From the Department of Neurology, University of
Alabama at Birmingham, Birmingham, Alabama 35294-3293, the
Neurology Service, Department of Veterans Affairs Medical
Center, Birmingham, Alabama 35294, and the Departments of
¶ Physics,
Applied Physics, and
** Biological Sciences, Stanford University, Stanford,
California 94305
Received for publication, January 27, 2003
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ABSTRACT |
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The ability of kinesin to travel long distances
on its microtubule track without dissociating has led to a variety of
models to explain how this remarkable degree of processivity is
maintained. All of these require that the two motor domains remain
enzymatically "out of phase," a behavior that would ensure that, at
any given time, one motor is strongly attached to the microtubule. The
maintenance of this coordination over many mechanochemical cycles has
never been explained, because key steps in the cycle could not be
directly observed. We have addressed this issue by applying several
novel spectroscopic approaches to monitor motor dissociation, phosphate release, and nucleotide binding during processive movement by a dimeric
kinesin construct. Our data argue that the major effect of the internal
strain generated when both motor domains of kinesin bind the
microtubule is to block ATP from binding to the leading motor. This
effect guarantees the two motor domains remain out of phase for many
mechanochemical cycles and provides an efficient and adaptable
mechanism for the maintenance of processive movement.
Members of the kinesin family of molecular motors are capable of
taking over 100 steps on their microtubule track without dissociating,
a feature that would be necessary for a transport motor that operates
in isolation (1-6). A variety of kinetic, structural, and
mechanical studies have revealed that this processive behavior requires
that two motors of kinesin remain in different structural and enzymatic
states during a processive run (7-10, 11, 12). This would
ensure that, at any given time, at least one of the two heads would
remain strongly attached to its track, preventing the motor from
prematurely detaching. Such coordination requires a way for the two
motor domains to communicate their structural states to each other
while walking processively. Several lines of evidence suggest that this
allosteric communication is mediated through the internal load
generated when both heads attach to the microtubule (1, 13-16). As
illustrated below in Fig. 1, kinesin initiates its mechanochemical
cycle with its attached head (green) nucleotide free and its
tethered head (magenta) containing ADP in the active site.
ATP binding to the attached head reorients its neck linker
(blue), which swings the tethered head forward to the next
tubulin-docking site. ADP is then released from the new, weakly bound
leading head (magenta) to produce an intermediate in which
both heads are strongly bound to the microtubule. This situation would
generate rearward strain on the neck linker of the leading
head, depicted as a left pointing arrow, and
forward strain on the corresponding structure of the
trailing head, depicted as a right pointing arrow.
It has been proposed that this strain generates processivity by
accelerating release of the trailing head (13, 17). In this mechanism,
release of the ADP-containing trailing head would be very slow in the
absence of forward strain and fast in its presence. In such a system,
the greater that forward strain accelerates kdMT, the greater the degree of
processivity. However, there is an internal inconsistency with this
scheme. If kinesin's processivity were dependent solely on
this mechanism, the motor would dissociate from the microtubule after
only a few steps. The reasoning behind this is illustrated in Fig. 1.
We have recently shown (18) that the effective rate of trailing head
dissociation (kdMT ~ 50 s An alternative possibility is that rearward strain on the leading head
slows ATP binding and subsequent hydrolysis, insuring that the leading
head would remain strongly attached until the trailing head had
dissociated. ATP would then rapidly bind to the leading head and cause
the trailing head to swing forward to the next tubulin-docking site.
Processive movement would be favored, because the rate of this forward
stepping movement, at ~800 s
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL METHODS
RESULTS
DISCUSSION
REFERENCES
1) is appreciably slower than that for ADP release
(kdADP = 170 s
1, this
work, and Refs. 8, 19-21), and ATP hydrolysis
(kh, 100 s
1, Refs. 8, 19, 21, 22).
This would lead to accumulation of a kinesin intermediate with both
heads attached to the microtubule, with the leading head nucleotide
free, and with ADP-Pi in the active site of the trailing
head. Given millimolar intracellular ATP concentrations and an apparent
second order rate constant of > 1 µM
1s
1 (2, 8, 15, 19, 20, 23),
ATP would then rapidly bind to the new leading head (>1000
s
1) and be hydrolyzed. This would generate an
intermediate with both heads weakly bound, and dissociation would
rapidly follow, as indicated in Fig. 1 by the red X. The
fact that kinesin is highly processive (1-6) argues that there is a
mechanism that prevents it from proceeding down this pathway.
1, is nearly sixteen
times faster than the rate of trailing head dissociation (18).
Furthermore, blocking ATP binding to the forward head while it was
experiencing rearward strain would prevent the motor from proceeding
down the pathway marked by the red X in Fig.
1.
View larger version (24K):
[in a new window]
Fig. 1.
Kinesin's first two steps. The two
motor domains of kinesin are distinguished by magenta
and green shading and are depicted walking on a track of
tubulin dimers. The neck linkers are depicted by the blue
lines that connect the motor domains to the black
coiled coil dimerization segment. The mechanochemical cycle is
initiated by ATP binding to the green, attached motor domain
and is characterized by the equilibrium constant
KATP. This leads to rapid (~800
s 1) docking of this motor domain's neck linker (depicted
as a straightening of the blue neck linker),
which throws the tethered magenta motor forward toward the
next tubulin docking site. Release of ADP from the new leading
(magenta) motor, occurring with forward rate constant
kdADP (170 ± 17 s
1),
is followed by ATP hydrolysis (kh, ~100
s
1), which leads to binding of both heads to the
microtubule. This places the two neck linkers under mechanical strain
(depicted as the rightward and leftward
pointing blue arrows). In the absence of any mechanism to
prevent it, ATP could then bind to the empty, leading motor domain
(magenta) and then become rapidly hydrolyzed. This would
produce motor dissociation from the microtubule after only two
turnovers. That this does not happen (symbolized by the red
X) implies that a mechanism must exist to prevent ATP from binding
to the leading head while it is experiencing rearward strain (see text
for details). Instead, hydrolysis is followed by dissociation of the
rear head, characterized by rate constant
kdMT, which occurs concomitantly with
phosphate release.
Determining whether processivity depends on the first mechanism, the
second, or to some degree on both requires the ability to unambiguously
measure the rates of key steps in the mechanochemical cycle and the
effects of strain on these rates. These include the rates of trailing
head dissociation, of ADP release from the tethered head after it
attaches to the microtubule, and of ATP binding to the new, leading
head. In this study, we apply the spectroscopic approaches developed in
our prior work to make these measurements (15, 18, 23). This has
allowed us to generate a model that provides an efficient yet adaptable
mechanism for insuring processivity in this motor.
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EXPERIMENTAL METHODS |
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Generation of K413FBIO-- K413W340F, a cysteine-light recombinant kinesin construct with tryptophan 340 replaced by phenylalanine, consists of the first 413 amino-terminal residues of human kinesin, and was generated as described in our previous study (18). K413BIO is a construct in which K413W340F is fused at the carboxyl terminus to a biotinyl transferase recognition peptide, followed in turn by a hexahistidine tag for affinity purification. The peptide sequence, with its attached biotin, was incorporated into the K413 sequence to allow for attachment of the motor to streptavidin-coated beads used in motility assays in vitro. The plasmid containing the K413W340F mutant kinesin construct served as a PCR template to amplify the kinesin insert with the following primers: upstream, 5'-AGATATACATATGGCGGACCTGGCC-3'; downstream, 5'-AAGTTGCATGTGCTCGAGAAAATTTCCTATAACTCCAAT-3'. The underlined sequences are the NdeI and XhoI restriction sites, respectively. The fragment was cloned into pCR2.1-TOPO (Invitrogen, Carlsbad, CA) and excised with NdeI and XhoI. The fragment was purified and then ligated into a pET-21 vector with an in-frame biotinyl transferase recognition peptide sequence at the XhoI (carboxyl terminus) site. The resultant construct was verified by sequence analysis.
In Vitro Motility Studies of K413BIO-- Single molecule kinesin bead motility assays were performed essentially as described previously (24). 500-nm diameter carboxy-modified latex beads (Bangs Laboratories, Fishers, IN) were covalently biotinated with biotin-x-cadaverine (Molecular Probes, Eugene, OR), coated with avidin-DN (Vector Laboratories, Burlingame, CA), and purified by repeated pelleting and resuspension followed by sonication to eliminate clumping. Diluted kinesin constructs were mixed with the beads and incubated at 4 °C for 4 h in assay buffer containing 80 mM PIPES,1 pH 6.9, 50 mM potassium acetate, 4 mM MgCl2, 2 mM dithiothreitol, 1 mM EGTA, 7 µM Taxol, various ATP concentrations, and 2 mg/ml bovine serum albumin as a blocking protein. The beads were diluted to 80 fM, and final kinesin dilutions were chosen such that, on average, fewer than half of the beads moved (typically 1:500,000 to 1:1,000,000 from ~100 µM stock). An oxygen-scavenging system (37) was added to the kinesin:bead mixture just prior to measurement.
Flow cells with a volume of ~20 µl were constructed by using two strips of doubly adhesive tape to form a channel between a microscope slide and a No. 11/2 coverslip (cleaned by sonication in 5 M ethanolic KOH and coated with polylysine). Ingredients were introduced in the following order: taxol-stabilized MTs (polymerized from purified bovine brain tubulin, Cytoskeleton, Inc., Denver, CO) followed by a 10-min incubation, assay buffer wash followed by a 10-min incubation, and then kinesin:bead mixtures. To ensure that measurements reflected single molecule properties, data was only collected from assays in which fewer than half of the tested beads moved. All chemicals used were from Sigma, except bovine serum albumin and the ingredients used in the oxygen scavenging system (Calbiochem, San Diego, CA). Kinesin velocities and run lengths were measured by centroid video tracking (25) with sub-pixel resolution using a commercial video tracking software package (Isee Imaging Systems, Raleigh, NC).
Fluorescence Methodologies--
Labeling of K413W340F and
K413W340FBIO with
5-(((2-iodoacetyl)amino)ethyl)aminonaphthalene-1 sulfonic
acid or tetramethyl rhodamine maleimide was carried out as
described previously (15, 18, 23). Labeling of phosphate-binding
protein by MDCC was carried out as described previously (26). Transient
kinetic measurements were made in an Applied Photophysics SX.18 MV
stopped-flow spectrometer with instrument dead time of 1.2 ms as
described previously (15, 18, 23). Unless otherwise described,
complexes of kinesin and microtubules were formed with a 5- to 10-fold
molar excess of microtubules over active sites. ADP was added to 3 µM to ensure that the tethered kinesin head of a
kinesin-microtubule complex contained ADP in the active site.
Experiments with phosphate-binding protein were carried out in
phosphate mop (0.2 unit/ml purine nucleoside phosphorylase plus 1 mM 7 methylguanosine) with a 10-fold molar excess of
MDCC-labeled phosphate-binding protein over kinesin active sites in
both syringes.
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RESULTS |
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In Vitro Motility of K413BIO
The experiments in this study utilize a cysteine-light dimeric
kinesin construct (K413), and it was necessary to establish that this
kinesin construct is processive. In our previous study, we provided
enzymatic evidence that K413 is capable of undergoing multiple
enzymatic cycles per diffusional encounter with the microtubule, a
prerequisite for processivity (18). Furthermore, attaching the biotin
transferase recognition sequence to the carboxyl terminus of K413 had
no appreciable effect on kcat (20 ± 2 s1), K0.5,MT (0.30 ± 0.1 µM), or kbi,ADP (1.28 ± 0.15 µM
1s
1) (data not shown).
However, enzymatic measures of processivity are indirect. To directly
assess processive behavior, we examined the in vitro
motility properties of K413BIO.
As with wild-type kinesin, average in vitro velocities
showed a Michaelis-Menten dependence on ATP concentration, defining values of Vmax and
KmATP of 703 ± 73 nm/s and 23 ± 9 µM, respectively (Fig.
2A). These compare with values
of 650-800 nm/s and ~80 µM for wild-type squid kinesin
measured in our laboratory (6, 27). Fig. 2B illustrates the
distribution of run lengths for K413BIO. Run length did not vary beyond
experimental error over the range of ATP concentrations examined, and
fitting data from all ATP concentrations to an exponential decay
revealed a mean run length of 276 ± 22 nm. Thus, although
in vitro velocities of K413BIO are similar to wild-type,
mean run length is reduced by a factor of 2-3. Having established that
K413BIO was processive, we decided to use it to test the two models of
how kinesin uses strain to walk processively: that forward
strain accelerates trailing head release or that rearward
strain inhibits ATP binding to the leading head.
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Evaluation of the Effect of Forward Strain on the Trailing Head
Dissociation of K413BIO:Microtubule by ATP--
If forward strain
on the trailing head accelerates its release, then we would predict
that release of the trailing head of a processively moving kinesin
dimer should be faster than that for a non-processive monomeric
construct, because the latter is incapable of generating internal
strain. The steps leading to ATP-induced dissociation from the
microtubule are summarized in Fig. 1. In this scheme, the observed
dissociation rate, ATP, depends on the binding constant
for ATP (KATP) and on the effective rate
constant for ATP-induced dissociation, ke. The
value of ke depends in turn on the values of the
forward rate constants for the two irreversible steps that lead to
dissociation, ATP hydrolysis (kh), and
subsequent dissociation from the microtubule
(kdMT). Under these conditions,
ATP is defined by the following,
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(Eq. 1) |
If strain accelerates trailing head dissociation,
ke should be greater for K413BIO than for K349.
We measured the rate of trailing head dissociation for K413BIO and
compared these results to those for monomeric K349. For K413BIO, this
was accomplished using two spectroscopic probes, AEDANS and TMR, whose
use we have previously described (18), whereas for K349 we utilized
AEDANS and turbidity. The solid line in Fig.
3 depicts the fitting of the AEDANS data
(open boxes) for K413BIO to Equation 1, whereas the
dotted line depicts the corresponding fitting of the AEDANS data (open diamonds) for K349. It is immediately apparent
that ATP is consistently faster for monomeric
kinesin than for trailing head dissociation by dimeric kinesin, the
opposite of what would be expected if strain accelerated trailing head
dissociation.
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The AEDANS probe monitors motor-microtubule association, and results with this probe for K349 are nearly identical to those using turbidity (data not shown). For K413BIO, the TMR probe monitors neck linker-neck linker reassociation, a step whose rate we have shown is kinetically controlled by dissociation of the trailing head (18). We would therefore predict that results using the TMR probe (closed triangles) should be superimposable on those using the AEDANS probe (open boxes), and Fig. 3 confirms this. Table I summarizes the values of KATP and ke for both probes. Nearly identical results were seen at 10 mM KCl (data not shown).
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Phosphate Release by K413BIO-Microtubule-- Phosphate release occurs concomitantly with trailing head dissociation (1, 2, 20). Therefore, measuring its release kinetics should provide an independent measure of ke. We accomplished this by mixing a complex of K413BIO-microtubules with a range of ATP concentrations in the presence of a fluorescently labeled phosphate-binding protein (28) and compared our results to K349. The resulting fluorescence transient consisted of an initial exponential, burst phase, followed by a linear increase, which could be described by the following relationship,
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(Eq. 2) |
Dissociation of K413BIO-Microtubule by ADP--
In the absence of
added nucleotide, kinesin attaches to the microtubule via only one
motor domain (7, 8, 29, 30). This is illustrated in Fig. 1 by the
upper left kinesin-microtubule complex. If ADP is added, it
will bind to the empty catalytic site of the attached
(green) head and dissociate the complex (17). This reaction
would occur in the absence of internal strain, because ADP does not
induce forward stepping and strong attachment of the other, tethered
(magenta) head. We would, therefore, predict that the
kinetics of K413BIO and K349 dissociation from the microtubule should
be identical. This is confirmed by comparing the solid (K413BIO) and dashed (K349) curves in the
inset of Fig. 4. ADP-induced dissociation was monitored by turbidity and by the AEDANS probe. Data
for turbidity (closed symbols) and AEDANS (open
symbols) are nearly identical and showed a hyperbolic dependence
on [ADP] for both K413BIO and K349, defining maximum rates and
apparent dissociation constants of 201 ± 27 s1 and
154 ± 58 µM and 219 ± 16 s
1 and
123 ± 19 µM, respectively.
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ADP dissociates wild-type kinesin from the microtubule at a rate of
~12 s1 (17, 31). This is considerably slower than
kcat under processive conditions, under
conditions where strain would be present, and this finding has been
used to support the argument that trailing head dissociation is
accelerated by forward strain (17). If processivity depended solely on
this mechanism, it would follow that any mutation in kinesin that
accelerates the rate of ADP-induced dissociation in the absence of
strain should reduce average run length proportionally. Nevertheless,
our data show that, although the rate of ADP-induced dissociation for
K413BIO is nearly 19-fold larger than for wild-type, mean run length is
only reduced 2- to 3-fold (Fig. 2).
Evaluation of the Effect of Rearward Strain on the Leading Head
We next set out to examine the effect of rearward strain on the
leading head by measuring the kinetics of 2'-deoxy-mant-ATP (2'dmT)
binding to a K413BIO-microtubule complex. Binding of 2'dmT was
monitored by FRET from kinesin tyrosine residues to the mant fluorophor, as previously described (18), and the experimental design
is illustrated in Fig. 5A.
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In the absence of microtubules, binding of 2'dmT to nucleotide-free
K413BIO produced a fluorescence increase characterized by a single
phase (Fig. 5B, microtubules). The rate
depended hyperbolically on [2'dmT], defining a maximum of 1033 ± 153 s
1 (Fig. 5B, inset,
dotted curve). By contrast, mixing a 1:10
K413BIO-microtubule complex with 2'dmT produced a fluorescence increase
that occurred in two distinct phases of similar amplitudes, separated
by a lag (Fig. 5B, +microtubules). The rate of
the first phase showed a hyperbolic dependence on 2'dmT concentration,
defining a maximum rate of 457 ± 56 s
1, an apparent
affinity of 80 ± 49 µM, and an apparent
dissociation rate constant of 107 ± 50 s
1 (Fig.
5B, inset, solid curve). The amplitude
of this phase is approximately half of that for an equal concentration
of K413 in the absence of microtubules. Repeating these experiments in the presence of microtubules alone produced no fluorescence change (data not shown). These findings led us to conclude that the first phase in this transient is due to 2'dmT binding to the attached, nucleotide-free head. The rate of the second rising phase also showed a
hyperbolic dependence on [2'dmT], defining a maximum rate of 39 ± 4 s
1 and an apparent affinity of 39 ± 10 µM (Fig. 6, inset, dotted curve).
Given that the amplitudes and the apparent affinities of the two phases
of the fluorescence transient are similar, we propose that the second
phase in the transient is due to binding of 2'dmT to the leading head
of a doubly attached kinesin-microtubule complex.
Why is the rate of ATP binding to the leading head so much slower than
that for the trailing head? One possibility is that it is rate-limited
by the dissociation of bound ADP. To determine if this is the case, we
measured the rate of 2'dmD dissociation from the tethered head by
mixing a complex of K413BIO-2'dmD plus a 10-fold molar excess of
microtubules with varying concentrations of ATP in the stopped flow.
The resulting fluorescence transient consisted of a single falling
phase whose rate depended hyperbolically on ATP concentration, defining
a maximum rate constant of 170 ± 17 s1 (Fig.
6, inset, solid
curve). This is over four times faster than the rate of binding of
2'dmT to this head (Fig. 6, inset, dotted curve).
Thus, nucleotide binding to the leading head of a doubly attached
kinesin-microtubule complex is rate-limited by some process other than
ADP release, and we propose that this process consists of a rearward
strain imposed on this head.
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We can test our hypothesis that ATP binding to the leading head is
strain-inhibited by examining the effect of AMPPNP on ADP-induced kinesin dissociation. Adding AMPPNP to a kinesin-microtubule complex induces the two neck linkers to separate from each other, in a manner
similar to what is seen when kinesin takes a forward step with ATP
binding (18). This occurs hand-in-hand with an acceleration of ADP
release from the tethered head (8, 19) and leads to strong binding of
both heads to the microtubule (29, 30) and to immobilization of both
neck linkers (18). Furthermore, at equilibrium, the stoichiometry of
nucleotide binding is 1 mol of AMPPNP:2.4 mol of active sites (32).
Taken together, these results indicate that AMPPNP binding to the
tethered head causes both heads to bind strongly, with the trailing
head containing AMPPNP, with the leading head nucleotide-free, and with
both heads under strain. This is illustrated in the left
half of Fig. 7A. Furthermore, adding ADP to this system will cause one of the two heads
to dissociate, leaving one head strongly bound, as illustrated in the
right half of Fig. 7A (29, 30). If rearward
strain inhibits nucleotide binding to the leading head, we would
predict that mixing a kinesin-microtubule complex plus 1 mM
AMPPNP in the stopped flow with ADP will dissociate from the leading
head very slowly when compared with ADP-induced dissociation in the absence of AMPPNP (Fig. 4).
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We measured the kinetics of leading head dissociation by mixing a
complex of 1:10 AEDANS-labeled K413BIO:microtuble plus 1 mM
AMPPNP with a range of ADP concentrations, as illustrated in Fig.
7A. An example of the fluorescence transient produced by mixing with 400 mM ADP is depicted as the red jagged
curve in the figure. Its rate demonstrated a hyperbolic dependence
on ADP concentration, defining a maximum of 0.28 s1,
nearly three orders of magnitude slower than seen in the absence of
AMPPNP (Fig. 4, inset). The apparent second order rate
constant for this process, at 0.016 µM
1
s
1, compares to a value of 1.31 µM
1 s
1 in the absence of
AMPPNP (Fig. 3, inset). Similar results were also seen using
the rhodamine probe (data not shown).
To be sure that the fluorescence changes detected with the AEDANS probe
are indeed due to the effects of ADP binding, we directly measured the
kinetics of 2'dmD binding to a 1:10 kinesin-microtubule complex in the
presence of 1 mM AMPPNP as described above (Figs. 5 and 6).
As shown in Fig. 7B, the rate of the fluorescence rise produced by mixing with 400 µM 2'dmD (final
concentration), at 0.29 s1, was nearly identical to the
rate of the fluorescence decrease seen with the AEDANS probe.
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DISCUSSION |
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The most significant finding of this study is that strain appears
to affect one discrete step in the kinesin mechanochemical cycle:
binding of ATP to the leading head. This conclusion is supported not
only by direct evidence from 2'dmT binding kinetics (Figs. 5 and 6) but
also from the effect of AMPPNP on nucleotide binding and
nucleotide-induced dissociation (Fig. 7). If rearward strain
effectively blocks ATP binding to the leading head, we can predict how
fast ATP binding can occur by using the values of the rate constants we
measured in this study. According to our model, ATP binding can only
occur after ADP dissociation from the leading head
(kdADP) and trailing head dissociation (ke). Because these two steps are irreversible
under the conditions of our experiments, at infinite ATP concentration,
the rate of binding will be equal to
1, which is in remarkable agreement with
the measured value of 39 ± 4 s
1 (Fig. 6). Our
conclusion is directly supported by recent single molecule mechanical
studies, which show that external load imposed against the direction of
motility reduces ADP binding (38). Hence, we conclude that
internal load insures that the two heads of a processive kinesin remain
out of phase for many mechanochemical cycles by hindering nucleotide
binding to the leading head.
The inset of Fig. 7 demonstrates that the second order rate constant for ADP binding is reduced at least two orders of magnitude in the presence of rearward strain. This implies that strain makes the catalytic site relatively inaccessible to nucleotide. Furthermore, we have shown that kinesin:nucleotide is an equilibrium mixture of two states (15). Taken together, these results suggest that the effect of rearward strain is to drive an equilibrium distribution of catalytic site conformations to favor one that is relatively "closed" and inaccessible to nucleotide binding.
Our results also provide a critical test of a recently proposed "inchworm" model of kinesin movement (33). In this model, the leading motor is always leading, the trailing motor is always trailing, and one motor remains enzymatically inactive throughout a processive run. Our data show that binding of 2'dmT occurs in two distinct phases (Fig. 5B, red transient), and the rates of both of these phases are considerably faster than kcat. This means that a processive kinesin moving on a microtubule reaches the steady state after two nucleotide-binding events. This is both consistent with and required by a hand-over-hand mechanism, such as the one depicted in Fig. 1. However, it is inconsistent with an inchworm mechanism, which would predict only one nucleotide-binding event before the steady state is reached.
Our model explains how processive movement by kinesin can be both
efficient and adaptable. By preventing ATP binding to the lead head,
internal strain guarantees that this head will remain strongly attached
to the microtubule at the moment that the trailing head dissociates.
ATP would then bind rapidly to the leading head (>1000
s1), but hydrolysis and subsequent dissociation would
still be relatively slow (ke = 48-55
s
1, Table I). This disparity would give the trailing head
time to swing forward and associate with the microtubule, because we have shown (18) that this process is very rapid (~800
s
1). Processivity would therefore result from two
features of the mechanochemical cycle: blocking of ATP binding to the
leading head by strain, and very rapid forward stepping of the trailing head and its concomitant docking to the microtubule surface (Fig. 1). A
particular advantage of this arrangement is that, if the tethered head
were to come across an obstacle during its forward swing, the entire
kinesin molecule would dissociate at a rate defined by
ke. This feature would enable kinesin to
sidestep an obstruction, diffuse to another microtubule, and continue
on with its journey.
Does forward strain have any effect on the trailing head? A variety of
mechanical studies have suggested that it accelerates trailing head
dissociation (1, 9, 29). However, our data with K413BIO does not
support this. The effective rate constant for dissociation,
ke, was in fact slower for dimeric
kinesin than for a monomeric construct. As we have shown (Equation 1),
ke is a composite rate constant and depends on
the rates of ATP hydrolysis (kh) and microtubule
dissociation (kdMT). Direct measurements using chemical quench methods have consistently shown that, although kh is ~100 s1 for dimeric
constructs, it is considerably faster for monomers, with estimates
placing it at >250 s
1 (2, 20, 22). On the other hand,
our previous studies with K349 show that the rate of docking of the
neck linker places an upper limit on kh of
~800 s
1 (23). We have performed fitting to the data in
Fig. 3 to obtain values of kdMT for
monomeric and dimeric kinesins, using values of
kh of 100 s
1 for K413 and the
limiting values of 300 s
1 and 800 s
1 for
K349. These reveal values of kdMT of
122 ± 27 s
1 for K413 and 143 ± 16 s
1 (kh = 300) and 111 ± 10 s
1 (kh = 800) for K349. Thus, even
when correcting for differences in the kinetics of ATP hydrolysis
between monomeric and dimeric constructs, we find that
kdMT is relatively unaffected by forward strain.
However, K413BIO is a mutant construct that has eliminated all the surface-reactive cysteines. Hence, it may still be possible that forward strain has some effect on the processivity of wild-type kinesin. Our kinetic characterizations of K349 and K413 has shown that only one step in the mechanochemical cycle is affected (18, 23, and this work). This is the rate of ADP-induced dissociation, which is accelerated 19-fold compared with wild-type (Fig. 4). Furthermore, although K413BIO is processive and has near wild-type in vitro velocities, its mean run length is reduced ~2- to 3-fold (Fig. 2). Thus, it is possible that forward strain may accelerate trailing head dissociation in wild-type kinesin. However, even if this were the case, the degree of acceleration would be relatively small, amounting to no more than a factor of 2 or 3. This degree of acceleration is almost identical to the value predicted by Uemura et al. (29) using unbinding force measurements. Thus, our data with K413BIO clearly shows that, although a forward strain-induced dissociation mechanism may modulate the length of a processive run, it is not required for processivity.
In summary, this study has shown that the internal strain generated by
kinesin during its mechanochemical cycle provides a mechanism that
supports processivity. The major effect of strain is to markedly slow
ATP binding to the leading head, an effect that guarantees that the two
motor domains remain out of phase of each other during multiple
mechanochemical cycles. Although strain may also accelerate
dissociation of the trailing head, our results show that this effect is
not necessary for processive movement. Finally, the
strain-dependent mechanism that we describe may have more
general applicability. Other molecular motors, such as myosins V and VI
are also processive (34-36). Like kinesin, these motors need a
mechanism to keep their individual motor units out of phase
enzymatically to prevent premature dissociation from actin. Several of
the methods developed in this study are directly applicable to these
motors and may be useful in future studies to elucidate the mechanisms
underlying their processivity.
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ACKNOWLEDGEMENT |
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We thank Dr. Shin'ichi Ishiwata (Waseda University, Japan) for his thoughtful review of our manuscript.
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FOOTNOTES |
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* 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.
§ To whom correspondence should be addressed: Dept. of Neurology, University of Alabama at Birmingham, MEB 510, 1813 6th Ave. South, Birmingham, AL 35294. Tel.: 205-934-1813; Fax: 205-975-7546; E-mail: stevensr@uab.edu.
Published, JBC Papers in Press, March 6, 2003, DOI 10.1074/jbc.M300849200
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ABBREVIATIONS |
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The abbreviations used are: PIPES, 1,4-piperazinediethanesulfonic acid; AMPPNP, 5'-adenylyl-b,g-imidodiphosphate; 2'dmD, 2'-deoxy-mant-ADP; 2'dmT, 2'-deoxy-mant-ATP; FRET, fluorescence resonance energy transfer; MDCC, N-[2-(1-maleimidyl)ethyl]-7-(diethylamino)coumarin-3-carboxamide; TMR, tetramethyl rhodamine maleimide; AEDANS, S-(((2-acetyl)amino)ethyl)aminonaphthalene-1-sulfonic acid.
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