From the Department of Biochemistry and Biophysics,
University of California, San Francisco, California 94143 and
¶ Cytokinetics, Inc., South San Francisco, California
94080
Received for publication, January 16, 2001, and in revised form, April 24, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Success of mitosis depends upon the coordinated
and regulated activity of many cellular factors, including kinesin
motor proteins, which are required for the assembly and function of the
mitotic spindle. Eg5 is a kinesin implicated in the formation of the
bipolar spindle and its movement prior to and during anaphase. We have determined the crystal structure of the Eg5 motor domain with ADP-Mg
bound. This structure revealed a new intramolecular binding site of the
neck-linker. In other kinesins, the neck-linker has been shown to be a
critical mechanical element for force generation. The neck-linker of
conventional kinesin is believed to undergo an ordered-to-disordered
transition as it translocates along a microtubule. The structure of Eg5
showed an ordered neck-linker conformation in a position never observed
previously. The docking of the neck-linker relies upon residues
conserved only in the Eg5 subfamily of kinesin motors. Based on this
new information, we suggest that the neck-linker of Eg5 may undergo an
ordered-to-ordered transition during force production. This
ratchet-like mechanism is consistent with the biological activity of Eg5.
Prior to the separation of sister chromatids in anaphase,
duplicated centrosomes are repositioned to opposite sides of the cell,
forming the mitotic spindle as they move. Centrosome separation is
dependent upon numerous proteins, including Eg5, a kinesin motor (1).
Eg5 slides the microtubules of the developing spindle past each other,
thereby pushing the centrosomes apart (2). This outward force is
balanced by other kinesin motors that provide an inward force (3,
4).
Elucidation of the specific roles played by Eg5 in this process has
been aided by the discovery of a small, cell-permeable molecule that
selectively inhibits Eg5 activity. This compound was named monastrol
because its presence causes the formation of a mono-astral spindle by
inhibiting centrosome separation (5, 6). The addition of monastrol
after bipolar spindle formation caused the spindle to collapse,
indicating that a force is constantly required to maintain spindle
integrity (6). In addition to its role as a cell biological reagent,
monastrol and its derivatives may be useful in the clinical setting as
anti-mitotic agents.
At least one Eg5 homologue has been found in every eukaryote (called
BimC in Aspergillus (7), cut7 in
Schizosaccharomyces pombe (8), cin8p in
Saccharomyces cerevisiae (9), Klp61F in
Drosophila (10) and Eg5 in Xenopus (11, 12) and
humans (13)). These kinesins and other homologues (identified by
sequence similarity) comprise the KinN2 kinesin subfamily (14). They share slow, plus end-directed, nonprocessive movement (15, 16), and a
unique homotetrameric structure (17). Like conventional kinesin,
two motor domains form a dimer via association of their stalks.
However, in a KinN2 motor, two dimers interact, anti-parallel to each
other, to form a rod with two motor domains at each end, a structure
often referred to as "bipolar." Both ends interact with
microtubules, bundling and sliding them past each other. The activity
of these proteins is restricted to the mitotic spindle and is
controlled, at least in part, by phosphorylation at a conserved C-terminal region by p34cdc2 (13, 18).
Recently, a general mechanism used by kinesin motors to produce
motility has been proposed (19). This model is based upon alternating
cycles of weak and strong microtubule binding that are dependent upon
whether ATP or ADP is bound to the protein. When ATP is hydrolyzed to
ADP, the affinity for the microtubule is weakened and the motor
releases. When ATP re-binds, a series of conformational changes take
place which trigger the movement of a mechanical element. This
positions the motor closer to the next binding site on the microtubule,
where it will bind tightly, allowing a step forward to take place.
Although the same general scheme appears to be utilized by all kinesin
motors, different kinesins perform numerous different activities in the
cell. This wide variety of functions is the result of subtle
alterations within the motor domain and the placement of the motor
within the ultra-structure of the protein.
To better understand how Eg5 functions, we have determined
the structure of its motor domain. The structure of the Eg5 motor revealed a unique conformation of the mechanical element. Unlike conventional kinesin, where the mechanical element is disordered in
the ADP state, the Eg5 mechanical element is structured in a
position not observed in any other kinesin. This observation may help
explain how Eg5 can work in arrays to efficiently slide microtubules
and why Eg5 is not a processive motor.
Cloning and Purification--
A fragment of Eg5 was amplified
from human placenta cDNA (CLONTECH) by
polymerase chain reaction using Pfu turbo
(Stratagene). The region encoding residues 1-491 was subcloned into
pET23d (Novagen) containing a Myc epitope and a 6-His tag. The
construct expressing untagged Eg5 (residues 1-368) used in this
study was created by introducing a stop codon 3' of the codon for K368
using QuickChange site-directed mutagenesis kit (Stratagene). Protein
expression in Escherichia coli BL21(DE3) cells was induced
with 0.5 mM
isopropyl-1-thio- Crystallization, Data Collection, and Data
Processing--
Crystals of Eg5 were grown by the sitting drop vapor
diffusion method. A 10 mg/ml protein solution was mixed 1:1 with and equilibrated against well solution (18% PEG-3350, 100 mM
PIPES (pH 6.8), 200 mM NaNO3). Imperfect
crystals that grew in about 2 days at 4 °C were crushed and used to
seed 1:1 drops of 10 mg/ml Eg5 and a modified well solution (15%
PEG-3350, 100 mM MES (pH 5.6), and 200 mM
NaNO3). Under these conditions, 100 × 100 × 50 µm crystals grew within a week. The crystals were cryopreserved by
transferring to well solution containing 15% glycerol and immersing in
liquid nitrogen.
Three independent data sets collected at Stanford Synchrotron Radiation
Laboratory (SSRL) (beamline 9-1;
The structure was solved by molecular replacement methods with the
program CNS (21) using the structure of KAR3 (22) as a search
model. The asymmetric unit contained two copies of Eg5, and both
monomers were refined independently by repeated cycles of manual
fitting in QUANTA (Molecular Simulations, Inc.) followed by simulated
annealing, B-factor refinement, and minimization in CNS. To reveal any
differences between the two monomers, noncrystallographic symmetry was
not employed during the refinement. Residues 1-15, 271-279, and
366-368 in the first monomer and residues 1-15, 271-284, and
366-368 in the second were not observed in the electron
density. The model also contains 351 waters, two Mg-ADP complexes, and two nitrate ions. The final Rcryst = 21.7% and
Rfree = 25.5%.
The structure of Eg5 was solved by molecular replacement methods
using the KAR3 motor (22) as a model. Details of the data collection
and refinement are presented in Table I.
The refined Eg5 model revealed a protein with the general features
expected of a kinesin motor, with six major
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-galactopyranoside, and cells were
harvested after 4 h of growth at 25 °C. Frozen cells were
thawed into 50 mM
PIPES1 (pH 6.8), 2 mM MgCl2, 1 mM EGTA, 1 mM ATP, and 1 mM Tris-(2-carboxyethyl)phosphine (TCEP) HCl and lysed in a microfluidizer. The lysate was
clarified by centrifugation and applied to a SP-Sepharose column
(Amersham Pharmacia Biotech). Protein was eluted with a linear gradient of 0-250 mM NaCl. Eg5 was identified in fractions by
SDS-polyacrylamide gel electrophoresis analysis, and the pooled
fractions were applied to Mono-S and Mono-Q columns (Amersham Pharmacia
Biotech), which were developed as the SP-Sepharose column. The final
protein pool was >95% pure as judged by SDS-polyacrylamide gel
electrophoresis, mass spectroscopy, and isoelectric
focusing-electrophoresis. Microtubule-stimulated ATPase assays
(data not shown) revealed our preparation to be as active as that
described by Lockhart and Cross (15).
=0.983) were processed with
DENZO and SCALEPACK (20) and revealed a unit cell of 53.1 × 78.6 × 94.1 Å in the space group P21 with
= 93.8°. The data set was 94.9% complete to 2.1 Å (85% complete in
the highest resolution shell).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-sheets surrounded by
six
-helices (Fig. 1A). The
kinesin motor structure has been described as an arrowhead (23, 24)
with a nucleotide binding site at the wider end of the arrowhead. In
Eg5, the nucleotide binding site is occupied by Mg-ADP.
Data collection and refinement statistics
View larger version (55K):
[in a new window]
Fig. 1.
Structure of the Eg5 motor domain and
its comparison to conventional KHC motor domain. A, the
Eg5 motor with major -sheets (gray) and
-helices
(gold) labeled as described for conventional kinesin by Kull
et al. (23). One molecule of Mg-ADP is located in
the nucleotide binding site. B, a sequence alignment of Eg5
and KHC motor domains. Structural elements are shown in gray
for
-sheets and gold for
-helices. Disordered regions
are indicated by light gray lettering. C,
overlay of the
-sheets of Eg5 (in pink) and KHC (in
blue). The remainder of the structure is Eg5. D,
overlay of helix
2, including a larger loop in Eg5 than in KHC.
E, overlay of helix
3 and the switch I regions of Eg5 and KHC. Note that switch I of Eg5 is found as a coil, whereas in KHC
the same region is a short
-helix. F, overlay of
helix
4, the switch II helix. Note that this helix is slightly
longer in Eg5 than in KHC. G, overlay of helix
5, a
region important for microtubule interactions. H,
overlay of helix
6, revealing a novel position of the neck-linker in
Eg5. In KHC, the neck-linker is disordered.
This structure of Eg5 is the first structure of a Kin N2 motor. However, it is the ninth structure of a motor from the kinesin superfamily. In analyzing the structure, we noticed many differences between Eg5 and other kinesin motors. Our challenge was to determine which of these features are important for Eg5 function in particular and which are important for kinesin motor function in general.
Here, we present a detailed comparison of Eg5 and one kinesin motor,
KHC. We chose kinesin heavy chain (KHC) because it shares 40% identity
with Eg5, the highest of all the motors with structures available (Fig.
1B). The comparison of divergent structures is facilitated
by superimposing elements that are known to be conserved in the
structures and then examining the differences this highlights in other
regions. The phosphate binding region (P-loop) is conserved in all
kinesin structures. Therefore, we used the P-loop region (Eg5 105-113,
KHC 85-93) to align Eg5 with KHC. To more easily view the results of
the comparison, Fig. 1, CH, presents a single region of
the overlapping structures at a time, with the Eg5 structure shown in
pink and the KHC structure in blue.
After superimposing KHC and Eg5, it becomes apparent that the core
-sheets are almost identical in the two structures (Fig. 1C). However, there is a region of divergence near the tip
of the protein, leading to the appearance of a slight tilting and lengthening of Eg5 with respect to KHC. The recently determined structure of the Kif1A motor shares this feature with Eg5 (25). It is
believed that the tip of the kinesin arrowhead may play an important
role in transient interactions with the microtubule during force
production. Therefore, this structural alteration may be a factor in
determining the affinity of kinesin motors for the microtubule rather
than a transient change that occurs as part of the ATP hydrolysis cycle.
Of the six helices that surround the core -sheets, helix
1 did
not appear different in the Eg5 and KHC structures (not shown). However, there was a dramatic difference in helix
2. This helix is
interrupted by a loop in all kinesins, and its function is not known.
As seen in Fig. 1D, this loop is larger in Eg5 than in KHC.
The size of the loop is variable among kinesin family members, but it
is largest in the Kin N2 family (see Fig. 3A for a limited
sequence comparison). This loop is located on the opposite face of the
protein from that which binds to the microtubule and is in proximity
to, but not a part of, the nucleotide binding site. One idea, which
remains unsubstantiated, is that this loop may somehow regulate motor
activity, perhaps by interacting with other proteins.
Kinesin motility is based upon nucleotide state sensing. In this way,
small changes (the presence or absence of the -phosphate) can be
transmitted to and amplified in other parts of the structure. This
activity relies upon loop components of the switch I and switch II
regions. When ATP binds, these loops make direct contact with the
-phosphate of ATP and also form interactions with each other
(22-26). These adjustments cause a cascade of secondary movements in
the protein, including docking of the neck-linker mechanical element
and increasing microtubule affinity. When ATP is hydrolyzed to ADP, the
interactions among switch I, switch II, and the nucleotide are
lost. This reverses the conformational changes that took place upon ATP
binding, resulting in the release of the neck-linker and a decrease in
microtubule affinity.
The switch I region is found at the end of helix 3. In KHC, switch I
is a short
-helix, whereas in Eg5 switch I is a loop (Fig.
1E). Is this structural difference the basis for the
functional differences between Eg5 and KHC? We think it is more likely
that the two structures may represent two different states that all kinesin motors assume at some point during force generation. As mentioned above, the
-phosphate acts to bring together switch I and
switch II. However, both the Eg5 and KHC structures contain ADP and
therefore no
-phosphate. Therefore, switch I with ATP bound likely
assumes the same conformation in all kinesins, whereas without the
-phosphate "tether" this region is flexible. This prediction is
supported by the many different positions of this region observed in
other kinesin motors (22-26). However, it is also possible that the
different switch I conformations may reflect real differences among
kinesin family members. Additional structures and biochemical
experiments should be able to answer this question in the future.
In addition to nucleotide sensing by switch I, the switch II region is
also critical for nucleotide sensing and force production. Switch II
consists of helix 4 (often referred to as the switch II helix or the
relay helix) and a loop that interacts with the nucleotide. A portion
of this loop is disordered and therefore not observed in the electron
density map of Eg5 and most other kinesins (22-26). Helix
4 of Eg5
and KHC differ in two ways. In Eg5, helix
4 is one and one-half
turns longer and slightly rotated with respect to helix
4 in KHC
(Fig. 1F).
The helix extension observed in Eg5 is formed by ordering a region of
the switch II loop that is disordered in the KHC structure. A longer
4 is also seen in the structure of Kif1A but only with ADP (not the
ATP analogue AMP-PCP) bound (25). In that study, the length of helix
4 was found to be dependent upon the nucleotide state. However, by
looking at all of the kinesin structures available, it becomes obvious
that there is not a strict correlation with helix
4 length and
nucleotide state (22-26). As a case in point, the two molecules of Eg5
in the asymmetric unit of the current crystal structure differ in the
length of the
4 helix although they are nearly identical elsewhere
(not shown). Therefore, it appears as though the length of
4 may
change during ATP hydrolysis but that this change requires a low energy
input in the absence of microtubules. In other words, crystals may trap
structural intermediates that occur within motors in the absence of
microtubules. This reflects a flexibility in this region that may be
required for helix
4 to adjust its position in response to ATP binding.
In addition to the length of helix 4, the position of this helix is
a key component in force generation. All known kinesin structures can
be classified as switch II helix-up or switch II helix-down.
Neck-linker docking is inhibited in the switch II helix-down position,
which is believed to be reflective of the ADP-bound state. Although the
position of
4 is slightly different in Eg5 and KHC, both of these
structures are part of the switch II helix-down group (25). Again, we
attribute the slightly different positions to trapping of these mobile
elements in the crystal structures.
In addition to switch I and switch II, helix 5 and its neighboring
loops undergo nucleotide-dependent rearrangements.
Structural changes in this region likely effect microtubule binding,
because this region is an important surface for interaction between the motor and the microtubule. The differences observed between Eg5 and KHC
in this area (Fig. 1G) may again indicate flexibility of
this region in the ADP-bound state in the absence of microtubules. Alternatively, this region may contribute to differences in
microtubule binding observed in the two proteins.
Helix 6 is virtually identical in the two proteins (Fig.
1H). However, the region at the end of helix
6 is very
different in the two structures. In the KHC model, there is no electron density in this region, whereas in Eg5 electron density is clearly visible. (Fig. 2). This region, termed
the neck-linker, is a critical mechanical component of the force
production cycle of kinesins and also serves to attach the motor to the
coiled-coil stalk (27). In recent years, much attention has been paid
to the neck-linker of kinesin motors. In all kinesin crystal structures
except Eg5, the neck-linker is either disordered (as in KHC) or found
docked to the motor, parallel to the longest motor edge (as in rat
KHC). The neck-linker position directly correlates with the switch II helix position. The only exception is the Eg5 neck-linker, which adopts a position perpendicular to the long edge of the protein. The neck-linker position is not stabilized by crystal contacts but
rather by a series of hydrogen bonds between the neck-linker and the
1/
2 lobe it docks against.
|
In addition to comparing the structure of Eg5 to the structures of other kinesin motors, we were interested in understanding how specific residues may play roles in specializing the activity of the different families of motor domains. In this analysis, we identified a subset of residues conserved among the KinN2 family that was not conserved in other kinesins. A sequence alignment of selected motor domains is presented in Fig. 2A, with Kin N2-specific residues highlighted in red. These residues were mapped onto the Eg5 structure and are shown in Fig. 2B.
Surprisingly, a number of the KinN2-conserved residues mapped to regions of Eg5 involved in neck-linker docking. Two residues in the neck-linker are specific to the KinN2 family, Lys-364 and Pro-365. These residues interact with residues Glu-49, and Thr-67, which are also conserved in the KinN2 family (Fig. 2C). The identification of these KinN2-conserved interactions lends more substance to the argument that the neck-linker conformation seen in Eg5 is not merely an experimental artifact but is a potential intermediate in the mechanochemical cycle.
This analysis identified other residues conserved specifically in the
Kin N2 family that may be important for the interaction of the
neck-linker with the core of the protein in other nucleotide states.
Although we do not yet have structural information on the ATP bound
state of Eg5, we can model where the neck-linker will go based on
information available from other kinesin structures (22-26). In the
ADP bound state, the parallel conformation of the neck-linker is
precluded by the down position of the switch II helix. When ATP binds,
the switch II helix moves up and the neck-linker is able to zip down
the side of the motor core. This position is shown in Fig.
2C. Interestingly, some of the residues on Eg5 that would
need to move to allow the neck-linker to dock are specifically conserved in the KinN2 family (Val-303, Arg-327, and Thr-328). Further
down the predicted pathway for the neck-linker, another group of
KinN2-specific residues is encountered at the "tip" of the protein
(Gly-252, Glu-253, Glu-254). These may represent the last specific
contact site the neck-linker makes with the motor core in the ATP bound
state. Future experiments will determine the contribution of these
conserved residues located in interesting regions.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The result of this analysis was that most of the differences between Eg5 and KHC are likely the result of capturing the motors in slightly different stages of the movements that they go through during a force generation cycle. Kinesin motors are built to move and contain modules that move in a regulated and coordinated manner during force generation. Crystal structures are useful in that they capture one particular state that a motor may assume during force production. This information is valuable only in the context of understanding that other conformations did and will exist immediately before and immediately after the particular state a crystal has trapped.
By comparing Eg5 to all other kinesin structures, only one feature stood out as truly unique. This was the position of the neck-linker, docked perpendicular to the motor in the presence of Mg-ADP. This conformation was not seen to be stabilized by crystal contacts and involved conserved residues. Taken together, these observations suggest that perpendicular neck-linker docking may play a role in the force generation cycle of Eg5.
Although Eg5 contains a motor domain similar to that found in all other
kinesins, it has evolved to perform a unique biological function. Unlike conventional kinesin, which walks along stationary microtubules, Eg5 has the job of putting microtubules into motion. A
model highlighting possible differences in the mechanisms of these two
types of kinesins is shown in Fig. 3. Eg5
works in arrays along the microtubule, and therefore to symbolize this,
two motors are illustrated. However, for the sake of clarity, only one
head is shown at either end of the Eg5 bipolar structure (Fig.
3A). With ADP bound, the Eg5 neck-linker (shown in
red) exits the motor core perpendicular to the long edge of
the motor, as seen in the structure reported here. When microtubules
are encountered, ADP release is stimulated, and ATP readily binds the
empty nucleotide binding site. This causes a cascade of conformational
changes that result in the "zipping" of the neck-linker down the
side of the microtubule-attached motor core. Because both ends of Eg5 are interacting with microtubules, the motors themselves cannot move.
Therefore, the microtubule must move as the neck-linker assumes a
position parallel to the length of the motor. When ATP is hydrolyzed,
the neck-linker is pushed out of the down position and the affinity of
the motor for the microtubule is decreased. Because the microtubule has
moved as a result of the previous ATP binding, release of the
microtubule brings the motor into position for binding the next site on
the microtubule.
|
The establishment of defined positions of the neck-linker in both the ADP and the ATP bound states may allow control over the direction and efficiency of microtubule sliding. If the neck-linker remained disordered in the ADP-bound state, each time the neck-linker "zipped down" in the presence of ATP, the microtubule might be propelled in a different direction. Because multiple motors bind the moving microtubules, randomization of the direction of the forces would cause a "canceling out" of the activities of many of the motors. Additionally, the structured nature of both the ADP and ATP bound states may help to maintain some of the rigidity that would be needed to slide microtubules efficiently. This could be referred to as a "ratcheting" mechanism, where the two binding sites are utilized to define the space through which the mechanical element can move and also to provide stability at both limits of its movement.
The model illustrated in Fig. 3A could be amended in a number of ways that may more accurately represent what occurs in cells. For example, how the four heads are coordinated with each other is not understood. Additionally, microtubules with the same polarity may interact with both ends of an Eg5 motor, resulting in microtubule bundling rather than sliding in opposite directions. Finally, it may be useful to envision an array of Eg5 motors, each moving the microtubule a distance less than that required for finding the next binding site. In this instance, movement of the microtubule by other motors may help microtubule release and facilitate the next encounter with the next binding site. The Eg5 structure presented here will allow these and other mechanistic details to be addressed.
Muscle myosin proteins use a similar ordered-to-ordered transition to generate directed movement (19). Like Eg5, myosin II acts in arrays which work together to move actin. Do other kinesins utilize an ordered-to-ordered neck-linker transformation? Crystal structures of other kinesins have revealed either a neck-linker in the zipped-down conformation or as disordered regions (22-27). Interestingly, electronmicroscopy studies of KHC have shown that although the neck-linker position was variable in the presence of ADP, a few positions were observed more frequently than others (28). One of these sites is consistent with the position of the neck-linker observed in the current structure of Eg5 (Fig. 4D in Ref. 28). Perhaps a stable neck-linker conformation in the presence of ADP is more transient in other kinesins, or it may not be as prone to forming as it is in Eg5.
Alternatively, a flexible neck-linker may be an adaptation required by two-headed "walking" kinesins as shown in Fig. 3B. Neck-linker flexibility may be important for the trailing motor to be propelled forward to the next binding site on the microtubule. The "stepping" or "hand-over-hand" processive mechanism proposed for conventional kinesin suggests that the neck-linker is flexible and can assume a number of different positions. Eg5 has been shown to be a nonprocessive motor (16). Therefore, neck-linker stability in both the ADP and ATP bound states of Eg5 may preclude this motor from being processive. Perhaps the need for controlled, efficient sliding of the mitotic spindle outweighs the benefits of processivity afforded by a flexible neck-linker. Alternatively, processivity may have been an evolutionary outgrowth of simpler, nonprocessive motors.
Recently, electronmicroscopy studies of Eg5 bound to
microtubules revealed a decreased flexibility of the two heads when
compared with similar experiments with KHC and other processive dimeric motors (29). This could reflect ordering of the neck-linker as
discussed here. Future experiments, including electronmicroscopy labeling of the neck-linker, site-directed mutagenesis, and
spectroscopic assays will determine whether the observed neck-linker
ordering is truly critical for Eg5 activity.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank M. Yu for help with protein expression and initial purification. E. Sablin, M. Butte, L. Brinen, C. Sindelar, M. Vinogradvia, S. Wang, R. Vale, and R. Cooke provided enormous help with the crystallography and scientific discussions. We also acknowledge the support staff at SSRL.
![]() |
FOOTNOTES |
---|
* This work was supported by grants from the National Institutes of Health (to R. F.) and the Damon Runyon-Walter Winchell Cancer Research Fund (to J. T.).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.
The atomic coordinates and the structure factors (code 1II6) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
§ To whom correspondence should be addressed: Dept. of Biochemistry and Biophysics, University of California, Box 0448, San Francisco, CA 94143. Tel.: 415-502-5848; Fax: 415-476-1902; E-mail: turner@msg.ucsf.edu.
Published, JBC Papers in Press, April 27, 2001, DOI 10.1074/jbc.M100395200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: PIPES, 1,4-piperazinediethanesulfonic acid; MES, 4-morpholineethanesulfonic acid; KHC, kinesin heavy chain.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Heald, R. (2000) Cell 102, 399-402[Medline] [Order article via Infotrieve] |
2. |
Sharp, D. J.,
McDonald, K. L.,
Brown, H. M.,
Matthies, H. J.,
Walczak, C.,
Vale, R. D.,
Mitchison, T. J.,
and Scholey, J. M.
(1999)
J. Cell Biol.
144,
125-138 |
3. | Walczak, C., Vernos, I., Mitchison, T. J., Karsenti, E., and Heald, R. (1998) Curr. Biol. 8, 903-913[Medline] [Order article via Infotrieve] |
4. | Sharp, D. J., Yu, K. R., Sisson, J. C., Sullivan, W., and Scholey, J. M. (1999) Nat. Cell Biol. 1, 51-54[CrossRef][Medline] [Order article via Infotrieve] |
5. |
Mayer, T. U.,
Kapoor, T. M.,
Haggerty, S. J.,
King, R. W.,
Schreiber, S. L.,
and Mitchison, T. J.
(1999)
Science
286,
971-974 |
6. |
Kapoor, T. M.,
Mayer, T. U.,
Coughlin, M. L.,
and Mitchison, T. J.
(2000)
J. Cell Biol.
150,
975-988 |
7. | Enos, A. P., and Morris, N. R. (1990) Cell 60, 1019-1027[Medline] [Order article via Infotrieve] |
8. | Hagan, I., and Yanagida, M. (1990) Nature 347, 563-566[CrossRef][Medline] [Order article via Infotrieve] |
9. | Hoyt, M. A., He, L., Loo, K. K., and Saunders, W. S. (1992) J. Cell Biol. 118, 109-120[Abstract] |
10. | Heck, M. M., Peereira, A., Pesavento, P., Yannoni, Y., Spradling, A. C., and Goldstein, L. S. (1993) J. Cell Biol. 123, 655-679 |
11. | LeGuellec, R., Paric, J., Couturier, A., Roghi, C., and Philippe, M. (1991) Mol. Cell. Biol. 11, 3395-3398[Medline] [Order article via Infotrieve] |
12. | Swain, K. E., LeGuellec, K., Philippe, M., and Mitchison, T. J. (1992) Nature 359, 540-543[CrossRef][Medline] [Order article via Infotrieve] |
13. | Blangy, A., Lane, H. A., d'Herin, P., Harper, M., Kress, M., and Nigg, E. A. (1995) Cell 83, 1159-1169[Medline] [Order article via Infotrieve] |
14. | Vale, R. D., and Fletterick, R. J. (1997) Annu. Rev. Cell Dev. Biol. 13, 745-777[CrossRef][Medline] [Order article via Infotrieve] |
15. | Lockhart, A., and Cross, R. A. (1996) Biochemistry 35, 2365-2373[CrossRef][Medline] [Order article via Infotrieve] |
16. | Crevel, I. M., Lockhart, A., and Cross, R. A. (1997) J. Mol. Biol. 273, 160-170[CrossRef][Medline] [Order article via Infotrieve] |
17. | Kashina, A. S., Baskin, R. J., Cole, D. G., Wedman, K. P., Saxton, W. M., and Scholey, J. M. (1996) Nature 379, 270-272[CrossRef][Medline] [Order article via Infotrieve] |
18. | Swain, K. E., and Mitchison, T. J. (1995) Proc. Natl. Acad. U. S. A. 92, 4289-4293[Abstract] |
19. |
Vale, R. D.,
and Milligan, R. A.
(2000)
Science
288,
88-95 |
20. | Otwinowski, Z., and Minor, W. (1997) in Methods in Enzymology (Carter, C. W. , and Sweet, R. M., eds), Vol. 276 , pp. 307-326, Academic Press, San Diego |
21. | Brunger, A. T., Kuriyan, J., and Karplus, M. (1987) Science 235, 458-460 |
22. | Gulick, A. M., Song, H., Endow, S. A., and Rayment, I. (1998) Biochemistry 37, 1769-1776[CrossRef][Medline] [Order article via Infotrieve] |
23. | Kull, F. J., Sablin, E. P., Lau, R., Fletterick, R. J., and Vale, R. D. (1996) Nature 380, 550-555[CrossRef][Medline] [Order article via Infotrieve] |
24. | Sablin, E. P., Kull, F. J., Cooke, R., Vale, R. D., and Fletterick, R. J. (1996) Nature 380, 555-559[CrossRef][Medline] [Order article via Infotrieve] |
25. | Kikkawa, M., Sablin, E. P., Fletterick, R. J., and Hirokawa, N. (2001) Nature 411, 439-445[CrossRef][Medline] [Order article via Infotrieve] |
26. | Sack, S., Muller, J., Marx, A., Thormahlen, M., Mandelkow, E. M., Brady, S. T., and Mandelkow, E. (1997) Biochemistry 36, 16155-16165[CrossRef][Medline] [Order article via Infotrieve] |
27. | Case, R. B., Rice, S., Hart, C. L., Ly, B., and Vale, R. D. (2000) Curr. Biol. 10, 157-160[CrossRef][Medline] [Order article via Infotrieve] |
28. | Rice, S., Lin, A. W., Safer, D., Hart, C. L., Naber, N., Carragher, B. O., Cain, S. M., Pechatnikova, E., Wilson-Kubalek, E. M., Whittaker, M., Pate, E., Cooke, R., Taylor, E. W., Milligan, R. A., and Vale, R. D. (1999) Nature 402, 778-784[CrossRef][Medline] [Order article via Infotrieve] |
29. |
Hirose, K.,
Henningsen, U.,
Schliwa, M.,
Toyoshima, C.,
Shimizu, T.,
Alonso, M.,
Cross, R. A.,
and Amos, L. A.
(2000)
EMBO J.
19,
5308-5314 |