Wellcome Trust/Cancer Research UK Institute and Department of Genetics, University of Cambridge, Tennis Court Road, Cambridge CB2 1QR, UK
* Author for correspondence (e-mail: ds139{at}mole.bio.cam.ac.uk)
Accepted 5 September 2002
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SUMMARY |
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Key words: Motor proteins, Axis formation, Cytoplasmic streaming, RNA transport, Kinesin, Drosophila
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INTRODUCTION |
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Conventional Kinesin, Kinesin I, was the first member of the Kinesin
superfamily to be identified, and is responsible for the ATP-dependent
transport of several distinct cargoes along microtubules, such as vesicles,
membranous organelles and pigment granules
(Brady, 1985;
Goldstein and Yang, 2000
;
Vale et al., 1985
;
Yang et al., 1989
). Kinesin I
is composed of two Kinesin heavy chains (KHC) and two Kinesin light chains
(KLC), each of which is encoded by several genes in vertebrates, but
single-copy genes in Drosophila. The KHC has an N-terminal motor
domain that contains both the microtubule and ATP binding sites, a central
coiled-coil domain that dimerises to form a bipartite stalk, and a globular
C-terminal tail. The KLC binds to the C-terminal region of the heavy chain
stalk through its N-terminal coiled-coil domain, while its C-terminal region
contains six tetra-trico peptide (TPR) domains, which interact with cargo
adaptors (Bowman et al., 2000
;
Gauger and Goldstein, 1993
;
Gindhart and Goldstein, 1996
;
Kamal et al., 2000
;
Verhey et al., 2001
).
In addition to the heavy chain of Kinesin I, mutants in staufen,
barentsz, mago nashi, tsunagi (Drosophila Y14) and
non-muscular cytoplasmic tropomyosin II (TmII; Tm1
FlyBase) block the localisation of oskar mRNA to the posterior pole
(Erdélyi et al., 1995;
Hachet and Ephrussi, 2001
;
Micklem et al., 1997
;
Mohr et al., 2001
;
Newmark and Boswell, 1994
;
Tetzlaff et al., 1996
;
van Eeden et al., 2001
).
Staufen protein colocalises with oskar mRNA throughout oogenesis in
both wild-type and mutant egg chambers, and its posterior localisation is
oskar mRNA-dependent and vice versa
(Ferrandon et al., 1994
;
St Johnston et al., 1991
). The
protein contains five dsRNA binding domains, and this RNA-binding activity is
essential for oskar mRNA localisation
(Ramos et al., 2000
;
St Johnston et al., 1992
).
Thus, Staufen presumably binds directly to oskar mRNA and is a
reliable marker of the localisation of the transcript. Barentsz, Mago nashi
and Tsunagi/Y14 are also likely to be components of the oskar mRNA
localisation complex, as they colocalise with the mRNA to the posterior pole
(Hachet and Ephrussi, 2001
;
Mohr et al., 2001
;
Newmark et al., 1997
;
van Eeden et al., 2001
).
Kinesin I has not been shown to interact with any of these proteins, however,
and it is unclear whether the motor interacts with the oskar mRNA
complex directly.
Glotzer et al. have proposed an alternative model for oskar mRNA
localisation, in which cytoplasmic flows circulate the mRNA around the oocyte,
and it is then trapped at the posterior by a pre-localised anchor
(Glotzer et al., 1997). In
this model, the microtubules and kinesin would be required to generate the
cytoplasmic flows that facilitate the diffusion of the mRNA towards this
anchor. The bulk movement of the cytoplasm in the egg chamber has been
previously studied by video-enhanced contrast microscopy, and by time-lapse
films using confocal imaging of fluorescent yolk granules
(Gutzeit, 1986a
;
Gutzeit and Koppa, 1982
;
Theurkauf et al., 1992
;
Bohrmann and Biber, 1994
). From
stage 7 onwards, nurse cell cytoplasm flows through the ring canals into the
oocyte, where it is efficiently mixed with the ooplasm by cytoplasmic
movements in the oocyte, called streaming. The ooplasmic streaming at stage 9
consists of random motions in several directions, but the nature of these
movements changes dramatically at stage 10b when streaming becomes
unidirectional, and around five times faster
(Bohrmann and Biber, 1994
).
Actin-depolymerising drugs block the nurse cell-to-oocyte movements, but not
streaming within the oocyte. By contrast, microtubule-disrupting drugs have no
effect on the nurse cell-to-oocyte flow, but block both the slow streaming in
the oocyte at stage 9 and the more rapid streaming at 10b
(Gutzeit, 1986b
;
Gutzeit, 1986a
;
Bohrmann and Biber, 1994
). The
microtubules rearrange to form parallel arrays 5-10 µm below the oocyte
cortex when the stage 10b streaming begins, and vesicles are observed in close
proximity to these microtubules, suggesting that the movement is driven by
vesicle transport along microtubules
(Theurkauf et al., 1992
).
We have analysed the role of Kinesin I in the oocyte and showed that it is required for all cytoplasmic movements and posterior localisations in the oocyte. Surprisingly, the Kinesin light chain is not required for any of the functions of kinesin in the oocyte, indicating that it associates with its cargo by a novel mechanism.
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MATERIALS AND METHODS |
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Whole-mount in situ hybridisation and antibody staining
Females were fattened for 24 hours, and the ovaries dissected in PBT
(PBS+0.1%Tween), fixed for 20 minutes in 4% paraformaldehyde/PBT, washed with
PBT and kept in methanol at -20°C.
For antibody staining, the ovaries were then washed with PBT, blocked with
PBT-10 (PBT+10% BSA) for 1 hour and incubated with the antibody in PBT-1 (+1%
BSA) for 12 hours. After washing the ovaries with PBT-1 several times for 30
minutes, they were incubated with the secondary antibody for at least 3 hours.
They were finally washed three times with PBT for 15 minutes and mounted in
Vectashield (Vector). All steps were performed at room temperature. The
following antibodies were used: rabbit anti-Staufen (1:1000)
(St Johnston et al., 1991),
monoclonal P1H4 anti-Dhc64C (1:200)
(McGrail and Hays, 1997
) and
rabbit anti-KHC (1:250, Cytoskeleton company).
For RNA in situ hybridisation, the ovaries were rehydrated after the methanol treatment with PBT, post-fixed for 5 minutes with 4% paraformaldehyde/PBT, and washed three times for 10 minutes with PBT. They were then incubated in the pre-hybridisation buffer (50% formamide, 5xSSC, 0.1% Tween-20 pH 4.5) for 1 hour at 70°C, and then overnight at 70°C with the oskar mRNA antisense digoxigenin (DIG)-UTP probe in hybridisation buffer (pre-hybridisation + Boerhinger Calf tRNA 0.1 mg/ml + heparin 0.05 mg/ml). After several washes at 70°C with a 1:1 mix of pre-hybridisation:PBT, and at room temperature with PBT, the ovaries were incubated for 1 hour with an anti-DIG antibody coupled to either Alkaline Phosphatase (Roche) or to Cy3 (Jackson ImmunoResearch). In the first case, the ovaries were then washed with PBT three times for 20 minutes and once with staining solution (100 mM NaCl, 50 mM MgCl2, 100 mM Tris pH 9.5, 0.01% Tween-20). The reaction was developed by the addition of NBT:BCIP kit from Promega following the manufacturer's instructions. Finally, they were washed several times with PBS and mounted in 50% glycerol. In the second case, the ovaries were washed and mounted as previously described for the antibody staining.
Analysis of cytoplasmic movements
Females were fattened for 24 hours and dissected in 10S Voltalef oil
(Altachem). All time-lapse movies of living egg chambers were taken using
either a Nipkow spinning disk confocal or a BioRad confocal MRC1024 and a
Nikon inverted microscope. On the Nipkow spinning disk confocal we made 15
minutes time lapse films by collecting images every 2 seconds. On the BioRad
confocal, the time-lapse movies were obtained by collecting z series
of five sections at 2 µm intervals every minute for 45 minutes. Three
distinct type of particles were analysed: GFP-Staufen particles,
uncharacterised vesicles that were visualised by exciting the sample with the
568 nm wavelength light, and collecting the emission through a OG515 or OPEN
filter; and the yolk granules. The tricolor images shown in
Fig. 2A-J were obtained by
creating a Photoshop RGB file in which three consecutive images of a time
lapse movie were inserted in the red, green and blue channel, respectively.
The Kalman images shown in Fig.
2K-P, Fig. 3 and
Fig. 6 were obtained by using
the Kalman averaging function of the confocal microscope to merge successive
scans in one composite image. The oocytes were laser scanned either once (1)
or in a continuous manner for 15 (15) or 30 (30) times, and the composite
image is shown.
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RESULTS |
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The Kinesin heavy chain is required for cytoplasmic streaming in the
oocyte
One way to distinguish between the models for oskar mRNA
localisation is to observe how the mRNA moves to the posterior in living
oocytes. A GFP-Staufen fusion protein rescues the oskar mRNA
localisation defect of a staufen null mutant and localises to the
posterior (Fig. 2A)
(Schuldt et al., 1998), and we
attempted to use this fusion to visualise the movement of
Staufen/oskar mRNA complexes in vivo. To detect particles that move
at similar speeds to kinesin-dependent fast axonal transport, we made 15
minute time-lapse films by collecting images every 2 seconds on a Nipkow
spinning disc confocal microscope (see Movie 1 at
http://www.welc.cam.ac.uk/~dstjlab/isa_movies/isa_mov_index.htm).
Although many GFP-Staufen particles undergo rapid movements in the nurse
cells, we could not detect any fast-moving particles in the oocyte, and
observed that most particles move at much slower rates that are hard to see at
this time scale. We therefore generated slower time lapse films by imaging
once a minute for 45 minutes on a standard confocal microscope, and observed
bright GFP-Staufen particles that move from the anterior towards the posterior
of the oocyte (see Movie 2 at
http://www.welc.cam.ac.uk/~dstjlab/isa_movies/isa_mov_index.htm).
These movements require functional kinesin, as most particles remain
stationary in Khc27 mutant germline clones (see Movie 3 at
http://www.welc.cam.ac.uk/~dstjlab/isa_movies/isa_mov_index.htm).
This can be easily visualised by importing three successive images from the
time lapse films into the red, green and blue channels of a Photoshop file
(Fig. 2A-D)
(Theurkauf and Hazelrigg,
1998
). If a GFP-Staufen particle moves between scans, it appears
as separate red, green and blue dots, and the distance between them can then
be measured to calculate its speed. However, stationary particles appear
white, due to the superposition of the red, green and blue signals. Although
almost all particles in wild-type oocytes are motile, the majority of
particles in Khc27 oocytes are stationary, except for
those in the vicinity of the ring canals
(Fig. 2A-D, Q, R). This defect
on the movement of GFP-Staufen particles was completely rescued by a wild-type
Khc transgene. Kinesin is not required, however, for the motion of
these particles in the nurse cells, or for their movement from the nurse cells
into the oocyte. Quantification of these data reveals that the particles show
similar movements in the nurse cells of wild-type and mutant egg chambers
(average velocities of 5.0 µm per minute), but are much less motile in
mutant oocytes (5 µm compared with 0.5 µm per minute). Indeed, the
majority of particles that do move in mutant oocytes lie in the vicinity of
the ring canals, and may be carried by the flow of cytoplasm from the nurse
cells.
Unfortunately, the GFP-Staufen particles are unlikely to represent
Staufen/oskar mRNA complexes in transit to the posterior, because
untagged Staufen is not found in large particles unless it is overexpressed,
and the GFP-Staufen is expressed at much higher levels than the endogenous
protein, which is already present in excess over oskar mRNA
(Ferrandon et al., 1994).
Furthermore, the GFP-Staufen particles do not accumulate at the anterior of
the oocyte in mago nashi and barentsz mutants, in contrast
to oskar mRNA and endogenous Staufen (I. M. P. and D. St J.,
unpublished). Thus, they most probably correspond to aggregates of
overexpressed protein. Although a proportion of the GFP-Staufen must associate
with oskar mRNA to mediate its posterior localisation, and
accumulates with it at the posterior pole, we have been unable to determine
how it gets there, presumably because it localises in particles that are
either too small, too dim or too rare to image in this way.
Although the particles probably do not reflect the localisation of oskar mRNA, they do undergo dramatic kinesin-dependent movements in the oocyte, which could be caused either by active transport or by cytoplasmic flows. To distinguish between these possibilities, we compared their movements with those of the surrounding cytoplasm, which can be follow by visualising a particle in the oocyte that reflects 568 nm light, and which may correspond to a peroxisome. In wild-type oocytes, the red particles always move in the same direction and at the same speed as the GFP-Staufen particles in their vicinity, even though their relative positions rule out the possibility that they are attached to the same motor or microtubule (see Movie 4 at http://www.welc.cam.ac.uk/~dstjlab/isa_movies/isa_mov_index.htm). These observations, and the fact that the particles move much slower than in other kinesin-dependent transport processes indicate that these movements correspond to cytoplasmic flows. As is the case for GFP-Staufen, the removal of kinesin blocks virtually all movement of the red particles in the oocyte, but has no effect on their motion within the nurse cells or on nurse cell to oocyte transport (Fig. 2E-J; see Movie 5 at http://www.welc.cam.ac.uk/~dstjlab/isa_movies/isa_mov_index.htm). The yolk vesicles also fail to move in mutant oocytes, although the uptake of the yolk from the follicle cells is unaffected, and the oocyte grows at the normal rate (I. M. P. and D. St J., unpublished).
As an alternative way to visualise the cytoplasmic movements in the oocyte, we used the Kalman averaging function of the confocal microscope to merge successive scans into a composite image. Although Kalman averaging is normally used to reduce noise in images of stationary objects, moving particles become increasingly blurred as the number of scans is increased, and the direction of blurring indicates the orientation of their motion. Using this technique to image the red fluorescent particles in wild-type egg chambers reveals multiple flows in different directions, which vary from oocyte to oocyte (Fig. 2K-P). The fastest flows are always seen at the anterior of the oocyte, while the region around the posterior pole remains fairly quiescent (Fig. 2K-M). By contrast, no flows can be detected in Khc mutant oocytes, and the particles gradually become fuzzy because of Brownian motion as the number of scans increases (Fig. 2N-P).
These results show that the Kinesin heavy chain is required for all detectable cytoplasmic movements within the oocyte. This raises the question of the relationship between these flows and oskar mRNA localisation, and we therefore examined the movements of the red fluorescent particles in other mutants that block the posterior localisation of oskar mRNA. The ooplasmic movements are indistinguishable from those in wild type in staufen, mago nashi, barentsz and TmII mutant egg chambers (Fig. 2G,H; see Movie 6 at http://www.welc.cam.ac.uk/~dstjlab/isa_movies/isa_mov_index.htm). These results show that the kinesin-dependent ooplasmic flows are caused by the movement of some other structure than the oskar mRNA localisation complex.
When the nurse cells transfer most of their contents into the oocyte at stage 10b, the oocyte cytoplasm starts to flow unidirectionally around the oocyte at a speed that is about five times faster than at stage 9 (Fig. 3A-C). Although rapid streaming at stage 10b is distinct from the stage 9 flows, it also depends on the Kinesin heavy chain, as it does not occur in stage10b Khc mutant oocytes (Fig. 3D-F). This streaming defect is completely rescued by a wild-type Khc transgene. This effect is apparent in an even single time section. Normally, the material that enters the anterior of the oocyte from the nurse cells is efficiently dispersed by streaming throughout the ooplasm. In Khc mutants, however, this material does not mix with the rest of the cytoplasm, and the oocyte constituents become stratified. For example, fluorescent particles that are transported from the nurse cells into the oocyte accumulate at the anterior, while yolk spheres, which are endocytosed from the follicle cells, are excluded from the anterior and remain cortical. Thus, Kinesin is required for both the slow chaotic cytoplasmic flows at stage 9, and the faster directional flows at stage 10b, suggesting that a common mechanism generates both movements.
The Kinesin heavy chain localises to the posterior pole of the
oocyte
To further understand the function of Kinesin I in oskar mRNA
localisation and cytoplasmic streaming within the oocyte, we examined the
distribution of the KHC itself, and found that it is strongly localised at the
posterior of the oocyte from stage 9-10a. This staining is specific, as it is
not observed in Khc mutant germline clones
(Fig. 4A,C). The microtubule
cytoskeleton in the stage 9 oocyte is polarised with the plusends towards the
posterior, and this is therefore where one would expect an active plus-end
directed motor, like Kinesin I, to accumulate. The activity of Kinesin I is
believed to require cargo binding (Coy et
al., 1999; Friedman and Vale,
1999
). This localisation therefore strongly suggests that the
motor protein transports something along microtubules to the posterior pole.
However, the KHC still accumulates at the posterior both in staufen
mutant oocytes, in which oskar mRNA is not localised
(Fig. 4B). Thus, Kinesin I
presumably transports something else to the posterior pole, in addition to
oskar mRNA.
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The Kinesin heavy chain is required for Dynein heavy chain
localisation to the posterior pole of the oocyte
Another candidate cargo for Kinesin I is the Dynein heavy chain (Dhc64C,
DHC), which also localises to the posterior of the oocyte during stage 9 of
oogenesis (Li et al., 1994)
(Fig. 4D). DHC localisation is
not required for the posterior localisation of Staufen and oskar
mRNA, as they both localise normally in a combination of hypomorphic
Dhc64C alleles that abolishes the posterior localisation of DHC
(McGrail and Hays, 1997
).
Furthermore, DHC localisation is independent of the posterior localisation of
oskar mRNA, because it shows a wild-type accumulation at the
posterior in staufen, barentsz and mago nashi mutant egg
chambers (Fig. 4E; I. M. P. and
D. St J., unpublished). By contrast, DHC shows no posterior enrichment in
Khc27 germline clones
(Fig. 4F). This localisation
defect is completely rescued by a wild type Khc transgene. Thus, the
KHC is required for the posterior localisation of both DHC and oskar
mRNA, even though neither is required for the localisation of the other. The
KHC still accumulates at the posterior in Dhc64C mutant oocytes (I.
M. P. and D St J., unpublished). Thus, Kinesin I presumably transports
something else to the posterior pole, in addition to oskar mRNA and
DHC.
The functions of kinesin and dynein appear to be interdependent in
neurones, because mutants or inhibitors of either motor block both anterograde
and retrograde fast axonal transport (Brady
et al., 1990; Martin et al.,
1999
; Stenoien and Brady,
1997
; Waterman-Storer et al.,
1997
). To test whether this is also the case in the oocyte, we
examined whether the hypomorphic Dhc64C mutant combination has any
effect on streaming. The cytoplasmic flows still occur in this mutant, but
they are significantly slower than normal. Although this is consistent with
the idea that dynein and kinesin are interdependent, it is not possible to
test whether the cytoplasmic flows would be completely abolished in the
absence of dynein, because the null mutants block oocyte determination
(McGrail and Hays, 1997
).
Kinesin light chain and Sunday driver are not
required for ooplasmic streaming and posterior localisation
The discovery that the KHC is required in the oocyte for the posterior
localisation of oskar mRNA, the posterior localisation of DHC and for
cytoplasmic streaming raises the question of whether these reflect three
independent functions of the motor, or whether they all depend on a common
underlying process. One way to address this question is to determine whether
the three functions require different factors to couple kinesin to its
cargoes. One of the main cargo adaptors in Drosophila neurones is
Sunday driver (Syd), as syd mutants cause the same defects in axonal
transport as null mutants in either the Kinesin heavy chain or light chain
(Bowman et al., 2000). We
therefore examined the phenotypes of syd null germline clones. The
posterior localisations of both Staufen and DHC are indistinguishable from
wild-type in syd mutants (Fig.
5A,B,D,E). Furthermore, the absence of Syd has no effect on the
rate of cytoplasmic streaming at either stage 9 or stage 10b
(Fig. 6A-C; I. M. P. and D. St
J., unpublished) (Table 1).
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The lack of a phenotype in syd germline clones prompted us to
investigate the role of the KLC in these processes, by generating germline
clones of a hypomorphic allele, klc1, or a null allele,
klc8ex94, which is a deletion of the entire coding region
(Gindhart et al., 1998).
Staufen still forms a normal posterior crescent in 100% of both the
klc1 and the klc8ex94 mutant egg
chambers (Fig. 5C). Occasionally, however, it can also be found in a dot near the posterior of the
oocyte, which is often connected to the posterior crescent (I. M. P. and D. St
J., unpublished). DHC also localises to the posterior in the absence of the
KLC, although the amount is reduced compared with wild type
(Fig. 5F). These subtle
phenotypes are completely different from those caused by loss of the KHC, and
indicate that the light chain may be required for the efficiency of posterior
localisation, but that it is not essential for this process. One possible
explanation for the lack of a requirement for the KLC is that the phenotype is
rescued by the perdurance of the wild-type protein that is synthesised in the
heterozygous germline stem cells before the clones were induced. To test this
possibility, we examined egg chambers from germline clones that had been
induced two weeks earlier, and observed the same effects on oskar
mRNA and DHC localisation. As the germline stem cells divide every 12-16
hours, they should have gone through over 20 divisions in this period, ruling
out the possibility that any wild-type protein survives.
Finally, we examined the cytoplasmic streaming in Klc mutant oocytes. Unlike the heavy chain, the KLC is not required for streaming at either stage 9 or stage 10b, although the movements are often less vigorous than in wild type (Fig. 6D-F; I. M. P. and D. St J., unpublished). Thus, the KLC is therefore dispensable for the three KHC-dependent processes that we have examined in the oocyte, suggesting that they may be related.
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DISCUSSION |
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A second essential function of the light chain is to couple the heavy chain
to its cargoes (Kamal and Goldstein,
2002). A genetic screen for mutants that disrupt axonal transport
led to the identification of a highly conserved membrane protein, Syd, which
binds to the TPR repeats of the KLC (Bowman
et al., 2000
). As syd mutants cause the same defects as
null mutants in the Klc and Khc, and the protein is found on
vesicles, it is likely to be a major cargo adaptor in axonal transport. The
mammalian homologue of Syd, JIP3/JSAP1, was isolated in an independent screen
for cargo adaptors, along with two other Jun N-terminal kinase interacting
proteins, JIP1 and JIP2, which also bind to the TPR domains of the light chain
and link the motor to several cytoplasmic and transmembrane protein cargoes
(Verhey et al., 2001
). The
amyloid precursor protein (APP) and the related APP-like proteins constitute a
second family of cargo adaptors that bind to the same region of the light
chain (Kamal et al., 2000
).
Studies in both mice and Drosophila have shown that these proteins
are required to couple kinesin to specific vesicular cargoes that are
transported along axons (Gunawardena and
Goldstein, 2001
; Kamal et al.,
2001
). The heavy chain probably makes some contribution to
cargo-binding, because it has been shown to interact with microsomal membranes
(Skoufias et al., 1994
). In
addition, the microsomal transmembrane protein Kinectin co-purifies with
kinesin, and binds directly to the C-terminal region of the heavy chain
(Kumar et al., 1995
;
Ong et al., 2000
;
Toyoshima et al., 1992
). There
is no Kinectin homologue in C. elegans or Drosophila,
however, whereas the mouse Kinectin knock out is viable and fertile, and shows
no obvious defects in kinesindependent transport processes
(Plitz and Pfeffer, 2001
).
Thus, all known specific cargo interactions with kinesin are mediated by the
light chain.
In light of the results above, it is very surprising that the light chain
is dispensable for the three functions of kinesin in the Drosophila
female germline. One trivial explanation is that there is a second light chain
gene in Drosophila, but this seems highly unlikely for several
reasons. First, the protein is not redundant in the nervous system, as a
strong axonal transport phenotype is observed in Klc mutants
(Gindhart et al., 1998).
Second, there is only one light chain gene in the `complete'
Drosophila genome sequence (63% sequence identity to human kinesin
light chain 1), and all of the light chain cDNAs in the extensive
Drosophila EST collections correspond with this gene
(Adams et al., 2000
;
Goldstein and Gunawardena,
2000
). Third, the `complete' genome sequence of another Dipteran
insect, the mosquito Anopheles gambiae, also contains only a single
Klc gene. Although it is possible that there is a second light chain
gene in the small region of each genome that has not been sequenced, it seems
very improbable that this would be the case in both organisms. Thus, our
results strongly suggest that the kinesin heavy chain can function without a
light chain in the oocyte, and that it must therefore interact with its cargo
or cargoes in some other way.
Although there is no precedent for light chain independent activities of
the KHC in higher eukaryotes, the distantly related kinesin heavy chains of
fungi, such as Neurospora crassa, function without any associated
light chains (Steinberg and Schliwa,
1995). Mutagenesis studies on the N. crassa kinesin have
identified a putative cargo-binding domain in the tail, and this region has
been conserved in animal KHCs (Seiler et
al., 2000
). It may therefore represent an alternative
cargo-binding domain that could account for the light chain independence of
the KHC functions in the oocyte. Interestingly, the glutamate receptor
interacting protein, GRIP1, has recently been shown to bind to this region of
the mouse KHC (Setou et al.,
2002
). GRIP1 has been proposed to target kinesin to dendrites, and
it is not yet known whether it functions as a cargo adaptor, or plays a role
in light chain independent transport.
Kinesin heavy chain is required for all ooplasmic streamings during
oogenesis
Twenty years ago it was suggested that the vigorous ooplasmic streaming and
the cytoplasmic movements in the nurse cells in stage 10b egg chambers are
independent processes (Gutzeit and Koppa,
1982). Our results demonstrate that this is indeed the case, not
only at stage 10b, but also earlier in oogenesis, as ooplasmic streaming is
completely abolished in Khc mutant egg chambers, whereas the
cytoplasmic movements in the nurse cells and from the nurse cells into the
oocyte are unaffected. It is unclear how kinesin creates these cytoplasmic
flows in the oocyte. Given its role in vesicle transport in other systems, an
attractive model is that it transports some organelle or vesicle along
microtubules, and that this then generates flows in the surrounding cytoplasm,
because of its viscosity. It seems unlikely that kinesin is directly
transporting any of the particles or vesicles that we have visualised in our
assays, as these particles move at speeds of about 0.1 µm/second at stage
9, which is significantly slower than other reported kinesin-dependent
transport processes (Goldstein and Yang,
2000
). This suggests that kinesin generates streaming by
transporting some other organelle or vesicle more rapidly along the
microtubules.
The nature of the cytoplasmic flows in the oocyte is variable and
temporally regulated (Theurkauf,
1994). The ooplasmic streaming at stage 9 is slow and
uncoordinated, whereas the movements at stage 10b are faster and
unidirectional, and resemble those of a `washing machine'. As both types of
ooplasmic streaming are completely abolished in Khc mutants, these
differences cannot be due to the motor protein. The type of streaming probably
depends, at least in part, on the organisation of the microtubule
cytoskeleton, which changes completely at the beginning of stage 10b, but
kinesin may also have distinct cargoes at the two stages, which could
influence the strength of the cytoplasmic flows.
The role of the Kinesin heavy chain in oskar mRNA
localisation?
In an attempt to understand the mechanism for oskar mRNA transport
to the posterior, we analysed the movement of a GFP-Staufen fusion protein in
living oocytes. Although this fusion protein localises to the posterior with
oskar mRNA and rescues the oskar mRNA localisation defect of
a staufen null mutant, we have been unable to resolve any movements
that unambiguously correspond to posterior transport. One possible explanation
for this failure is that most of the fluorescent GFP-Staufen particles do not
contain oskar mRNA, which is expressed at much lower levels than the
fusion protein. Thus, the relevant oskar mRNA/GFP-Staufen complexes
may be too rare or too weakly fluorescent to follow in time-lapse films.
Although we have been unable to determine how GFP-Staufen reaches the
posterior, our results do reveal several important features of this process
that are relevant to the discussion of the models for the mechanism of
oskar mRNA localisation.
One model proposes that cytoplasmic flows circulate oskar mRNA
around the oocyte, so that it can then be efficiently trapped at the posterior
by a pre-localised cortical anchor
(Glotzer et al., 1997).
Indeed, this mechanism would account for our failure to detect any directed
transport of GFP-Staufen to the posterior pole. Our observation that the KHC
is required for all cytoplasmic flows in the oocyte also supports this model,
as it provides an explanation for why the KHC is required to localise
oskar mRNA. However, several other considerations make this mechanism
unlikely. First, the cytoplasmic flows are much weaker at the posterior of the
oocyte than elsewhere, presumably because there are fewer microtubules in this
region, and many oocytes show little or no cytoplasmic movement near the
posterior pole. It is therefore hard to imagine how cytoplasmic flows could
efficiently deliver the mRNA to a posterior anchor. Second, the hypothetical
anchor would have to localise to the posterior before oskar mRNA and
in an oskar mRNA independent manner, and no proteins that meet these
criteria have been identified so far. Indeed, the only proteins that fulfil
the second criterion are the KHC and the components of the dynein/dynactin
complex. Third, oskar mRNA localises to the centre of the oocyte in
mutants that alter the organisation of the microtubule cytoskeleton, such as
gurken, pka and par-1, and it is hard to reconcile this with
trapping by a cortical anchor, as there is no plasma membrane or cortical
cytoskeleton in this region
(González-Reyes et al.,
1995
; Lane and Kalderon,
1994
; Roth et al.,
1995
; Shulman et al.,
2000
; Tomancak et al.,
2000
). The localisation of oskar mRNA still correlates
with the position of microtubule plus ends in these mutants, because
Kin-ßGal forms a dot in the centre of the oocyte with the mRNA, and this
is more consistent with the model in which oskar mRNA is transported
along microtubules towards the posterior pole. Finally, the KHC accumulates at
the posterior during the stages when oskar mRNA and DHC are
localised, strongly suggesting that it plays a direct role in transporting
them there.
Another model for oskar mRNA localisation proposes that the KHC
functions to transport the RNA away from the minus ends of the microtubules at
the anterior and lateral cortex towards the plus ends in the interior of the
oocyte, and that the lack of microtubules at the posterior somehow allows the
mRNA to accumulate at this pole (Cha et
al., 2002). Two aspects of our data do not fit this cortical
exclusion model. First, unlike Cha et al., we never saw any oskar
mRNA or Staufen at the posterior of the oocyte in Khc germline
clones, regardless of whether we performed fluorescent or wholemount in situ
hybridisation or antibody staining. This observation seems incompatible with a
model in which kinesin removes oskar mRNA from the anterior and
lateral cortex, but is not required for its localisation to the posterior
pole. Second, the demonstration that endogenous kinesin localises to the
posterior cortex, like kinesin-ßGal, provides further evidence that the
plus ends of the microtubules are enriched in this region, and strongly
suggests that kinesin mediates transport to this pole. These localisations are
not visible until stage 9, however, which is when oskar mRNA starts
to accumulate at the posterior. Thus, our results can be reconciled with those
of Cha et al., by proposing that the plus ends lie in the middle of the oocyte
at stage 8, when they observe a kinesin-dependent accumulation of
oskar mRNA in the central dot, and that they are only recruited to
the posterior at stage 9, coincident with the onset of oskar mRNA
localisation.
In light of the posterior localisation of endogenous kinesin, we think it
most likely that this motor does transport oskar mRNA to the
posterior of the oocyte, even though we have been unable to see this movement.
The link between the KHC and the oskar mRNA localisation complex need
not be direct, however. The KHC probably transports something else to the
posterior of the oocyte, in addition to oskar mRNA and dynein,
because mutants that abolish either oskar mRNA localisation (such as
staufen and barentsz) or DHC localisation
(Dhc64C6-6/Dhc64C6-12) have no effect on the
posterior localisation of the KHC, even though the motor activity of the KHC
is thought to require binding to a cargo. The KHC is also required for
cytoplasmic streaming, and presumably induces these flows by moving a large
structure, such as a vesicle or organelle, along microtubules. This structure
should therefore accumulate at the posterior of the oocyte during stage 9,
because this is where the microtubule plus ends and the KHC itself localise.
Thus, oskar mRNA and dynein could reach the posterior at stage 9 by
hitch-hiking on the large cargo that drives streaming. This proposal is
consistent with several other observations. First, the fact that cytoplasmic
streaming, oskar mRNA localisation and dynein localisation all share
the very unusual property of being light chain independent suggests that they
all depend on a single KHC-mediated transport process, which could be the
transport of the cargo that induces streaming to the posterior. Second, it has
been shown in a number of other systems that plus and minus end directed
microtubule motors, such as kinesin and dynein, are found on the same
organelles (Gross et al.,
2002; Martin et al.,
1999
; Welte et al.,
1998
). Third, if dynein and oskar mRNA interact with the
kinesin cargo independently of each other, this would explain why both their
posterior localisations require the KHC, but do not require each other.
Finally, there is already evidence that links oskar mRNA localisation
with vesicle trafficking, as mutants in rab11, a small GTPase
implicated in the regulation of endocytic vesicle recycling, disrupt the
posterior localisation of oskar mRNA
(Dollar et al., 2002
;
Jankovics et al., 2002). Furthermore, Rab11 itself localises to the posterior
of the oocyte. The effect of Rab11 on oskar mRNA localisation may be
indirect, however, as these mutants also disrupt the organisation of the
microtubule cytoskeleton.
It is unclear why dynein localises to the posterior, but one possibility is that it is needed to recycle kinesin to the minus ends of the microtubules, so that it can mediate another round of posterior localisation. The only known phenotype of the Dhc64C mutants that specifically disrupt the posterior localisation of DHC is a reduction in the rate of cytoplasmic streaming, and this may due to the gradual depletion of the pool of KHC available for transport. However, this localisation may be important for recycling dynein away from the minus ends of microtubules, so that it can mediate further rounds of minus end-directed transport.
If the hitch-hiking model for oskar mRNA localisation is correct,
Staufen, Barentsz, Mago nashi and Y14 would be required to couple the mRNA to
the vesicle or organelle that is transported by kinesin. In this context, it
is interesting to note that mammalian Staufen homologues have been shown to
associate with the endoplasmic reticulum
(Kiebler et al., 1999;
Marión et al., 1999
;
Wickham et al., 1999
). The
localisation of Vg1 mRNA to the vegetal pole of Xenopus oocytes
requires the RNA-binding protein VERA/Vg1 RBP, which co-fractionates with
markers for the endoplasmic reticulum, and this has led to the suggestion that
Vg1 mRNA is transported in association with ER vesicles
(Deshler et al., 1997
). Thus,
hitchhiking on vesicles may represent a general mechanism for mRNA
transport.
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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---|
Adams, M. D., Celniker, S. E., Holt, R. A., Evans, C. A.,
Gocayne, J. D., Amanatides, P. G., Scherer, S. E., Li, P. W., Hoskins, R. A.,
Galle, R. F. et al. (2000). The genome sequence of
Drosophila melanogaster. Science
287,2185
-2195.
Bohrmann, J. and Biber, K. (1994).
Cytoskeleton-dependent transport of cytoplasmic particiles in previtellogenic
to mid-vitellogenic ovarian follicles of Drosophila: time-lapse
analysis using video-enhanced contrast microscopy. J. Cell
Sci. 107,849
-858.
Bolivar, J., Huynh, J. R., Lopez-Schier, H., Gonzalez, C., St
Johnston, D. and Gonzalez-Reyes, A. (2001). Centrosome
migration into the Drosophila oocyte is independent of BicD and egl, and of
the organisation of the microtubule cytoskeleton.
Development 128,1889
-1897.
Boswell, R. E., Prout, M. E. and Steichen, J. C. (1991). Mutations in a newly identified Drosophila melanogaster gene, mago nashi, disrupt germ cell formation and result in the formation of mirror-image symmetrical double abdomen embryos. Development 113,373 -384.[Abstract]
Bowman, A. B., Kamal, A., Ritchings, B. W., Philp, A. V., McGrail, M., Gindhart, J. G. and Goldstein, L. S. (2000). Kinesin-dependent axonal transport is mediated by the sunday driver (SYD) protein. Cell 103,583 -594.[Medline]
Brady, S. T. (1985). A novel brain ATPase with properties expected for the fast axonal transport motor. Nature 317,73 -75.[Medline]
Brady, S. T., Pfister, K. K. and Bloom, G. S. (1990). A monoclonal antibody against kinesin inhibits both anterograde and retrograde fast axonal transport in squid axoplasm. Proc. Natl. Acad. Sci. USA 87,1061 -1065.[Abstract]
Brendza, R. P., Serbus, L. R., Duffy, J. B. and Saxton, W.
M. (2000). A function for kinesin I in the posterior
transport of oskar mRNA and Staufen protein. Science
289,2120
-2122.
Cha, B., Koppetsch, B. S. and Theurkauf, W. E. (2001). In vivo analysis of Drosophila bicoid mRNA localization reveals a novel microtubule-dependent axis specification pathway. Cell 106,35 -46.[Medline]
Cha, B. J., Serbus, L. R., Koppetsch, B. S. and Theurkauf, W. E. (2002). Kinesin I-dependent cortical exclusion restricts pole plasm to the oocyte posterior. Nat. Cell Biol. 22, 22.
Chou, T.-B. and Perrimon, N. (1996). The
autosomal FLP-DFS technique for generating germline mosaics in Drosophila
melanogaster. Genetics 144,1673
-1679.
Chou, T.-B., Noll, E. and Perrimon, N. (1993).
Autosomal P[ovoD1] dominant female-sterile insertions in
Drosophila and their use in generating germ-line chimeras.
Development 119,1359
-1369.
Clark, I., Giniger, E., Ruohola-Baker, H., Jan, L. and Jan, Y. (1994). Transient posterior localisation of a kinesin fusion protein reflects anteroposterior polarity of the Drosophila oocyte. Curr. Biol. 4,289 -300.[Medline]
Clark, I., Jan, L. Y. and Jan, Y. N. (1997).
Reciprocal localization of Nod and kinesin fusion proteins indicates
microtubule polarity in the Drosophila oocyte, epithelium, neuron and
muscle. Development 124,461
-470.
Coy, D. L., Hancock, W. O., Wagenbach, M. and Howard, J. (1999). Kinesin's tail domain is an inhibitory regulator of the motor domain. Nat. Cell Biol. 1, 288-292.[CrossRef][Medline]
Deshler, J. O., Highett, M. I. and Schnapp, B. J.
(1997). Localization of Xenopus Vg1 mRNA by Vera protein and the
endoplasmic reticulum [see comments]. Science
276,1128
-1131.
Dollar, G., Strukhoff, E., Michaud, J. and Cohen, R. S.
(2002). Rab11 polarization of the Drosophila oocyte: a
novel link between membrane trafficking, microtubule organization, and
oskar mRNA localization and translation.
Development 129,517
-526.
Ephrussi, A., Dickinson, L. K. and Lehmann, R. (1991). oskar organizes the germ plasm and directs localization of the posterior determinant nanos. Cell 66, 37-50.[Medline]
Ephrussi, A. and Lehmann, R. (1992). Induction of germ cell formation by oskar. Nature 358,387 -392.[CrossRef][Medline]
Erdélyi, M., Michon, A., Guichet, A., Bogucka Glotzer, J. and Ephrusssi, A. (1995). A requirement for Drosophila cytoplasmic tropomyosin in oskar mRNA localization. Nature 377,524 -527.[CrossRef][Medline]
Ferrandon, D., Elphick, L., Nüsslein-Volhard, C. and St Johnston, D. (1994). Staufen protein associates with the 3'UTR of bicoid mRNA to form particles which move in a microtubule-dependent manner. Cell 79,1221 -1232.[Medline]
Friedman, D. S. and Vale, R. D. (1999). Single-molecule analysis of kinesin motility reveals regulation by the cargo-binding tail domain. Nat. Cell Biol. 1, 293-297.[CrossRef][Medline]
Gauger, A. K. and Goldstein, L. S. (1993). The
Drosophila kinesin light chain. Primary structure and interaction with kinesin
heavy chain. J. Biol. Chem.
268,13657
-13666.
Gindhart, J. G., Jr, Desai, C. J., Beushausen, S., Zinn, K. and
Goldstein, L. S. (1998). Kinesin light chains are essential
for axonal transport in Drosophila. J. Cell Biol.
141,443
-454.
Gindhart, J. G., Jr and Goldstein, L. S. (1996). Tetratrico peptide repeats are present in the kinesin light chain. Trends Biochem. Sci. 21, 52-53.[CrossRef][Medline]
Glotzer, J. B., Saffrich, R., Glotzer, M. and Ephrussi, A. (1997). Cytoplasmic flows localize injected oskar RNA in Drosophila oocytes. Curr. Biol. 7, 326-337.[Medline]
Goldstein, L. S. and Gunawardena, S. (2000). Flying through the drosophila cytoskeletal genome. J. Cell Biol. 150,F63 -F68.[Medline]
Goldstein, L. S. and Yang, Z. (2000). Microtubule-based transport systems in neurons: the roles of kinesins and dyneins. Annu. Rev. Neurosci. 23, 39-71.[CrossRef][Medline]
González-Reyes, A., Elliott, H. and St Johnston, D. (1995). Polarization of both major body axes in Drosophila by gurken-torpedo signalling. Nature 375,654 -658.[CrossRef][Medline]
Gross, S. P., Tuma, M. C., Deacon, S. W., Serpinskaya, A. S.,
Reilein, A. R. and Gelfand, V. I. (2002). Interactions and
regulation of molecular motors in Xenopus melanophores. J. Cell
Biol. 156,855
-865.
Gunawardena, S. and Goldstein, L. S. (2001). Disruption of axonal transport and neuronal viability by amyloid precursor protein mutations in Drosophila. Neuron 32,389 -401.[Medline]
Gutzeit, H. (1986a). The role of microfilaments in cytoplasmic streaming in Drosophila follicles. J. Cell Sci. 80,159 -169.[Abstract]
Gutzeit, H. (1986b). The role of microtubules in the differentiation of ovarian follicles during vitellogenesis in Drosophila. Roux's Arch. Dev. Biol. 195,173 -181.
Gutzeit, H. O. and Koppa, R. (1982). Time-lapse film analysis of cytoplasmic streaming during late oogenesis of Drosophila. J. Embryol. Exp. Morphol. 67,101 -111.
Hachet, O. and Ephrussi, A. (2001). Drosophila Y14 shuttles to the posterior of the oocyte and is required for oskar mRNA transport. Curr. Biol. 11,1666 -1674.[CrossRef][Medline]
Hackney, D. D. (1994). The rate-limiting step
in microtubule-stimulated ATP hydrolysis by dimeric kinesin head domains
occurs while bound to the microtubule. J. Biol. Chem.
269,16508
-16511.
Hackney, D. D., Levitt, J. D. and Wagner, D. D. (1991). Characterization of alpha 2 beta 2 and alpha 2 forms of kinesin. Biochem. Biophys. Res. Commun. 174,810 -815.[Medline]
Hurd, D. D. and Saxton, W. M. (1996). Kinesin
mutations cause motor neuron disease phenotypes by disrupting fast axonal
transport in Drosophila. Genetics
144,1075
-1085.
Jankovics, F., Sinka, R. and Erdelyi, M.
(2001). An interaction type of genetic screen reveals a role of
the Rab11 gene in oskar mRNA localization in the developing
Drosophila melanogaster oocyte. Genetics
158,1177
-1188.
Kamal, A. and Goldstein, L. S. (2000). Connecting vesicle transport to the cytoskeleton. Curr. Opin. Cell Biol. 12,503 -508.[CrossRef][Medline]
Kamal, A. and Goldstein, L. S. (2002). Principles of cargo attachment to cytoplasmic motor proteins. Curr. Opin. Cell Biol. 14,63 -68.[CrossRef][Medline]
Kamal, A., Stokin, G. B., Yang, Z., Xia, C. H. and Goldstein, L. S. (2000). Axonal transport of amyloid precursor protein is mediated by direct binding to the kinesin light chain subunit of kinesin-I. Neuron 28,449 -459.[Medline]
Kamal, A., Almenar-Queralt, A., LeBlanc, J. F., Roberts, E. A. and Goldstein, L. S. (2001). Kinesin-mediated axonal transport of a membrane compartment containing beta-secretase and presenilin-1 requires APP. Nature 414,643 -648.[CrossRef][Medline]
Kiebler, M. A., Hemraj, I., Verkade, P., Kohrmann, M., Fortes,
P., Marion, R. M., Ortin, J. and Dotti, C. G. (1999). The
mammalian staufen protein localizes to the somatodendritic domain of cultured
hippocampal neurons: implications for its involvement in mRNA transport.
J. Neurosci. 19,288
-297.
Kim-Ha, J., Smith, J. L. and Macdonald, P. M. (1991). oskar mRNA is localized to the posterior pole of the Drosophila oocyte. Cell 66, 23-35.[Medline]
Kumar, J., Yu, H. and Sheetz, M. P. (1995). Kinectin, an essential anchor for kinesin-driven vesicle motility. Science 267,1834 -1837.[Medline]
Kuznetsov, S. A., Vaisberg, Y. A., Rothwell, S. W., Murphy, D.
B. and Gelfand, V. I. (1989). Isolation of a 45-kDa fragment
from the kinesin heavy chain with enhanced ATPase and microtubule-binding
activities. J. Biol. Chem.
264,589
-595.
Lane, M. E. and Kalderon, D. (1994). RNA localization along the anteroposterior axis of the Drosophila oocyte requires PKA-mediated signal transduction to direct normal microtubule organization. Genes Dev. 8,2986 -2995.[Abstract]
Li, M.-g., McGrail, M., Serr, M. and Hays, T. H. (1994). Drosophila cytoplasmic dynein, a microtubule motor that is asymmetrically localized in the oocyte. J. Cell Biol. 126,1475 -1494.[Abstract]
Marión, R. M., Fortes, P., Belosco, A., Dotti, C. and
Ortín, J. (1999). A human sequence homologue of
Staufen is an RNA-binding protein that is associated with polysomes and
localizes to the rough endoplasmic reticulum. Mol. Cell.
Biol. 19,2212
-2219.
Martin, M., Iyadurai, S. J., Gassman, A., Gindhart, J. G., Jr,
Hays, T. S. and Saxton, W. M. (1999). Cytoplasmic dynein, the
dynactin complex, and kinesin are interdependent and essential for fast axonal
transport. Mol. Biol. Cell
10,3717
-3728.
McGrail, M. and Hays, T. S. (1997). The
microtubule motor cytoplasmic dynein is required for spindle orientation
during germline cell divisions and oocyte differentiation in Drosophila.Development 124,2409
-2419.
Micklem, D. R., Dasgupta, R., Elliott, H., Gergely, F., Davidson, C., Brand, A., González-Reyes, A. and St Johnston, D. (1997). The mago nashi gene is required for the polarisation of the oocyte and the formation of perpendicular axes in Drosophila. Curr. Biol. 7, 468-478.[Medline]
Mohr, S. E., Dillon, S. T. and Boswell, R. E.
(2001). The RNA-binding protein Tsunagi interacts with Mago Nashi
to establish polarity and localize oskar mRNA during Drosophila oogenesis.
Genes Dev. 15,2886
-2899.
Newmark, P. A. and Boswell, R. E. (1994). The
mago nashi locus encodes an essential product required for germ plasm
assembly in Drosophila. Development
120,1303
-1313.
Newmark, P. A., Mohr, S. A., Gong, L. and Boswell, R. E.
(1997). mago nashi mediates the posterior follicle
cell-to-oocyte signal to organize axis formation in Drosophila.Development 124,3197
-3207.
Ong, L. L., Lim, A. P., Er, C. P., Kuznetsov, S. A. and Yu,
H. (2000). Kinectin-kinestin binding domains and their
effects on organelle motility. J. Biol. Chem.
275,32854
-32860.
Plitz, T. and Pfeffer, K. (2001). Intact
lysosome transport and phagosome function despite kinectin deficiency.
Mol. Cell. Biol. 21,6044
-6055.
Rahman, A., Kamal, A., Roberts, E. A. and Goldstein, L. S.
(1999). Defective kinesin heavy chain behavior in mouse kinesin
light chain mutants. J. Cell Biol.
146,1277
-1288.
Ramos, A., Grunert, S., Adams, J., Micklem, D. R., Proctor, M.
R., Freund, S., Bycroft, M., St Johnston, D. and Varani, G.
(2000). RNA recognition by a Staufen double-stranded RNA-binding
domain. EMBO J. 19,997
-1009.
Riechmann, V. and Ephrussi, A. (2001). Axis formation during Drosophila oogenesis. Curr. Opin. Genet. Dev. 11,374 -383.[CrossRef][Medline]
Roth, S., Neuman-Silberberg, F. S., Barcelo, G. and Schüpbach, T. (1995). cornichon and the EGF receptor signaling process are necessary for both anterior-posterior and dorsal-ventral pattern formation in Drosophila. Cell 81,967 -978.[Medline]
Ruohola, H., Bremer, K. A., Baker, D., Sedlow, J. R., Jan, L. Y. and Jan, Y. N. (1991). Role of neurogenic genes in establishment of follicle cell fate and oocyte polarity during oogenesis in Drosophila. Cell 66,433 -449.[Medline]
Saxton, W. M., Hicks, J., Goldstein, L. S. B. and Raff, E. C. (1991). Kinesin heavy chain is essential for viability and neuromuscular functions in Drosophila, bu mutants show no defects in mitosis. Cell 64,1093 -1102.[Medline]
Schuldt, A. J., Adams, J. H. J., Davidson, C. M., Micklem, D.
R., St Johnston, D. and Brand, A. (1998). Miranda mediates
the asymmetric protein and RNA localisation in the developing nervous system.
Genes Dev. 12,1847
-1857.
Seiler, S., Kirchner, J., Horn, C., Kallipolitou, A., Woehlke, G. and Schliwa, M. (2000). Cargo binding and regulatory sites in the tail of fungal conventional kinesin. Nat. Cell Biol. 2,333 -338.[CrossRef][Medline]
Setou, M., Seog, D., Tanaka, Y., Kanai, Y., Takei, Y., Kawagishi, M. and Hirokawa, N. (2002). Glutamate-receptor-interacting protein GRIP1 directly steers kinesin to dendrites. Nature 417,83 -87.[Medline]
Shulman, J. M., Benton, R. and St. Johnston, D. (2000). The Drosophila homolog of C. elegans PAR-1 organizes the oocyte cytoskeleton and directs oskar mRNA localisation to the posterior pole. Cell 101, 1-20.[Medline]
Skoufias, D. A., Cole, D. G., Wedaman, K. P. and Scholey, J.
M. (1994). The carboxyl-terminal domain of kinesin heavy
chain is important for membrane binding. J. Biol.
Chem. 269,1477
-1485.
St Johnston, D., Beuchle, D. and Nüsslein-Volhard, C. (1991). Staufen, a gene required to localize maternal RNAs in the Drosophila egg. Cell 66, 51-63.[Medline]
St Johnston, D., Brown, N. H., Gall, J. G. and Jantsch, M. (1992). A conserved double-stranded RNA-binding domain. Proc. Natl. Acad. Sci. USA 89,10979 -10983.[Abstract]
Steinberg, G. and Schliwa, M. (1995). The Neurospora organelle motor: a distant relative of conventional kinesin with unconventional properties. Mol. Biol. Cell 6,1605 -1618.[Abstract]
Stenoien, D. L. and Brady, S. T. (1997). Immunochemical analysis of kinesin light chain function. Mol. Biol. Cell 8,675 -689.[Abstract]
Stewart, R. J., Thaler, J. P. and Goldstein, L. S. (1993). Direction of microtubule movement is an intrinsic property of the motor domains of kinesin heavy chain and Drosophila ncd protein. Proc. Natl. Acad. Sci. USA 90,5209 -5213.[Abstract]
Tetzlaff, M. T., Jäckle, H. and Pankratz, M. J. (1996). Lack of Drosophila cytoskeletal tropomyosin affects head morphogenesis and the accumulation of oskar messenger RNA required for germ cell formation. EMBO J. 15,1247 -1254.[Abstract]
Theurkauf, W. E. (1994). Premature microtubule-dependent cytoplasmic streaming in cappuccino and spire mutant oocytes. Science 265,2093 -2096.[Medline]
Theurkauf, W. E. and Hazelrigg, T. I. (1998).
In vivo analyses of cytoplasmic transport and cytoskeletal organization during
Drosophila oogenesis: characterization of a multi-step anterior localization
pathway. Development
125,3655
-3666.
Theurkauf, W. E., Smiley, S., Wong, M. L. and Alberts, B. M.
(1992). Reorganization of the cytoskeleton during
Drosophila oogenesis: implications for axis specification and
intercellular transport. Development
115,923
-936.
Tomancak, P., Piano, F., Riechmann, V., Gunsalus, K. C., Kemphues, K. J. and Ephrussi, A. (2000). A Drosophila melanogaster homologue of Caenorhabditis elegans par-1 acts at an early step in embryonic-axis formation. Nat. Cell Biol. 2,458 -460.[CrossRef][Medline]
Toyoshima, I., Yu, H., Steuer, E. R. and Sheetz, M. P. (1992). Kinectin, a major kinesin-binding protein on ER. J. Cell Biol. 118,1121 -1131.[Abstract]
Vale, R. D., Reese, T. S. and Sheetz, M. P. (1985). Identification of a novel force-generating protein, kinesin, involved in microtubule based motility. Cell 42, 39-50.[Medline]
van Eeden, F. and St Johnston, D. (1999). The polarisation of the anterior-posterior and dorsal-ventral axes during Drosophila oogenesis. Curr. Opin. Genet. Dev. 9, 396-404.[CrossRef][Medline]
van Eeden, F. J., Palacios, I. M., Petronczki, M., Weston, M. J.
and St Johnston, D. (2001). Barentsz is essential for the
posterior localization of oskar mRNA and colocalizes with it to the posterior
pole. J. Cell Biol. 154,511
-523.
Verhey, K. J., Lizotte, D. L., Abramson, T., Barenboim, L.,
Schnapp, B. J. and Rapoport, T. A. (1998). Light
chain-dependent regulation of Kinesin's interaction with microtubules.
J. Cell Biol. 143,1053
-1066.
Verhey, K. J., Meyer, D., Deehan, R., Blenis, J., Schnapp, B.
J., Rapoport, T. A. and Margolis, B. (2001). Cargo of kinesin
identified as JIP scaffolding proteins and associated signaling molecules.
J. Cell Biol. 152,959
-970.
Waterman-Storer, C. M., Karki, S. B., Kuznetsov, S. A., Tabb, J.
S., Weiss, D. G., Langford, G. M. and Holzbaur, E. L. (1997).
The interaction between cytoplasmic dynein and dynactin is required for fast
axonal transport. Proc. Natl. Acad. Sci. USA
94,12180
-12185.
Welte, M. A., Gross, S. P., Postner, M., Block, S. M. and Wieschaus, E. F. (1998). Developmental regulation of vesicle transport in Drosophila embryos: forces and kinetics. Cell 92,547 -557.[Medline]
Wickham, L., Duchaîne, T., Luo, M., Nabi, I. R. and
DesGrosseillers, L. (1999). Mammalian Staufen is a
double-stranded-RNA- and Tubulin-binding protein which localizes to the rough
endoplasmic reticulum. Mol. Cell. Biol.
19,2220
-2230.
Yang, J. T., Laymon, R. A. and Goldstein, L. S. B. (1989). A three-domain structure of kinesin heavy chain revealed by DNA sequence and microtubule binding analyses. Cell 56,879 -889.[Medline]