* Institute of Cell Biology, Swiss Federal Institute of Technology, ETH-Hönggerberg, CH-8093 Zürich, Switzerland; and Max-Planck-Unit for Structural Molecular Biology, D-22603 Hamburg, Germany
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Abstract |
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We have decorated microtubules with monomeric and dimeric kinesin constructs, studied their
structure by cryoelectron microscopy and three-dimensional image reconstruction, and compared the results with the x-ray crystal structure of monomeric and
dimeric kinesin. A monomeric kinesin construct
(rK354, containing only a short neck helix insufficient
for coiled-coil formation) decorates microtubules with
a stoichiometry of one kinesin head per tubulin subunit
(-
-heterodimer). The orientation of the kinesin head
(an anterograde motor) on the microtubule surface is
similar to that of ncd (a retrograde motor). A longer kinesin construct (rK379) forms a dimer because of the
longer neck helix forming a coiled-coil. Unexpectedly, this construct also decorates the microtubule with a stoichiometry of one head per tubulin subunit, and the orientation is similar to that of the monomeric construct.
This means that the interaction with microtubules
causes the two heads of a kinesin dimer to separate sufficiently so that they can bind to two different tubulin
subunits. This result is in contrast to recent models and
can be explained by assuming that the tubulin-kinesin
interaction is antagonistic to the coiled-coil interaction
within a kinesin dimer.
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Introduction |
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INTRACELLULAR transport, mitosis, flagellar beating,
and many other motile phenomena in cells are based
on the interaction between microtubules and their
motor proteins such as kinesins or dynein. The diverse
family of kinesin-related motor proteins can be subdivided
into two main classes, depending on whether they move
their cargoes (e.g., vesicles, organelles) towards the plus
end or the minus end of microtubules (anterograde or retrograde transport in axons). Their best known representatives are kinesin itself (a plus end-directed motor) and ncd
(a minus end-directed motor). These proteins consist of
several domains, the "head" or motor domain (containing
the microtubule and ATP binding sites), a "stalk," which is
largely -helical and causes two kinesin chains to dimerize
via a coiled-coil interaction, and a "tail" connecting the
stalk to the cargo (for reviews see Goldstein, 1993
; Brady, 1995
; Cole and Scholey, 1995
).
The core motor domain contains ~330 residues. Its
x-ray structure has been solved recently for human kinesin
and Drosophila ncd; despite their opposite directionality
the two structures are remarkably similar (Kull et al., 1996;
Sablin et al., 1996
). There is a central
sheet of eight
strands sandwiched between three
helices on either side.
The residues interacting with microtubules are confined to
the "rear" surface (containing loops L8, L12, and helices
4-
6; see Kull et al., 1996
; Woehlke et al., 1997
), whereas
the "front" surface contains the nucleotide-binding loops
(for review see Vale and Fletterick, 1997
). A recent x-ray structure of rat kinesin in the monomeric and dimeric state
has revealed additional features (Kozielski et al., 1997a,b;
Sack et al., 1997
). Notably, the NH2-terminal entry and
COOH-terminal exit of the polypeptide chain into the
core motor domain have a
strand structure, forming an
additional antiparallel
sheet (
0,
9) or integrated into
the central sheet (
10). Conserved residues that play a
role in the directionality of the motors (Case et al., 1997
)
are located around
9 for kinesin and near
0 for ncd. The
neck helix
7 (Ala339-Trp370) extends from the lower tip
of the structure, roughly within the plane of the core sheet.
Judging from model peptides, the first half of the neck helix has a low coiled-coil potential, presumably because of
its unusually high charge and nonstandard heptad residues, but the second half is similar to a leucine zipper and
causes tight association (Morii et al., 1997
; Tripet et al.,
1997
; Thormählen et al., 1998b
). Accordingly, the construct rK354 is monomeric in solution and in the crystal,
whereas rK379 is dimeric (Kozielski et al., 1997a
).
The interaction between kinesin and tubulin has been
probed by various biochemical and biophysical methods.
Binding and cross-linking studies suggest that the motor
domain interacts mainly with -tubulin (Song and Mandelkow, 1993
; Kikkawa et al., 1995
), although some interactions also occur with
-tubulin (Walker, 1995
; Tucker
and Goldstein, 1997
). Reports on the stoichiometry of
binding have been controversial, claiming either one motor head bound per tubulin heterodimer (Harrison et al.,
1993
; Song and Mandelkow, 1993
), or two (Huang et al.,
1994
; Lockhart et al., 1995
). The latter value was interpreted within the framework of an "alternating catalysis"
model of ATP exchange (Hackney, 1994
; Gilbert et al., 1995
; Ma and Taylor, 1997
), whereby one head was firmly
bound (and unable to exchange ATP) whereas the other
was loosely tethered (and able to exchange ADP for
ATP). A recent reinvestigation of the stoichiometry with
scanning transmission electron microscopy, x-ray scattering, and other methods has confirmed the value of one
head per tubulin heterodimer (Thormählen et al., 1998a
;
Marx et al., 1998
), consistent with the results reported below. Measurements of the movement of kinesin-coated
beads along microtubules by laser trap microscopy has revealed a step size of 8 nm, equivalent to the axial spacing
of tubulin heterodimers (Svoboda et al., 1993
; Coppin et
al., 1996
; Hua et al., 1997
; Inoue et al., 1997
; Schnitzer and
Block, 1997
). Depending on the model of motion ("walking," "tightrope," and others; Cross, 1995
; Howard, 1996
)
the observed step size for the kinesin dimer implies that a
single head must be able to translocate up to 16 nm. This is
remarkably large considering the head's small dimensions
of ~6 × 3.5 × 3 nm.
Several image reconstructions of microtubules decorated with kinesin and ncd have been presented recently.
There is good agreement on the shape and position of monomeric constructs (Hirose et al., 1995, 1996
; Hoenger et al.,
1995
; Kikkawa et al., 1995
; Arnal et al., 1996
; Hoenger and
Milligan, 1997
; Sosa et al., 1997b
; for comparison see Figs.
2 and 3 below). Roughly speaking, if we view a microtubule in standard orientation (with the fast-growing "plus"
end up) the tubulin subunits show an anticlockwise slew, the bound motor heads are attached mostly to
-tubulin
(with some overlap to
-tubulin) with the long axis
roughly vertical, and with a clockwise displacement relative to the tubulin subunits (similar to Fig. 4 below). This
shape could be fitted well with the x-ray structure of monomeric ncd, showing the sequences implicated in microtubule interactions (loops L2, L8, L11, L12, helices
4, and
6 on the rear surface) touching the microtubule surface
(Sosa et al., 1997a
).There was however a curious discrepancy when dimeric constructs of kinesin or ncd were used
(Arnal et al., 1996
; Hirose et al., 1996
). Microtubules decorated with dimeric ncd showed two complete heads per tubulin heterodimer, one bound directly to the microtubule
surface (very similar to the monomeric constructs), the
second one attached to the first head, with an anticlockwise slew, and pointing towards the microtubule minus
end. This arrangement was consistent with the stoichiometry of two motor heads per tubulin subunit (Huang et al.,
1994
; Crevel et al., 1996
). In the case of dimeric kinesin,
however, the additional mass accounted for only ~20% of
a head, and it pointed towards the microtubule plus end.
This mass increase was interpreted to mean that a second head was present but only partially visible because of disorder. The differences between kinesin and ncd in the orientation of the second head were thought to reflect the
different mechanisms of motility (plus versus minus end
directed).
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An alternative explanation for kinesin, presented here,
is that there is only one bound head per tubulin heterodimer so that an additional mass observed with a
dimeric construct would be simply explainable by the longer
neck, rather than an additional but disordered head. We
find that the electron density decorating the microtubule
surface agrees well with the x-ray structure of the kinesin monomer, but not with the x-ray structure of the kinesin dimer. This implies that the two heads of dimeric kinesin
separate substantially upon binding to the microtubule lattice such that each head attaches to a different tubulin subunit. The orientation of the kinesin monomer is similar to
that of the ncd monomer described by Sosa et al. (1997a),
and the
-helical neck projects from the upper end of the
molecule in an anticlockwise direction. Thus the crest of a
protofilament appears to be embraced between loop L11
and the neck helix
7. The observed separation of the
heads would be sufficient to allow each head to make large
steps along the microtubule, in spite of the small head size.
The conserved "gearbox" residues that are important for
the directionality of motors (Case et al., 1997
; Henningsen
and Schliwa, 1997
) are located in the upper part of the
bound kinesin, near the head-neck junction.
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Materials and Methods |
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Assembly of Microtubules
Tubulin was prepared by phosphocellulose chromatography preceded by a
MAP-depleting step as described (Mandelkow et al., 1985). Microtubules
were polymerized in reassembly buffer (0.1 M Pipes, pH 6.9, 1 mM MgCl2,
0.5 mM EGTA, 1 mM DTT, 20 µM taxol) and 1 mM GTP from a solution
containing 6-15 mg/ml of PC-tubulin on incubation at 37°C for 20 min.
Expression of Kinesin
Plasmids coding for rat kinesin constructs were cloned and expressed as
described in Kozielski et al.(1997a). Briefly, to obtain pErK379 the
BamHI-SauI fragment of the rat kinesin gene (provided by S. Brady, University of Texas, Dallas, TX) was inserted into a pET-3a vector modified
to contain the same sites with a stop codon. Plasmid pErK354 was cloned by
PCR using plasmid pErK379 as a template. The plasmids were expressed
in Escherichia coli BL21 (DE3) cells. Recombinant rK354 and rK379
could be obtained in soluble form from the bacterial extracts. E. coli cells
were grown in Luria-Bertani (LB) medium and expression of kinesin was induced with 0.4 mM isopropylthio--D-galactoside (IPTG) at a cell density corresponding to A600 = 0.6-0.8. Cells were harvested after 16 h of induction at 25°C, resuspended in lysis buffer (50 mM Pipes, pH 6.9, 60 mM
NaCl, 1 mM MgCl2, 0.5 mM EGTA, 2 mM DTT, 1 mM PMSF), and then
lysed with a French press. Purification was done by ion exchange chromatography on phosphocellulose and MonoQ followed by gel filtration on a
G-200 Hiload 16/60 column (Pharmacia Biotechnology Inc., Piscataway, NJ).
Kinesin-Microtubule Binding Assays
Stoichiometries and dissociation constants were determined essentially as
described (Thormählen et al., 1998a). Briefly, taxol-stabilized microtubules at a concentration of 4 µM were incubated with kinesin (
20 µM kinesin heads) in the presence of 2 mM 5'-adenylylimidodiphosphate
(AMP-PNP)1. Microtubules with bound kinesin were pelleted in an ultracentrifuge (TL100; Beckman Instruments, Inc., Fullerton, CA) at 67,000 g and 22°C for 10 min. The pellets and supernatants were analyzed by SDS-PAGE. Coomassie blue-stained dried gels were digitized and evaluated
by using the software TINA (Raytest, Straubenhardt, FRG).
Decoration of Microtubules with Kinesin
Microtubules were polymerized for 20 min at 37°C in 80 mM Pipes, pH
6.8, 2 mM MgCl2, at a concentration of 2.5 mg/ml and in the presence of
5% (vol/vol) DMSO, 2 mM GTP, and 20 µM taxol. Polymerized tubulin
was washed twice by centrifugation in a centrifuge (Eppendorf Inc., Madison, WI), and resuspended in GTP-free buffer. Decoration of polymerized
microtubules with rK354 was performed in solution at a final tubulin concentration of 1 mg/ml (~10 µM) and a final kinesin concentration of 1.5 mg/ml (~30-40 µM, depending on construct) and in the presence of 2 mM
AMP-PNP. This stoichiometry was found previously (Hoenger and Milligan, 1997) to yield complete decoration. Samples were incubated for 2 min and subsequently adsorbed to holey carbon grids for 1 min, and then
quick-frozen in liquid propane by using a plunger essentially following the standard procedures described initially by Dubochet et al. (1985)
. Decoration of microtubules with rK379 was performed directly on the grid to
avoid bundling of tubules, either in the presence of 2 mM AMP-PNP, or in
the absence of nucleotides. To this end, microtubules at a concentration of
1 mg/ml were adsorbed on holey carbon grids for 1 min, washed briefly,
and then incubated with a rK379 solution at 3 mg/ml (to obtain the same
stoichiometry as for rK354 = approximately two kinesin double heads per
tubulin dimer) for 2 min and quick-frozen as described above.
Cryoelectron Microscopy
Cryoelectron microscopy was performed on a CM12 microscope (Philips
Electron Optics, Mahwah, NJ) equipped with an ultra-high vacuum chamber (MIDILAB; Gross et al., 1990) and a modified GATAN cryoholder
that allowed the grid to be transferred from the MIDILAB to the holder
without passing through air. The MIDILAB allowed storage of four frozen grids simultaneously at liquid N2 temperature in ultra-high vacuum,
which were readily available for a fast transfer into the microscope. In addition, we used the carbon evaporating capabilities of this chamber to coat
the frozen grids with a thin layer of carbon (30 Hz = ~3 nm) before observation to reduce charging effects on the grids. Micrographs were recorded at a magnification of 35,000× on SCIENTIA electron microscopy film
(AGFA, Leverkusen, Germany) (developed for 4 min in KODAK D19)
at an electron dose of <5 e
/Å2. Images were taken at a defocus range between
1 and
1.5 µm. On the CM12 twin lens microscope this defocus
range places the first zero node of the contrast transfer function between
1/27 and 1/20 Å.
Image Processing and Three-Dimensional Reconstruction
Because of the lattice properties of microtubules (Mandelkow et al., 1986;
Wade et al., 1990
; Sosa and Milligan, 1996
; Sosa et al., 1997b
) we were
searching for microtubules that exhibit a Moirè pattern (Mandelkow and
Mandelkow, 1985
; Wade et al., 1990
) corresponding to 15-protofilament/
2-start helical microtubules (Sosa et al., 1997b
). They accounted for ~5%
of the total number of microtubules present in a solution polymerized in
the presence of 5% DMSO (see above). 15-protofilament/2-start helical
microtubules also exhibit a characteristic diffraction patterns as shown in Fig. 2. Micrographs were digitized using a Leaf scanner (Leaf Systems,
Westborough, MA) operated at a pixel size of 20 µm on the negative,
which amounts to 0.57 nm on the sample. Suitable 15-protofilament/2-start microtubules were helically reconstructed by using the software
package PHOELIX (Whittaker et al., 1995a
). Some of them were long
enough to be prescreened for multiple seams by back projection (Wade et
al., 1995
; Sosa and Milligan, 1996
), but such cases were not observed. One
data set represents a near or far side of an intact microtubule; the microtubules had an average length of ~1.2 µm corresponding to ~150 tubulin
dimers per protofilament. A total of 19 data sets were used for the reconstruction of microtubules decorated with rK379 in the presence of AMP-PNP, 10 data sets went into the reconstructions of microtubules decorated with rK379 in the absence of nucleotide, 8 data sets were processed for microtubules decorated with rK354 (+AMP-PNP), and 14 data sets were
used to reconstruct the undecorated microtubule control map. Table I
summarizes the specific data for each reconstruction. All data sets were
truncated to a maximum resolution of 20 Å. Three-dimensional maps
were visualized by using the software packages SUPRIM (Schroeter and
Bretaudiere, 1996
), and SYNUVIEW (Hessler et al., 1996
). The three-dimensional maps of decorated microtubules were supplemented with the
x-ray structure of rK379 by using "O" (Jones et al., 1991
). All calculations
were performed on an INDIGO workstation (Impact-10000; Silicon
Graphics, Inc., Mountain View, CA). The polarity of the maps presented
here was determined according to Hoenger and Milligan (1996)
. All maps
are shown in a standard orientation (plus end up).
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Difference Mapping and t Tests
Difference maps were calculated to investigate the possibility of a residual signal from the second head (missing or dislocated) of rK379-decorated microtubules. All maps were brought to the same phase origin and subtracted from each other. We performed t tests to assess the statistical significance of the differences obtained. To this end we calculated variance maps from single and double head-decorated maps that contained 8 (rK354) and 19 (rK379) individual phase-aligned maps. For comparison, the maps from rK354-decorated microtubules were also tested against the undecorated microtubules (14 individual maps). This provided a calibration of the signal since the difference is due to a well-defined motor head domain. All calculations were carried out with the "varimap" and "t test" routines provided by PHOELIX.
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Results |
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Electron Microscopy Three-dimensional Reconstructions Reveal Only a Single Head of Kinesin per Tubulin Heterodimer, Both for Monomeric and "Dimeric" Kinesin
Kinesin constructs containing the motor domain but without or with only a short neck domain are monomeric in solution, while increasing lengths of the neck lead to dimerization because of their coiled-coil interactions (Huang et al.,
1994; Correia et al., 1995
; Jiang et al., 1997
). Thus, since
monomeric kinesin head domains bind to microtubules
with a stoichiometry of one head per tubulin subunit
(=
-
-heterodimer, mostly at
-tubulin; Song and Mandelkow, 1993
), one would expect dimeric constructs to
bind to two tubulin heterodimers, i.e., with the same stoichiometry of one kinesin head per tubulin heterodimer.
This is illustrated in Fig. 1 for the case of the dimeric construct rK379; results on other monomeric or dimeric constructs obtained with different methods are described elsewhere (Thormählen et al., 1998a
). In this regard, kinesin
differs from the retrograde motor ncd whose dimeric constructs bind with a stoichiometry of two heads per tubulin
subunit (Crevel et al., 1996
). Note however that the results on dimeric kinesin are in contrast to those derived from
two recent cryoelectron microscopy studies (Arnal et al.,
1996
; Hirose et al., 1996
), which were interpreted in terms
of two bound heads per tubulin subunit, similar to ncd (see
Discussion).
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Cryoelectron micrographs of microtubules decorated
with kinesin constructs rK354 (monomeric in solution)
and rK379 (dimeric) are shown in Fig. 2. The microtubules
chosen for processing had 15 protofilaments because this
allows helical image reconstructions without the complication of a seam asymmetry that is typical for the more common 13-protofilament microtubules (Sosa and Milligan, 1996; Sosa et al., 1997b
). The diffraction patterns of the
decorated particles reveal an additional set of strong layer
lines ~1/8 nm, corresponding to the axial spacing of the
decorating kinesin molecules (Fig. 2, B-D). This is equivalent to the spacing of tubulin heterodimers, consistent with
biochemical and other data showing that kinesin interacts
predominantly with
-tubulin (Song and Mandelkow,
1993
; Kikkawa et al., 1995
; Thormählen et al., 1998a
). The
reconstructed densities derived from the decorated microtubules (Fig. 3; averages of ~10,000-20,000 asymmetric
units = tubulin dimers) display two features. There is a
hollow cylinder (mean radius ~12.1 nm) consisting of 15 density peaks that represent the protofilaments seen in an
end-on projection (from plus to minus end), slewing in an
anticlockwise direction (see undecorated microtubule, Fig.
3 A; the left half is overlaid with the density contour lines).
After decoration with kinesin a further density peak appears attached to each protofilament, centered at a radius of ~16.6 nm and displaced in a clockwise direction (Fig. 3,
B-D). This density is very similar for both the monomeric
construct rK354 (Figs. 3 B and 4 A) and the "dimeric" construct rK379, and its shape is nearly independent of the nucleotide status of kinesin (AMP-PNP in Figs. 3, B and C,
and 4 B, or no nucleotide, Figs. 3 D and 4 C). Overall,
these maps are similar to those obtained with human kinesin hK349 (Hoenger and Milligan, 1997
; Sosa et al., 1997b
).
By contrast, microtubules decorated with a dimeric construct of the retrograde motor ncd showed two additional
densities attached to each protofilament, one centered at
an ~16.6-nm radius (as before), the second at ~19.9 nm,
displaced in an anticlockwise direction (Fig. 3 E; see Hoenger and Milligan, 1997
; Sosa et al., 1997a
).
A statistical analysis using the t test procedure was applied to obtain a comparison of motor related mass densities between microtubules decorated with rK354 and rK379 (Fig. 3, F-H). Both maps were reconstructed from motor head domains in the presence of AMP-PNP. As a control, we compared the map of rK354-decorated microtubules with the nondecorated microtubules to obtain a direct measure of the difference signal resulting from a well-defined motor head domain (Fig. 3 H). The comparison between maps from microtubules decorated with rK354 or rK379 revealed no significant mass attributable to a second head in the vicinity of the first head attached to tubulin (Fig. 3, F and G). We found slight differences in mass at the end of every attached head in the plus-end direction (Fig. 3, F and G, small arrow). This would be consistent with the 25-amino acid mass increase of rK379 over rK354 within the neck helix. A second faint difference peak is located on the outer surface of the attached motor (Fig. 3, F and G, long arrow). However, these differences are small, they disappear at the 98% confidence level and thus could not account for a second attached head.
The fact that both rK354 and rK379 show only one attached head per tubulin heterodimer is consistent with the
stoichiometry (Fig. 1). Nevertheless, it was still unexpected considering that the arrangement of heads in the
crystal structure is very different. This implies a major conformational change between the solute and bound states
of dimeric kinesin (Kozielski et al., 1997a; Marx et al.,
1998
; and see below). The volumes associated with the
motor mass are clearly equivalent to the volume of one
head only (Fig. 3, B-D). As shown later, the motor-related
volume can be filled completely with the x-ray-derived
map of one motor domain (Fig. 5). There may be subtle
differences in the attached kinesin heads between the no-nucleotide and AMP-PNP states, but at the resolution
shown here they are minute and shall therefore not be discussed in further detail. What matters is that the no-nucleotide state is the same as with AMP-PNP present: there is
only one head attached to a tubulin subunit, and the second one is not visible.
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The results described above become even more obvious
on the three-dimensional surface-rendered maps shown in
Fig. 4 where the volumes are adjusted approximately to
the molecular weights of the components. The volume of
the tubulin component is shown in blue. The monomeric
construct rK354 (Fig. 4 A, green), the dimeric construct
rK379 with AMP-PNP (Fig. 4 C, yellow) or without nucleotide (Fig. 4 C, red) have similar densities, equivalent to
one motor head per tubulin heterodimer, and there is no
evidence for additional motor mass in the case of microtubules decorated with "dimeric" rK379 (Fig. 4, B and C).
By contrast, a large additional volume equivalent to a second tethered head is found on microtubules decorated
with dimeric ncd (Fig. 4 D, white and red; Sosa et al., 1997a).
Orientation of the Bound Kinesin Head on Microtubules: Rear Surface Towards Tubulin, Neck Helix on Upper End and Tangential to Microtubule
The next step in the analysis was to interpret the low resolution kinesin density attached to microtubules in terms of
the high resolution x-ray model of kinesin. The structures
of the motor domains of human kinesin and Drosophila
ncd have been solved and shown to be closely related
(Kull et al., 1996; Sablin et al., 1996
). They consist of a core
sheet sandwiched between three
helices on either side;
the front face contains the nucleotide binding site, the rear
face contains the presumptive microtubule-binding region.
The two heads of dimeric ncd have been fitted into the image reconstruction (Sosa et al., 1997a
). More recently, the x-ray structures of the monomeric construct rK354 from
rat brain kinesin and the dimeric construct rK379 have
been solved (Kozielski et al., 1997a
; Sack et al., 1997
). The
core motor domains were found to be similar to those of
human kinesin and ncd, and in addition the structures revealed the positions of the NH2- and COOH-terminal regions, including the COOH-terminal neck helix,
7. The
coiled-coil interaction of this helix leads to the dimerization, as in the construct rK379. It would therefore be natural to assume that the crystal structures of kinesin monomers and dimers would fit into the reconstructed densities
observed for the same constructs bound to microtubules
(Fig. 5).
To orient the kinesin molecule on the microtubule surface, we first note that the reconstructed kinesin density
has an elongated shape, ~6 × 3.5 × 3 nm, and the long
axis runs parallel to the protofilaments (best seen in Fig. 5,
E and F). The x-ray model of kinesin and ncd have similar
dimensions (the "flower bouquet structure;" see Fig. 1 in
Kull et al., 1996) so the long axes of the image reconstruction and x-ray models must roughly coincide. The next criterium comes from sequence conservation and mutagenesis experiments. All conserved loops implied in microtubule
binding are on the rear face of the structure (e.g., L12, L8,
helices
4 and
5; Kull et al., 1996
), and an exchange of
critical residues on this surface affects both microtubule
binding and motility (Woehlke et al., 1997
). Thus the rear
surface of the kinesin head must be placed at the interface
between the motor and tubulin. This leaves two main
choices, the flower bouquet of the motor domain could be
either "up" or "down." The orientation up (as in Fig. 1 of
Kull et al., 1996
) would mean that the nucleotide-binding site is near the top end (towards the plus end of microtubules), the neck helix near the bottom; down is the opposite. The down orientation is strongly preferred by several
observations. First, the overall match between the x-ray
structure of the motor domain and the reconstructed density is better (note the pointed tip on the upper right of the
reconstructed density in Fig. 5 E, which marks the beginning of the neck helix). Second, the down orientation places loop L11 (not visible in the x-ray structure because
of disorder) near its position in the analogous motor ncd
(where L11 is ordered and visible), i.e., at the upper end of
the density, pointing clockwise. The up orientation is sterically forbidden because loop L11 would penetrate into the
microtubule surface. Third, Arnal et al. (1996)
and Hirose
et al. (1996)
presented difference maps of dimeric kinesin
constructs with monomeric ones, which both revealed the
location of additional mass towards the plus end of the microtubule, pointing anticlockwise. In our down orientation
this would coincide with the transition to the neck helix
(Fig. 5, E and F, upper right)
These considerations yield the orientation of the motor
domain shown in Fig. 5 (e.g., Fig. 5, A, C, and E, yellow; or
B, D, and F, blue). In front view (Fig. 5 E), the kinesin
head (yellow) appears suspended on the neck helix protruding at the upper end to the right. The orientation can
be visualized by the "left hand down" rule: because the strands run roughly vertical in this orientation, the view
can be compared with that of a left hand pointing down
and viewed from inside (thumb at upper right representing the neck helix). The microtubule-binding loops (on the
back of the hand) would lean against the protofilament in
the background (Fig. 5 E, red wire frame). Fig. 5 C shows a
side view of the kinesin structure (yellow) extending from
the left edge of a microtubule; the neck projects roughly
towards the observer helix (stretched out "thumb"). In the
top view (Fig. 5 A, yellow), the front of a microtubule is
seen from above (plus end), the neck helix of the bound
kinesin (yellow) projects to the right (anticlockwise), and
loop L11 to the left (clockwise). This view illustrates how the microtubule-binding surface of kinesin is embraced
between loop L11 and the neck helix on both sides, and
that the neck helix points tangentially to the microtubule
surface, which has consequences for models of motility
(see Discussion). In this picture, the "gearbox" residues
that are involved in the directionality (Case et al., 1997
)
and lie around strand
9 of kinesin, would correspond to
the upper end of the second finger, near the force-transmitting neck helix.
Whereas the core of the head fits well into the reconstructed density, most of the neck helix appears to lie outside the volume enclosed by the map (except the initial
segment). This can be explained by at least two factors: (a)
helices (single or coiled-coils) have very low contrast in
cryoelectron microscopy at ~2-nm resolution; and (b) the
necks could moreover be disordered or partly attached and merged with the density of tubulin so that they would
not be visible as distinct entities.
Gedankenspiel: How Could Dimeric Kinesin Be Oriented on the Microtubule Surface?
In the above discussion we have concentrated on one kinesin head bound to the microtubule surface for every tubulin heterodimer. The orientation is appropriate for the monomeric construct rK354. However, rK379 is dimeric in
solution and in the crystals, owing to its coiled-coil interaction (Kozielski et al., 1997a; Marx et al., 1998
), and one
might expect that it would bind to microtubules in a
dimeric form that could be related to the observed x-ray structure. The image reconstruction results show that this
is not the case. Nevertheless it is worthwhile to ask
whether and how the x-ray dimer structure might theoretically fit onto the microtubule lattice.
We can make the theoretical assumption that one of the
heads in a kinesin dimer (the "tightly" bound head) would
bind in the orientation compatible with the reconstructed
density and ask where the second ("loosely" bound) head
would come to lie if it had the same arrangement as in the
x-ray structure. In the crystals, the two heads are related
by a rotation of ~120 degrees around an axis close to that
of the coiled-coil neck; the two neck helices have a somewhat different conformation so that we can distinguish heads A and B within a dimer, whereas the two core motor domains are nearly superimposable. If we consider
head A as the firmly bound head (Fig. 5 B, D, and F, blue
head, right), the head B (yellow) would have to penetrate
into the microtubule surface, which is sterically impossible. Conversely, if head B were the tightly bound head
(Fig. 5, yellow head, left column), the tethered head A
(blue) would mostly stay clear of the microtubule surface, this would therefore be the only theoretically allowed
choice if one wanted to preserve the x-ray structure of the
dimer. Head A would lie above the tightly bound head B,
displaced in an anticlockwise direction. However, in practice this region shows no density in the reconstructions obtained with rK379. Note that this hypothetical position of
the loosely bound head is different from the observed second head of ncd bound to microtubules (Fig. 4 D; Sosa
et al., 1997a), emphasizing again the different behavior of
these two motors.
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Discussion |
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Orientation and Stoichiometry of Kinesin Heads Bound to Microtubules
The initial aim of this study was to explain the structure of
kinesin-decorated microtubules in terms of the x-ray structures of monomeric and dimeric kinesin (Kozielski et al.,
1997a; Sack et al., 1997
).We hoped that the arrangement
of dimeric kinesin heads on the microtubule surface would
shed light on the mechanism of kinesin's movement along
microtubules. This expectation was partly fulfilled, but in a
manner that had not been anticipated. At the same time,
our results explain the curious differences between previous image reconstructions of microtubules decorated with
plus or minus-end directed motors (Arnal et al., 1996
; Hirose et al., 1996
), but again in an unforeseen fashion.
Monomeric kinesin (rK354) binds with a stoichiometry
of one motor domain per tubulin heterodimer, and in an
orientation comparable to that of monomeric ncd (Sosa
et al., 1997a). It can be visualized by the left hand down
rule (see above). Since the NH2- and COOH-terminal residues are visible in our structure, we can describe the disposition of the neck helix relative to the microtubule and
the core motor domain. Viewing the microtubule from
outside and in standard orientation (plus end up), the motor domain lies mostly over
-tubulin, its long axis is
roughly vertical, and the neck helix (thumb) projects from
the upper end towards the right. This direction would be roughly tangential to the microtubule surface (Figs. 5 E
and 6 B; but note that it is conceivable that the neck helix
protrudes from the head in a different angle when attached to tubulin). The gearbox residues are at the upper
end ("second finger"), near the neck helix. The orientation
is roughly equivalent to the standard view of Sack et al.
(1997)
but rotated upside down. Viewed from the top (Fig.
5 A) the motor domain appears to wrap around the crest
of the protofilament, with loop L11 (disordered in kinesin but visible in ncd) pointing clockwise and the neck helix
pointing anticlockwise. This orientation of the neck is in
contrast to the radial orientation expected for a lever-type
mechanism of motility, sometimes postulated in analogy to
the conformational change observed with the actin-dependent motor myosin (for reviews see Rayment, 1996
;
Holmes, 1997
; Vale and Fletterick, 1997
).
|
The dimeric kinesin construct rK379 decorates microtubules essentially in the same way and in the same orientation as monomeric constructs. Judging from the noise level
at higher radii and between the heads, there is no evidence
for a second head tethered to the first (Fig. 3, F and G).
Since the construct is dimeric in solution and in the crystals (Kozielski et al., 1997b), it follows that the two heads
of the kinesin dimer have come apart upon interaction with the microtubule, for reasons discussed below. The resulting stoichiometry is one head per tubulin heterodimer,
consistent with other biochemical and structural data (Fig.
1; Song and Mandelkow, 1993
; Thormählen et al., 1998a
),
but formally in contrast to models derived from nucleotide
exchange experiments (e.g., Hackney, 1994
; Ma and Taylor, 1997
). Since the bound kinesin head is monomeric, there is no way to fit the entire crystal structure of the
dimer into the kinesin density observed by cryoelectron
microscopy (Fig. 5). This structure of kinesin "dimers" is
therefore clearly different from that of ncd dimers where
two heads are found attached on one tubulin subunit (Fig.
4 D; Sosa et al., 1997a
).
Implications for Models of Kinesin Motility
The result that the two heads of a dimeric kinesin construct bind to microtubules in a monomeric fashion has
several interesting consequences. The first is that the
strength of the tubulin-motor interaction partly overrides
the coiled-coil interaction within the kinesin dimer, forcing
the heads to come apart so that the individual kinesin molecules can bind to different tubulin heterodimers, relieving
the constraints imposed by the coiled-coil necks (Thormählen et al., 1998b).
The separation of heads during microtubule attachment
removes one of the major obstacles in modeling the motion of kinesin along microtubules: How could a kinesin
dimer, consisting of two small heads only ~6-nm long and
tied together directly at their base, ever achieve the observed step size of 8 nm? One theoretical possibility would
be that the entire dimeric complex would "hop" 8 nm at a
time, but this would mean that both heads would transiently dissociate from the microtubule simultaneously,
which would be difficult to reconcile with the observed
processivity (requiring at least one head to hold on to the
microtubule; Berliner et al., 1995; Gilbert et al., 1995
; Vale
et al., 1996
). Because of this constraint, most models assume that the two kinesin heads move sequentially in a
"hand-over-hand" fashion, either along one protofilament ("tightrope"), or along neighboring protofilaments ("waddling;" for review see Cross, 1995
). These models are consistent with important observations, such as kinetics of nucleotide exchange ("alternating head catalysis;" Hackney,
1994
; Ma and Taylor, 1997
; Gilbert et al., 1998
), or with
the movement of motors along protofilament tracks (Ray
et al., 1993
). However, to generate an 8-nm, center-of-mass step size, a single head would have to move 16 nm at
a time. This would in turn require the heads to have long
and semi-independent necks. Accordingly, the models for
kinesin movement are usually shown with long and free
necks, but this is clearly in contradiction to the observed
crystal structure of kinesin dimers (Kozielski et al., 1997a
).
The resolution of this dilemma is to separate the heads
specifically upon interaction with microtubules, as observed in this study.
The gain in freedom for the heads can be estimated as
follows. The length of the neck helix (Ala339-Trp370) is
~4.6 nm (0.15 nm per helical residue). This is followed by
a disordered region which extends to another predicted
coiled-coil starting around residue Val406. Assuming an
average chain length of 0.2-0.3 nm per residue, this would
amount to ~10-15 nm, giving a total of ~15-20 nm per
head. This would generate a "long leash," allowing the two
heads to be separated by twice this value (~30-40 nm) more than sufficient to cover the distance of 16 nm per
head. In addition, we note that the strands 9 and
10 connecting the motor domain to the neck helix and harboring
the gearbox residues (Fig. 6, white) are only weakly attached to the core motor and could possibly detach during
some stage of the movement, giving even more freedom of
movement (Romberg et al., 1998
).
Even if the two heads of a kinesin dimer are separated
on the microtubule surface one would expect that longer
constructs, or whole kinesin dimers, would stay in contact
further downstream, for example from the second coiled-coil region onwards (e.g., starting around residue 406). In
principle, it should therefore be possible to detect a pairing of kinesin stalk domains on the microtubule surface in
the case of longer constructs. For full-length kinesin, this is
technically difficult to show experimentally because the
long stalks prevent periodic binding of kinesin to microtubules (a prerequisite for image analysis). However, intermediate constructs (up to 500 residues in length) tend to
reveal an additional 16 nm layer line, twice the repeat of a
tubulin heterodimer (Thormählen et al., 1998a; and Fig. 6
A). This is interpreted as a pairing of kinesin monomers
into dimers in the axial direction, arguing for a tightrope
model where the two monomers of a dimer bind to successive tubulin subunits along one protofilament, as illustrated in the models of Fig. 6 (B, front view, and C, lateral
view of left edge). The 16-nm periodicity would arise from
the stalk domain common to both monomers of a dimer.
Comparison between Kinesin and ncd
Why do double-headed kinesin and ncd constructs look
and move differently on a microtubule, in spite of their
similar motor domain? This question has been addressed
in several recent studies based on image reconstruction
and biochemical data. There appears to be a consensus
that monomeric kinesin and ncd occupy the same binding
site on microtubules, both from binding studies experiments (Lockhart et al., 1995; Song and Endow, 1996
) and
image reconstruction (Hoenger and Milligan, 1997
). The
firmly bound head of ncd was compared with its x-ray
structure (Sablin et al., 1996
), resulting in the orientation
proposed by Sosa et al. (1997a)
. In the present work we
find a very similar orientation (left hand down when microtubule is plus end up) by comparing the image reconstruction of rat kinesin with its x-ray structure (Sack et al.,
1997
).
Differences on the level of observation and interpretation exist when one compares double-headed kinesin and
ncd. The dimeric construct ncd-295-700 clearly attaches
to microtubules as a dimer, one head firmly bound, the
second tethered to the first, as in Fig. 4 D (Arnal et al.,
1996
; Hirose et al., 1996
; Sosa et al., 1997a
). Both heads
had roughly the volume expected for their molecular weight. In the case of "dimeric" kinesin the "second head"
had only ~20% of its expected mass, which was interpreted in terms of disorder (Arnal et al., 1996
; Hirose et al.,
1996
; Amos and Hirose, 1997
). On the basis of our results
we would question that interpretation; the additional density could simply be accounted for by the additional mass
of the dimeric construct, compared with the monomeric
construct (401-340 = 61 residues or 18%). Thus, the reconstructions of Arnal et al. (1996)
and Hirose et al. (1996)
would be compatible with our interpretation of only one
head bound per tubulin dimer. In our case the difference
amounts to only 379-354 = 25 residues or 7%, which is not
enough to make a visible contribution at the limited resolution. Alternatively, partial decoration by the second
head may be more likely with the slightly longer constructs
used by Arnal et al. (1996)
and Hirose et al. (1996)
. This
partial decoration may reveal a reduced mass as found in
their reconstructions. Nevertheless this would still support our hypothesis of a (preferred) disassembly of the dimer
neck region once one of the motor binds to tubulin.
The differences in interpretation had consequences for
proposed mechanisms of motility. The second head of ncd
and the (interpreted) second head of kinesin had different
orientations on the microtubule lattice, and it was thought
that this was related to their different directionality. Our
results argue that this view should be abandoned. The determination of directionality appears to be more subtle than apparent from gross head-head interactions, involving short gearbox sequences that may mediate between
the core motor domain and the neck helix (Case et al.,
1997; Henningsen and Schliwa, 1997
; Romberg et al.,
1998
). Moreover, it is likely that the directionality is built
into individual motor molecules since there are monomeric kinesins (e.g., KIF1A, KIF1B; Nangaku et al., 1994
;
Okada et al., 1995
), and since even single chains of dimeric
motors can generate movement (Berliner et al., 1995
, Inoue et al., 1997
).
The question that remains to be explained is Why do
dimeric ncd constructs remain joined upon attaching to
microtubules, while kinesin constructs come apart? The
answer is likely to be the strength of the intra-dimer interaction, coupled possibly with conformational differences.
The intra-dimer interaction is thought to rest mainly on
the coiled-coil interaction in the neck and stalk region (although this might be modulated by interactions with the
motor domain and with tubulin). Since the NH2 and
COOH termini of the motor domain are spatially very
close to one another the transition to the neck helix would
also be expected to be similar (thus far the structure of the
transition region is only known for kinesin; see Sack et al.,
1997). Fig. 7 illustrates the predicted coiled-coil probability (using the COILS algorithm; Lupas et al., 1991
); the kinesin neck is COOH-terminal to the motor domain, the
ncd neck is NH2-terminal. The strength of the coiled-coil
interaction appears to be weaker in the initial neck (nearest the motor) than in segments further away (Morii et al., 1997
; Tripet et al., 1997
). This argues in favor of a mechanism whereby the two heads of a dimer separate during
some step in the motile cycle, even in the case of ncd. The
coiled-coil predictions alone do not clearly distinguish between ncd's tendency to stay dimeric on a tubulin subunit,
whereas kinesin heads are forced apart and settle on different tubulin subunits. However, the predictions provide
a testable hypothesis that can be addressed experimentally using mutant forms of kinesin or ncd.
|
A final point concerns the possibility of a lever arm
mechanism of motility. This mechanism is currently discussed for the case of the actin-dependent motor myosin
since, in this case, speed of movement depends on the
length of the neck helix (Anson et al., 1996; Uyeda et al.,
1996
), since the neck helix undergoes nucleotide-dependent angular motions (Whittaker et al., 1995b
), and since
the x-ray structure of myosin shows related changes in the
nucleotide-binding domain (Fisher et al., 1995
; Smith and Rayment, 1996
). The lever arm hypothesis presumes that
the neck points away from the actin filament, towards the
myosin filament, and generates movement by swinging
(for reviews see Rayment, 1996
; Holmes, 1997
). In the
case of kinesin (and probably ncd), the neck helix does not
point away from the microtubule surface, but rather tangentially to it (Fig. 6), and
unlike with myosin
the coiled-coil becomes separated upon binding to microtubules. It is therefore not likely that myosin-type models
can be applied to microtubule motors. Further structural
studies with variants of motor proteins should reveal
whether an angular change in the neck or stalk region is
involved in the movement.
![]() |
Footnotes |
---|
Received for publication 8 October 1997 and in revised form 2 February 1998.
S. Sack is a recipient of a fellowship from the Friedrich-Ebert Foundation. The project was supported by the Deutsche Forschungsgemeinschaft and the Swiss Science Foundation.We are grateful to P. Tittmann (ETH, Zurich, Switzerland) for help with specimen preparation; M. Whittaker (The Scripps Institute, La Jolla, CA) for help with the transfer of programs to Zürich; S. Brady (University of Texas, Dallas, TX) for providing the original clone of rat kinesin; R. Milligan (The Scripps Institute) and E. Mandelkow (Hamburg) for many discussions on structural and biochemical aspects.
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References |
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---|
1. | Amos, L.A., and K. Hirose. 1997. The structure of microtubule-motor complexes. Curr. Opin. Cell Biol. 9: 4-11 |
2. | Anson, M., M.A. Geeves, S.E. Kurzawa, and D.J. Manstein. 1996. Myosin motors with artificial lever arms. EMBO (Eur. Mol. Biol. Organ.) J. 15: 6069-6074 [Abstract]. |
3. | Arnal, I., F. Metoz, S. DeBonis, and R.H. Wade. 1996. Three-dimensional structure of functional motor proteins on microtubules. Curr. Biol. 6: 1265-1270 |
4. | Berliner, E., E.C. Young, K. Anderson, H.K. Mahtani, and J. Gelles. 1995. Failure of a single-headed kinesin to track parallel to microtubule protofilaments. Nature. 373: 718-721 |
5. | Brady, S.T.. 1995. A kinesin medley: biochemical and functional heterogeneity. Trends Cell Biol. 5: 159-164 . |
6. | Case, R.B., D.W. Pierce, N. Hom-Booher, C.L. Hart, and R.D. Vale. 1997. The directional preference of kinesin motors is specified by an element outside of the motor catalytic domain. Cell. 90: 959-966 |
7. | Cole, D., and J.M. Scholey. 1995. Structural variations among the kinesins. Trends Cell Biol. 5: 259-262 . |
8. |
Coppin, C.M.,
J.T. Finer,
J.A. Spudich, and
R.D. Vale.
1996.
Detection of sub-8-nm movements of kinesin by high-resolution optical-trap microscopy.
Proc. Natl. Acad. Sci. USA.
93:
1913-1917
|
9. | Correia, J.J., S.P. Gilbert, M.L. Moyer, and K.A. Johnson. 1995. Sedimentation studies on the kinesin motor domain constructs K401, K366, and K341. Biochemistry 34: 4898-4907 |
10. | Crevel, I.M., A. Lockhart, and R.A. Cross. 1996. Weak and strong states of kinesin and ncd. J. Mol. Biol. 257: 66-76 |
11. | Cross, R.A.. 1995. On the hand-over-hand footsteps of kinesin heads. J. Muscle Res. Cell Motil. 16: 91-94 |
12. | Dubochet, J., M. Adrian, J. Lepault, and A.W. McDowall. 1985. Cryo-electron microscopy of vitrified biological specimens. Trends Biochem. Sci. 10: 143-146 . |
13. | Fisher, A.J., C.A. Smith, J.B. Thoden, R. Smith, K. Sutoh, H.M. Holden, and I. Rayment. 1995. X-ray structures of the myosin motor domain of Dictyostelium discoideum complexed with MgADP.BeFx and MgADP.AlF4. Biochemistry. 34: 8960-8972 |
14. | Gilbert, S.P., M.R. Webb, M. Brune, and K.A. Johnson. 1995. Pathway of processive ATP hydrolysis by kinesin. Nature. 373: 671-676 |
15. | Gilbert, S.P., M.L. Moyer, and K.A. Johnson. 1998. Alternating site mechanism of the kinesin ATPase. Biochemistry 37: 792-799 |
16. | Goldstein, L.S.B.. 1993. With apologies to Scheherazade: Tails of 1001 kinesin motors. Annu. Rev. Genet. 27: 319-351 |
17. | Gross, H., K. Krusche, and P. Tittmann. 1990. Recent progress in high resolution shadowing for biological TEM. XIIth Intl. Cong. Electron Microsc. Seattle, WA. 510-511. |
18. | Hackney, D.D.. 1994. Evidence for alternating head catalysis by kinesin during microtubule-stimulated ATP hydrolysis. Proc. Natl. Acad. Sci. USA. 91: 6865-6869 [Abstract]. |
19. | Harrison, B.C., S.P. Marchese-Ragona, S.P. Gilbert, N. Cheng, A.C. Steven, and K.A. Johnson. 1993. Decoration of the microtubule surface by one kinesin head per tubulin heterodimer. Nature. 362: 73-75 |
20. | Henningsen, U., and M. Schliwa. 1997. Reversal of the direction of movement of a molecular motor. Nature. 389: 93-96 |
21. | Hessler, D., S.J. Young, and M.H. Ellisman. 1996. A flexible environment for the visualization of three-dimensional biological structures. J. Struct. Biol. 116: 113-119 |
22. | Hirose, K., A. Lockhart, R.A. Cross, and L.A. Amos. 1995. Nucleotide dependent angular change in kinesin motor domain bound to tubulin. Nature. 376: 277-279 |
23. |
Hirose, K.,
A. Lockhart,
R.A. Cross, and
L.A. Amos.
1996.
Three-dimensional
cryoelectron microscopy of dimeric kinesin and ncd motor domains on microtubules.
Proc. Natl. Acad. Sci. USA.
93:
9539-9544
|
24. | Hoenger, A., and R.A. Milligan. 1996. Polarity of 2-D and 3-D maps of tubulin sheets and motor decorated sheets. J. Mol. Biol. 263: 114-119 |
25. | Hoenger, A., and R.A. Milligan. 1997. Motor domains of kinesin and ncd interact with microtubule protofilaments with the same binding geometry. J. Mol. Biol. 265: 553-564 |
26. | Hoenger, A., E.P. Sablin, R.D. Vale, R.J. Fletterick, and R.A. Milligan. 1995. Three-dimensional structure of a tubulin-motor-protein complex. Nature. 376: 271-274 |
27. | Holmes, K.C.. 1997. The swinging lever-arm hypothesis of muscle contraction. Curr. Biol 7: 112-118 |
28. | Howard, J.. 1996. The movement of kinesin along microtubules. Annu. Rev. Physiol. 58: 703-729 |
29. | Hua, W., E.C. Young, M.L. Fleming, and J. Gelles. 1997. Coupling of kinesin steps to ATP hydrolysis. Nature. 388: 390-393 |
30. |
Huang, T.G.,
J. Suhan, and
D.D. Hackney.
1994.
Drosophila kinesin motor domain extending to amino acid position 392 is dimeric when expressed in Escherichia coli.
J. Biol. Chem.
269:
16502-16507
|
31. |
Inoue, Y.,
Y.Y. Toyoshima,
A.H. Iwane,
S. Morimoto,
H. Higuchi, and
T. Yanagida.
1997.
Movements of truncated kinesin fragments with a short or
an artificial flexible neck.
Proc. Natl. Acad. Sci. USA.
94:
7275-7280
|
32. |
Jiang, W.,
M.F. Stock,
X. Li, and
D.D. Hackney.
1997.
Influence of the kinesin
neck domain on dimerization and ATPase kinetics.
J. Biol. Chem.
272:
7626-7632
|
33. | Jones, T.A., J.Y. Zou, S.W. Cowan, and M. Kjeldgaard. 1991. Improved methods for building protein models in electron density maps and the location of errors in these models. Acta Crystallogr. A. 47: 110-119 |
34. | Kikkawa, M., T. Ishikawa, T. Wakabayashi, and N. Hirokawa. 1995. Three- dimensional structure of the kinesin head-microtubule complex. Nature. 376: 274-277 |
35. | Kozielski, F., S. Sack, A. Marx, M. Thormählen, E. Schönbrunn, V. Biou, A. Thompson, E.-M. Mandelkow, and E. Mandelkow. 1997a. The crystal structure of dimeric kinesin and implications for microtubule-dependent motility. Cell. 91: 985-994 |
36. | Kozielski, F., E. Schönbrunn, S. Sack, J. Müller, S.T. Brady, and E. Mandelkow. 1997b. Crystallization and preliminary X-ray analysis of the single-headed and double-headed motor protein kinesin. J. Struct. Biol. 119: 28-34 |
37. | Kull, F.J., E. Sablin, P. Lau, R. Fletterick, and R. Vale. 1996. Crystal structure of the kinesin motor domain reveals a structural similarity to myosin. Nature. 380: 550-554 |
38. | Lockhart, A., I.M. Crevel, and R.A. Cross. 1995. Kinesin and ncd bind through a single head to microtubules and compete for a shared MT binding site. J. Mol. Biol. 249: 763-771 |
39. | Lupas, A., M. Van Dyke, and J. Stock. 1991. Predicting coiled coils from protein sequences. Science. 252: 1162-1164 |
40. |
Ma, Y.Z., and
E.W. Taylor.
1997.
Interacting head mechanism of microtubule-kinesin ATPase.
J. Biol. Chem.
272:
724-30
|
41. | Mandelkow, E.-M., and E. Mandelkow. 1985. Unstained microtubules studied by cryo-electron microscopy: substructure, supertwist and disassembly. J. Mol. Biol. 181: 123-135 |
42. | Mandelkow, E.-M., M. Herrmann, and U. Rühl. 1985. Tubulin domains probed by limited proteolysis and subunit-specific antibodies. J. Mol. Biol. 185: 311-327 |
43. | Mandelkow, E.-M., R. Schultheiss, R. Rapp, M. Müller, and E. Mandelkow. 1986. On the surface lattice of microtubules: Helix starts, protofilament number, seam and handedness. J. Cell Biol. 102: 1067-1073 [Abstract]. |
44. | Marx, A., M. Thormählen, S. Sack, and E. Mandelkow. 1998. Crystal structure of kinesin and interaction with microtubules. Eur. Biophysics J. In press. |
45. | Morii, H., T. Takenawa, F. Arisaka, and T. Shimizu. 1997. Identification of kinesin neck region as a stable alpha-helical coiled coil and its thermodynamic characterization. Biochemistry. 36: 1933-1942 |
46. | Nangaku, M., R. Sato-Yoshitake, Y. Okada, Y. Noda, R. Takemura, H. Yamazaki, and N. Hirokawa. 1994. KIF1B, a novel microtubule plus end-directed monomeric motor protein for transport of mitochondria. Cell 79: 1209-1220 |
47. | Okada, Y., H. Yamazaki, Y. Sekine-Aizawa, and N. Hirokawa. 1995. The neuron-specific kinesin superfamily protein KIF1A is a unique monomeric motor for anterograde axonal transport of synaptic vesicle precursors. Cell. 81: 769-780 |
48. | Ray, S., E. Meyhofer, R.A. Milligan, and J. Howard. 1993. Kinesin follows the microtubule's protofilament axis. J. Cell Biol. 121: 1083-1093 [Abstract]. |
49. | Rayment, I.. 1996. Kinesin and myosin: molecular motors with similar engines. Structure. 4: 501-504 |
50. |
Romberg, L.,
D.W. Pierce, and
R.D. Vale.
1998.
Role of the kinesin neck region in processive microtubule-based motility.
J. Cell Biol.
140:
1407-1416
|
51. | Sablin, E.P., F.J. Kull, R. Cooke, R.D. Vale, and R.J. Fletterick. 1996. Crystal structure of the motor domain of the kinesin-related motor ncd. Nature. 380: 555-559 |
52. | Sack, S., J. Müller, A. Marx, M. Thormählen, E.-M. Mandelkow, S.T. Brady, and E. Mandelkow. 1997. X-ray structure of motor and neck domains from rat brain kinesin. Biochemistry. 36: 16155-16165 |
53. | Schnitzer, M.J., and S.M. Block. 1997. Kinesin hydrolyses one ATP per 8-nm step. Nature. 388: 386-90 |
54. | Schroeter, J., and J.-P. Bretaudiere. 1996. SUPRIM: easily modified image processing software. J. Struct. Biol. 116: 131-137 |
55. | Smith, C.A., and I. Rayment. 1996. X-ray structure of the magnesium(II).ADP.vanadate complex of the Dictyostelium discoideum myosin motor domain to 1.9 Å resolution. Biochemistry. 35: 5404-5417 |
56. | Song, H., and S.A. Endow. 1996. Binding sites on microtubules of kinesin motors of the same or opposite polarity. Biochemistry. 35: 11203-11209 |
57. |
Song, Y.H., and
E. Mandelkow.
1993.
Recombinant kinesin motor domain
binds to ![]() |
58. | Sosa, H., and R.A. Milligan. 1996. Three-dimensional structure of ncd-decorated microtubules obtained by a back-projection method. J. Mol. Biol. 260: 743-755 |
59. | Sosa, H., D.P. Dias, A. Hoenger, M. Whittaker, E. Wilson-Kubalek, E. Sablin, R.J. Fletterick, R.D. Vale, and R.A. Milligan. 1997a. A model for the microtubule-Ncd motor protein complex obtained by cryo-electron microscopy and image analysis. Cell. 90: 217-224 |
60. | Sosa, H., A. Hoenger, and R.A. Milligan. 1997b. Three different approaches for calculating the 3-dimensional structure of microtubules decorated with kinesin motor domains. J. Struct. Biol. 118: 149-158 |
61. | Svoboda, K., C.F. Schmidt, B.J. Schnapp, and S.M. Block. 1993. Direct observation of kinesin stepping by optical trapping interferometry. Nature. 365: 721-727 |
62. | Thormählen, M., A. Marx, S.A. Müller, Y.-H. Song, E.-M. Mandelkow, U. Aebi, and E. Mandelkow. 1998a. Interaction of monomeric and dimeric kinesin with microtubules. J. Mol. Biol. 275: 795-809 |
63. | Thormählen, M., A. Marx, S. Sack, and E. Mandelkow. 1998b. The coiled-coil helix in the neck of kinesin. J. Struct. Biol. In press. |
64. |
Tripet, B.,
R.D. Vale, and
R.S. Hodges.
1997.
Demonstration of coiled-coil interactions within the kinesin neck region using synthetic peptides![]() |
65. |
Tucker, C., and
L.S.B. Goldstein.
1997.
Probing the kinesin-microtubule interaction.
J. Biol. Chem.
272:
9481-9488
|
66. |
Uyeda, T.Q.,
P.D. Abramson, and
J.A. Spudich.
1996.
The neck region of the
myosin motor domain acts as a lever arm to generate movement.
Proc. Natl.
Acad. Sci. USA.
93:
4459-4464
|
67. | Vale, R.D., and R.J. Fletterick. 1997. The design plan of kinesin motors. Annu. Rev. Cell Dev. Biol. 13:745-777. |
68. | Vale, R.D., T. Funatsu, D.W. Pierce, L. Romberg, Y. Harada, and T. Yanagida. 1996. Direct observation of single kinesin molecules moving along microtubules. Nature. 380: 451-453 |
69. | Wade, R.H., D. Chrètien, and D. Job. 1990. Characterization of microtubule protofilament numbers; how does the surface lattice accommodate? J. Mol. Biol. 212: 775-786 |
70. | Wade, R.H., R. Horowitz, and R.A. Milligan. 1995. Toward understanding the structure and interactions of microtubules and motor proteins. PROTEINS: structure, function, and genetics 23:502-509. |
71. |
Walker, R.A..
1995.
Ncd and kinesin motor domains interact with both alpha-
and beta-tubulin.
Proc. Natl. Acad. Sci. USA.
92:
5960-5964
|
72. | Whittaker, M., B.O. Carragher, and R.A. Milligan. 1995a. PHOELIX: a package for semi-automated helical reconstruction. Ultramicroscopy. 58: 245-259 |
73. | Whittaker, M., E.M. Wilson-Kubalek, J.E. Smith, L. Faust, R.A. Milligan, and H.L. Sweeney. 1995b. A 35Å movement of smooth muscle myosin on ADP release. Nature. 378: 748-751 |
74. | Woehlke, G., A.K. Ruby, C.L. Hart, B. Ly, N. Hom-Booher, and R.D. Vale. 1997. Microtubule interaction site of the kinesin motor. Cell. 90: 207-216 |