change in the amplitudes of the lifetime components with
little change in the associated time constants, which remain at about 1 and 5 ns. This finding is to some extent linked to fitting the data to
a biexponential function. When fitted to a triexponential function
there was no improvement in fit for the data obtained in the presence
of ATP and ADP·AlF
|
One possible interpretation of the data is that Trp-501 is located in a partially solvent exposed, dynamic region of the protein such that the side chain interacts with solvent water and other nearby side chains on the nanosecond time scale to effect collisional quenching. Changes in gross protein conformation associated with different nucleotide states cause a change in the weighting of the various interactions experienced by Trp-501 but not of their nature. The large and nucleotide-independent anisotropy value (~0.2) of Trp-501 and the long rotational correlation time (~60 ns) argue that the Trp-501 side chain itself is relatively static, and hence the residues with which it interacts may provide the dynamics associated with the fluorescence relaxation process (1-5 ns). The microstates may correspond to different rotamers of the Trp-501 side chain which interconvert on a time scale of >60 ns to expose the indole ring to different environments (Fig. 4). It appears unlikely that the relay loop is melted in the open state, otherwise the anisotropy values would be expected to be close to that of the denatured state (A = 0.1). However, because the anisotropy decay appears limited by the rotation of the whole domain, it is possible that Trp-501 shows different rotational dynamics in different biochemical states but on a time scale of >60 ns. The failure to resolve the relay loop in many crystal structures therefore appears to arise from static disorder. This conclusion is in accord with cross-linking studies (20) that show that the relay loop bearing Trp-501 remains in close contact with the converter domain throughout the actin-activated ATPase cycle (between residues Ile-499 and Arg-738 at least).
Another complication that must be considered is that although
x-ray crystallography of myosin may yield a particular structure for a
particular bound nucleotide, kinetic studies show that a number of
states are often significantly occupied in solution. In particular, the
myosin-ADP·BeFx and AMP·PNP complexes show multiple states
whose equilibrium distribution is particularly temperature-sensitive
over the 0-20 °C range (12). We have therefore focused on other
complexes (ADP and ADP·AlF·ATP, M*·ADP·Pi, and M
·ADP states (Reaction 1)
with occupancies of ~29, 70, and 1% at 20 °C (12). It is
therefore pertinent to question whether the M*·ADP·Pi
state itself could be associated with a single 5-ns lifetime component
and whether the observed heterogeneity (Table I) arises from the other
nucleotide states rather than microstates within a single nucleotide
state. Relevant to this argument is the possibility that the 5-ns
component is underestimated in the presence of ATP after selection
by the WG335 cut-off filter because the spectrum is blue shifted.
Correction on the basis of the observed enhancement (1.38) compared
with the integrated corrected emission spectra (1.85, Table I) suggest that the corrected ratio in the presence of ATP may be
a1 = 0.2 for the 1-ns component and
a2 = 0.8 for the 5-ns component. Given the
uncertainties in the exact distributions, it is possible that the
M*·ADP·Pi state could be associated with essentially a
single 5-ns lifetime state, and the 1ns component arises from the
M
·ATP and M
·ADP states. However, there is still a requirement
for a statically quenched microstate of M*·ADP·Pi to
account for the relatively low quantum yield observed in the presence
of ATP compared with that of free tryptophan and NATA. Therefore
M*·ADP·Pi comprises at least two microstates. In the
presence of saturating ADP, >95% of the myosin should be in the
M
·ADP state, yet at least three spectroscopic microstates are
present (statically quenched, 1-ns and 5-ns components). Likewise,
there are at least three microstates for the nucleotide-free myosin
(Table II). It should be noted that the microstates within each class
are not necessarily identical in local conformation, and different
quenching groups might be involved in different biochemical states.
Indeed, the quench in fluorescence emission intensity observed in the
M
·ADP is accompanied by a blue shift, as is the enhancement
observed in M*·ADP·Pi, implying the intermediate
intensity of the apo-state is not simply a mixture of the these two
(12).
Interestingly, the fluorescence enhancement and quantum yield observed
in the presence of ADP·AlF4 is very similar to that seen
in the presence of ATP. Given that the steady-state ATPase complex
contains a small but significant fraction of quenched nucleotide states
(M·ATP and M
·ADP), it might be expected that M*·ADP·AlF4 would show up to a 30% higher fluorescence
emission than in the presence of ATP. These data suggest that
M*·ADP·AlF4 could be in equilibrium with a small amount
of M
·ADP·AlF4 state at 20 °C. Temperature and
pressure jump experiments are in progress to investigate the kinetics
of reequilibration between different bound-nucleotide states that are
not resolved by rapid mixing methods.
The ATP-induced enhancement of the relay loop tryptophan (Trp-501 or equivalent residue) appears to be a conserved property of myosin II molecules. It is difficult to compare the magnitude of enhancement in different species in the literature because the observed values are instrument-dependent, but the direction of the change is common. On the other hand, although ADP generally induces a smaller change in fluorescence emission, its effect appears to be species-specific. A quench is observed with D. discoideum myosin (12), but an enhancement is observed with a smooth muscle myosin construct (10). This difference could reflect either a different gross conformation (i.e. ADP is able to partially induce the closed conformation in smooth myosin) or a difference in the local amino acid structure around the relay loop tryptophan. Comparisons of the x-ray structures (5, 6) reveal several side chains and backbone carbonyl groups that could contribute to quenching, and these are generally highly conserved residues. Candidates within the vicinity (<1 nm) of the Trp-501 side chain are Glu-490, Gln-491, Glu-493, Tyr-494, Ile-199 (backbone carbonyl), Asn-500, Trp-501 (backbone carbonyl), Thr-502 (backbone carbonyl), Phe-501 (backbone carbonyl), Lys-690 (including backbone carbonyl), and Gly-691 (backbone carbonyl). Residues in bold are within 0.5 nm of Trp-501 in the closed-state (M·ADP·Vi) and are the prime candidates for involvement in quenching. Unfortunately, there are no structures of the D. discoideum motor domain with sufficient resolution in the relay loop to define the position of the corresponding residues in the open state (19). Nevertheless, this region can be compared with motor domains from other species. However, from the limited comparison possible to date, there is no residue that can be uniquely identified with the dequenching process observed in the presence of ATP.
The finding that the Trp-501 is effectively immobile on the time scale of fluorescence emission has bearing on fluorescence energy transfer measurements. For example, Yengo et al. (10) assumed free rotation or random orientation of the equivalent Trp-512 residue in smooth myosin in calculating the distance from mant-nucleotide bound at the active site. Previous anisotropy studies on mant-nucleotide complexes showed that the acceptor mant group is relatively static when bound to myosin (31, 32). Thus, such distance calculations using the Forster equation will be extremely dependent on the relative orientation of the tryptophan donor and mant-acceptor dipoles. Angles in the range of about 35o from the optimal dipole orientation could account for the underestimate of the separation by fluorescence resonance energy transfer of 26 Å (10) compared with 45-50 Å measured from the crystal structure of the M·mant·ADP·BeFx complex. It should be noted that Trp-501 was not resolved in this structure, but the adjacent residue Asn-500 was assigned and therefore limits the position of Trp-501 (32). It is likely that the apparent 8 Å movement of the Trp-512 residue in different nucleotide states (10) is a reflection of different orientations of the tryptophan (as might occur with different rotamer distributions) rather than translational movement. Alternatively, the fluorescence resonance energy transfer measurements (10) may have a differential contribution from the numerous tyrosine residues, particularly as 285-nm excitation was used. In principle, changes in the predominant tryptophan side chain orientation should be revealed in the time-resolved fluorescence anisotropy studies because the motor domain is not spherical, and therefore different probe orientations should have different limiting rotational correlation times. However, the dimensions of the motor domain of ~100 × 60 × 40 Å would not result in a strong dependence, particularly if the tryptophan reoriented around the shorter axes. Notably, we found that the rotational correlation time of a single tryptophan D. discoideum motor domain mutant, F129W, measured under the same conditions was around 100-120 ns compared with 60 ns for Trp-501. This suggests that the emission dipole of the former is aligned predominantly along the long axis of the motor domain and/or that Trp-501 has a small degree of independent motion.
In conclusion, the time-resolved fluorescence data presented here show
that Trp-501 experiences at least three different microenvironments. The coupling between the occupancy of the nucleotide site and the
Trp-501 environment does not correspond to a simple one-to-one relationship. Rather, nucleotide binding allows the Trp-501 side chain
to redistribute among different environments on a time scale of >5 ns
and <20 µs. It is therefore not possible to equate a particular
nucleotide state or a particular gross conformation (open or closed
state), to a unique set of coordinates for Trp-501 and its surrounding
residues, as might be implied by some crystal structures. This behavior
has been reported in other probe studies that reflect gross
conformational rearrangements of the myosin motor domain (33, 34).
Nevertheless, the crystal structures are valuable in defining the
potential environments that surround the Trp-501 side chain. We find no
evidence that the relay loop is dynamically disordered on the
nanosecond time scale in the open conformation, as might be surmised
from the crystal data. In this respect our data are concordant with
that of Shih and Spudich (20) showing that the relay loop does not make
any large scale rearrangements relative to the converter region during
ATPase activity.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank William Shih and Jim Spudich for an advanced copy of their manuscript and discussions. We thank Tony Parker for access to the Central Laser Facility.
![]() |
Note Added in Proof |
---|
A crystal structure (Protein Data Bank accession code 1G8X) of an engineered D. discoideum myosin motor domain locked in the open state has recently been published in which the two molecules in the unit cell have the same gross conformation, but different rotamers of Trp-501 are resolved (Kliche, W., Fujita-Becker, S., Kollmar, M., Manstein, D. J., and Kull, F. J. (2001) EMBO J. 20, 40-46). This structure also identifies the side chain of Arg-747 as a potential quenching group in the open state.
![]() |
FOOTNOTES |
---|
* This work was supported in part by the Biotechnology and Biological Sciences Research Council and the Wellcome Trust.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ Visit funded by a Hungarian State Eötvös fellowship.
** To whom correspondence should be addressed. E-mail: crb5@le.ac.uk.
Published, JBC Papers in Press, February 22, 2001, DOI 10.1074/jbc.M010886200
2 A. Málnási-Csizmadia and C. R. Bagshaw, unpublished observations.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
ATPS, adenosine
5'-3-O-(thio)triphosphate;
NATA, N-acetyl
L-tryptophanamide;
TES, N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid;
AMP·PNP, adenosine 5'-(
,
-imino)triphosphate;
mant-, 2'(3')-O-(N-methylanthraniloyl);
BeFx, beryllium
fluoride with undetermined F content.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Morita, F., and Yagi, K. (1966) Biochem. Biophys. Res. Commun. 22, 297-302[Medline] [Order article via Infotrieve] |
2. | Bagshaw, C. R., Eccleston, J. F., Trentham, D. R., Yates, D. W., and Goody, R. S. (1972) Cold Spring Harbor Symp. Quant. Biol. 37, 127-135 |
3. | Werber, M. M., Szent-Györgyi, A. G., and Fasman, G. D. (1972) Biochemistry 11, 2872-2883[Medline] [Order article via Infotrieve] |
4. | Rayment, I., Rypniewski, W. R., Schmidt-Bäse, K., Smith, R., Tomchick, D. R., Benning, M. M., Winkelmann, D. A., Wesenberg, G., and Holden, H. M. (1993) Science 261, 50-58[Medline] [Order article via Infotrieve] |
5. | Fisher, A. J., Smith, C. A., Thoden, J., Smith, R., Sutoh, K., Holden, H. M., and Rayment, I. (1995) Biochemistry 34, 8960-8972[Medline] [Order article via Infotrieve] |
6. | Smith, C. A., and Rayment, I. (1996) Biochemistry 35, 5404-5417[CrossRef][Medline] [Order article via Infotrieve] |
7. | Dominguez, R., Freyzon, Y., Trybus, K. M., and Cohen, C. (1998) Cell 94, 559-571[Medline] [Order article via Infotrieve] |
8. | Houdusse, A., Kalabokis, V. N., Himmel, D., Szent-Györgyi, A. G., and Cohen, C. (1999) Cell 97, 459-470[Medline] [Order article via Infotrieve] |
9. | Batra, R., and Manstein, D. J. (1999) Biol. Chem. Hoppe-Seyler 380, 1017-1023 |
10. |
Yengo, C. M.,
Chrin, L. R.,
Rovner, A. S.,
and Berger, C. L.
(2000)
J. Biol. Chem.
275,
25481-25487 |
11. |
Onishi, H,
Konishi, K.,
Fujiwara, K.,
Hayakawa, K.,
Tanokura, M.,
Martinez, H. M.,
and Morales, M. F.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
11203-11208 |
12. | Málnási-Csizmadia, A., Woolley, R. J., and Bagshaw, C. R. (2000) Biochemistry 39, 16135-16146[CrossRef][Medline] [Order article via Infotrieve] |
13. |
Hiratsuka, T.
(1992)
J. Biol. Chem.
267,
14949-14954 |
14. | Park, S., Ajtai, K., and Burghardt, T. P. (1997) Biochemistry 36, 3368-3372[CrossRef][Medline] [Order article via Infotrieve] |
15. | Park, S., and Burghardt, T. P. (2000) Biochemistry 39, 11732-11741[CrossRef][Medline] [Order article via Infotrieve] |
16. | Geeves, M. A., and Holmes, K. C. (1999) Annu. Rev. Biochem. 68, 687-728[CrossRef][Medline] [Order article via Infotrieve] |
17. | Bagshaw, C. R., Eccleston, J. F., Eckstein, F., Goody, R. S., Gutfreund, H., and Trentham, D. R. (1974) Biochem. J. 141, 351-364[Medline] [Order article via Infotrieve] |
18. | Johnson, K. A., and Taylor, E. W. (1978) Biochemistry 17, 3432-3442[Medline] [Order article via Infotrieve] |
19. |
Gulick, A. M.,
Bauer, C. B.,
Thoden, J. B.,
Pate, E.,
Yount, R. G.,
and Rayment, I.
(2000)
J. Biol. Chem.
275,
398-408 |
20. |
Shih, W. M.,
and Spudich, J. A.
(2001)
J. Biol. Chem.
276,
19491-19494 |
21. | Manstein, D. J., Schuster, H.-P., Morandini, P., and Hunt, D. M. (1995) Gene (Amst.) 162, 129-134[CrossRef][Medline] [Order article via Infotrieve] |
22. | Lakowicz, J. R. (1999) Principles of Fluorescence Spectroscopy , 2nd Ed. , Kluwer Academic Publishers, Norvell, MA |
23. | Torgerson, P. M. (1984) Biochemistry 23, 3002-3007[Medline] [Order article via Infotrieve] |
24. | Lakowicz, J. R., and Gryczynski, I. (1992) Biophys. Chem. 45, 1-6[CrossRef][Medline] [Order article via Infotrieve] |
25. | Highsmith, S., Polosukhina, K., and Eden, D. (2000) Biochemistry 39, 12330-12335[CrossRef][Medline] [Order article via Infotrieve] |
26. | Chen, Y., and Barkley, M. D. (1998) Biochemistry 37, 9976-9982[CrossRef][Medline] [Order article via Infotrieve] |
27. | Dahms, T. E. S., and Szabo, A. G. (1997) Methods Enzymol. 278, 202-221[Medline] [Order article via Infotrieve] |
28. | Beecham, J. M., and Brand, L. (1985) Annu. Rev. Biochem. 54, 43-71[CrossRef][Medline] [Order article via Infotrieve] |
29. | Sillen, A., Fernando-Díaz, J., and Engelborghs, Y. (2000) Protein Sci. 9, 158-169[Abstract] |
30. | Gastmans, M., Volckaert, G., and Engelborghs, Y. (1999) Proteins Struct. Funct. Genet. 35, 464-474[CrossRef][Medline] [Order article via Infotrieve] |
31. |
Rosenfeld, S. S.,
Xing, J.,
Rener, B.,
Lebowitz, J.,
Kar, S.,
and Cheung, H. C.
(1994)
J. Biol. Chem.
269,
30187-30194 |
32. | Bauer, C. B., Kuhlman, P. A., Bagshaw, C. R., and Rayment, I. (1997) J. Mol. Biol. 274, 394-407[CrossRef][Medline] [Order article via Infotrieve] |
33. |
Baker, J. E.,
Brust-Mascher, I.,
Ramachandran, S.,
LaConte, L. E. W.,
and Thomas, D. D.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
2944-2949 |
34. | Shih, W. M., Gryczynski, Z., Lakowicz, J. R., and Spudich, J. A. (2000) Cell 102, 683-694[Medline] [Order article via Infotrieve] |