Department of Cell Biology and Anatomy, Graduate School of Medicine, University of Tokyo, Tokyo, 113, Japan
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Abstract |
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Cytoplasmic dynein, a minus end-directed,
microtubule-based motor protein, is thought to drive
the movement of membranous organelles and chromosomes. It is a massive complex that consists of multiple
polypeptides. Among these polypeptides, the cytoplasmic dynein heavy chain (cDHC) constitutes the major
part of this complex. To elucidate the function of cytoplasmic dynein, we have produced mice lacking cDHC
by gene targeting. cDHC/
embryos were indistinguishable from cDHC+/
or cDHC+/+ littermates at the
blastocyst stage. However, no cDHC
/
embryos were
found at 8.5 d postcoitum. When cDHC
/
blastocysts
were cultured in vitro, they showed interesting phenotypes. First, the Golgi complex became highly vesiculated and distributed throughout the cytoplasm. Second, endosomes and lysosomes were not concentrated
near the nucleus but were distributed evenly throughout the cytoplasm. Interestingly, the Golgi "fragments" and lysosomes were still found to be attached to microtubules.
These results show that cDHC is essential for the formation and positioning of the Golgi complex. Moreover, cDHC is required for cell proliferation and proper distribution of endosomes and lysosomes. However, molecules other than cDHC might mediate attachment of the Golgi complex and endosomes/lysosomes to microtubules.
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Introduction |
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MICROTUBULES are thought to play important roles
in various cellular activities, including mitosis,
determination of cellular shape, and the transport, organization, and distribution of organelles in higher
eukaryotic cells. The involvement of microtubules in the
transport, organization, and distribution of organelles requires the activity of motor proteins that interact with
both microtubules and organelles (Hirokawa 1993, 1998
;
Holzbaur and Vallee, 1994
; Schroer, 1994
). Among these
motor proteins, minus end-directed, microtubule-based
motor proteins are thought to drive the centrosomal movement of membranous organelles (Hirokawa 1993
,
1998
; Holzbaur and Vallee, 1994
; Schroer, 1994
). Cytoplasmic dynein (Lye et al., 1987
; Paschal et al., 1987
), the
most abundant minus end-directed motor protein, is a
high-molecular mass complex primarily composed of multiple polypeptides, two heavy chains of ~550,000 D, three to four 74,000-D intermediate chains, four light intermediate chains of ~55,000 D, and light chains of 8,000-22,000
D. The heavy chain, containing the sites for ATP hydrolysis and microtubule binding, probably makes up the bulk
of the head (Koonce et al., 1992
; Mikami et al., 1993
;
Zhang et al., 1993
), while the intermediate and light intermediate chains lie at the base of the molecule. Biochemical and immunocytochemical studies using anti-cytoplasmic dynein antibodies have shown that the motor is
localized on late endosomes and lysosomes (Lin and Collins, 1992
) and the TGN (Fath et al., 1994
), suggesting its
role in transporting these organelles in nonneuronal cells.
In neurons it is believed to be involved in retrograde axonal transport (Schnapp and Reese 1989
; Schroer et al.,
1989
; Hirokawa et al., 1990
). Furthermore, colocalization of cytoplasmic dynein with kinetochores and mitotic spindles suggested its possible involvement in mitosis (Pfarr et al.,
1990
; Steuer et al., 1990
).
To elucidate its function, several studies have been
performed by various researchers. Depletion of cytoplasmic dynein blocked the in vitro movement of endosomes/
lysosomes on microtubules (Blocker et al., 1997) and
prevented the centrosomal localization of exogenously applied Golgi-derived vesicles in semiintact cells (Corthesy-Theulaz et al., 1992
). Injection of antibodies into cells induced the collapse of mitotic spindles (Vaisberg et al., 1993
). Overexpression of the dynamitin subunit of the dynactin complex, which is believed to dissociate dyneins
from their target organelles, leads to dispersion of the
Golgi complex, endosomes, and lysosomes. The overexpression of dynamitin was also shown to block the ER to
Golgi transport (Burkhardt et al., 1997
; Presley et al.,
1997
). Recently, the molecular cloning of dynein genes by
reverse transcriptase PCR using degenerate primers has
resulted in the identification of several members of the cytoplasmic dynein family (Gibbons et al., 1994
; Tanaka et al.,
1995
; Vaisberg et al., 1996
). Using isoform-specific antibodies, immunolocalization of each isoform has revealed
that DHC2 (DLP4 of Tanaka and DHC1B of Gibbons) is
localized on the ER-Golgi intermediate compartment (Vaisberg, E.A., P.M. Grissom, S.R. Gill, T.A. Schroer,
and J.R. McIntosh. 1996. Mol. Biol. Cell. 7:403a) and
DHC3 (DLP2 of Tanaka and DHC7C of Gibbons) is localized on unidentified vesicular structures (Vaisberg et al.,
1996
). Injection of anti-DHC2 antibody leads to dispersion
of the Golgi complex (Vaisberg et al., 1996
), which is in
contrast to the findings of a previous injection experiment using anti-cytoplasmic dynein heavy chain antibody (Vaisberg et al., 1993
). Therefore, there is a possibility that previous immunolocalization and antibody injection studies
may not have discriminated the unique function of one cytoplasmic dynein member from that of another. Generating mutants that specifically lack each dynein gene is the
ideal way to determine the function of each cytoplasmic dynein heavy chain isoform. Mutational studies using
yeast (Eshel et al., 1993
; Li et al., 1993
) have shown the involvement of cytoplasmic dynein in spindle orientation
and anaphase chromosome segregation (Saunders et al.,
1995
). Fungal cytoplasmic dynein mutants exhibited abnormalities in nuclear positioning and movement (Plamann et al., 1994
; Xiang et al., 1994
), while Drosophila
(Gepner et al., 1996
) mutants were shown to be lethal, suggesting that cytoplasmic dynein is essential for proliferation. Although these studies have increased our knowledge regarding the functions of cytoplasmic dynein, its
role in intracellular transport remains elusive because of
the lack of appropriate mammalian mutants that lack the
functional cytoplasmic dynein heavy chain (cDHC)1 gene.
To elucidate this, we generated mice lacking cytoplasmic dynein, the major species of numerous minus end- directed motor proteins, by targeted disruption of its heavy chain (cDHC [DHC1 of Vaisberg and DHC1A of Gibbons]), which is essential for the functioning of cytoplasmic dynein.
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Materials and Methods |
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Generation of Mutant Mice
A 13-kb EcoRI fragment containing the cDHC gene was obtained from a
129/Sv mouse genomic library using a rat cDHC cDNA fragment, which
contains the first coding exon (Zhang et al., 1993) and was subcloned into
pBluescript. The sequence from the translation start ATG in the first coding exon was shown in Fig. 1 e. For construction of the targeting vector,
the second half of the first coding exon (after the EagI site in Fig. 1 e) and
a part of the following intron were deleted. The loxP sequence, splice-
acceptor/splice-enhancer (Friedrich and Soriano, 1991; Watakabe et al.,
1993
) sequences, and the PGK-neo gene (McBurney et al., 1991
) in the reverse transcriptional orientation to the cDHC gene were inserted into this
deleted region. For negative selection, a diphtheria toxin A (Yagi et al.,
1993
) fragment cassette was introduced into the downstream polylinker site. The replacement-type cDHC construct (Fig. 1 a) was linearized with
NotI before electroporation. The J1 line of ES cells used for these experiments were cultured essentially as described previously (Harada et al.,
1994
). ES cells carrying a disrupted cDHC gene were injected into
C57BL/6 embryos at the blastocyst stage as described previously (Harada
et al., 1994
).
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Immunoblotting
Immunoblotting was performed largely as described previously (Harada
et al., 1994). Wild-type (cDHC+/+) and heterozygous mutant (cDHC+/
)
ES cells were cultured as described. Cos, NRK, and NIH 3T3 cells were
grown in DME (GIBCO BRL, Gaithersburg, MD) supplemented with 10% FCS. All types of cells were grown to confluency, washed twice with
PBS, collected by scraping into ice-cold PBS, pelleted down by centrifugation, and suspended in 25 mM TrisCl, pH 6.8, 2% SDS. Cell suspension
was sonicated and then boiled to lower the viscosity. Crude extracts were
made by centrifuging the homogenates at 20,000 g for 15 min at 4°C. Protein concentrations were determined by BCA Protein Assay Reagent
(Pierce Chemical Co., Rockford, IL). Equal amounts of crude extracts
were separated with polyacrylamide gel. Proteins were electrophoretically
transferred to nylon filters (Millipore Corp., Bedford, MA). Nylon filters
with transferred proteins were blocked with 2% skim milk in TBS, incubated in anti-cDHC (DHC1) antisera or anti-DHC2 antisera (Vaisberg et
al., 1996
) (gifts from E.A.Vaisberg and J.R. McIntosh) for up to 6 h at
37°C, rinsed in TBS containing 0.05% Tween 20, and incubated for 2 h with 125I-labeled protein A. Binding was detected by autoradiography using an imaging analyzer (model BAS-2000; Fuji-Film, Tokyo, Japan).
Blastocyst Culture and Immunofluorescence
Blastocysts were collected by flushing the oviducts of female mice at 3.5 d postcoitum (dpc) and cultured individually for 3 d on gelatinized coverslips in embryonic stem (ES) cell medium. Cultured blastocysts were fixed in 2% paraformaldehyde, PBS for 10 min at room temperature (RT). Some samples were fixed in 2% paraformaldehyde and 0.2% glutaraldehyde for 10 min at RT and blocked in 1 mg/ml NaBH4 for 30 min at 4°C. They were permeabilized with cold methanol for 15 s, washed in PBS twice, blocked in 1% BSA/PBS for 5 min at RT, and then incubated with the first antibodies in the blocking solution for 1 h at 37°C. Subsequently, samples were extensively washed in PBS, blocked in 1% BSA/PBS for 5 min at RT, and incubated with the second antibodies for 1 h at 37°C. As for anti-Arp1 and Glued antibodies, samples were fixed in 2% paraformaldehyde, washed in PBS twice and in 20 mM glycine/PBS once, permeabilized in 0.1% saponin in 20 mM glycine/PBS for 20 min at RT, and incubated with the first antibody in 0.1% saponin in 20 mM glycine/PBS for 1 h at 37°C. The following washing, blocking, and incubation processes were the same as described above.
For the first antibodies, the following antibodies or antisera were used:
anti-cDHC antisera (Vaisberg et al., 1996) (a gift from E.A.Vaisberg and
J.R. McIntosh); anti-GM130 antisera (Nakamura et al., 1995
) (a gift from
N. Nakamura and G. Warren); CTR433 (Jasmin et al., 1989
) (a gift from
M. Bornens); anti-Arp1 monoclonal antibody (45A) (Schafer et al., 1994
)
and anti-p150Glued monoclonal antibody (150B) (Blocker et al., 1997
) (gifts
from T. Schroer); anti-PDI antibody (StressGen Biotechnologies Corp., Victoria, British Columbia); anti-LAMP2 antibody ABL-93 (Chen et al.,
1986
); DM1A (Sigma Chemical Co., St. Louis, MO); and YL1/2 (BIOSYS
S.A., Compiegne, France) for tubulin. The samples were observed either
using a confocal laser scanning microscope (model MRC-1000; Bio-Rad
Laboratories, Hercules, CA) or using an AXIOPHOT microscope (Carl
Zeiss, Inc., Thornwood, NY). Images were transferred to a Macintosh
computer for editing and were printed with a Fujix Pictrography Digital
Printer.
Conventional Electron Microscopy
Blastocysts grown on gelatinized coverslips in ES media for 3 d were
washed once with PBS and fixed in 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M cacodylate buffer for 2 h. Sections were processed as
described previously (Harada et al., 1990) and viewed under an electron
microscope (model 2000EX; JEOL, Inc., Tokyo, Japan) at 100 kV.
Genotyping of Blastocysts
Both intact and cultured blastocysts were genotyped by hemi-nested PCR
method. Cultured blastocysts were lysed in 50-100 µl lysis buffer (1%
SDS, 10 mM TrisCl, pH 8.0, 25 mM EDTA, 75 mM NaCl) containing proteinase K (100 µg/ml) for 10-20 min at RT and stored at 20°C until use.
For samples for electron microscopy, the inner cell masses (ICM) of fixed
cultured blastocysts were removed with a finely drawn glass micropipette
on an inverted microscope (model Diaphot; Nikon Corp., Tokyo, Japan),
washed with PBS once, lysed in the lysis buffer, and stored at
20°C until
use. The lysates were thawed, overlaid with mineral oil, heat-denatured at
95°C for 35 min, and then subjected to ethanol precipitation. Ethanol precipitates were dissolved in 10 µl distilled water and were used as PCR
templates. Intact blastocysts were lysed in 10 µl of autoclaved distilled water and frozen immediately at
20°C, and the lysates were directly used as
PCR templates. The first round PCR was carried out in a reaction buffer
containing 10 mM Tris-Cl, pH 8.3, 50 mM KCl, 1.5 mM MgCl2, 0.001%
(wt/vol) gelatin, 200 µM each of dATP, dCTP, dGTP, and dTTP, 1 µM
each of PCR primers, and 1 U of Taq DNA polymerase (Perkin-Elmer
Corp., Norwalk, CT) using three primers (Fig. 1a): the forward primer "a"
(5'-CCTGGGTGTTCAAGACAAGCTGGTTC-3'), located upstream of
the insertion site; the reverse primer "c" (5'-GTTATGGGGGCTGAGGTCTTGTCGT-3'), located near the 3' end of the first coding exon; and
the reverse primer "d" (5'-TAAGGGCCAGCTCATTCCTCCACT-3'),
located within the polyA signal of the PGK-neo gene. PCR schedules
were employed for 25 cycles: 94°C for 30 s, 63°C for 30 s, and 72°C for 90 s. For reamplification, 2 µl of the first round PCR mix was removed and
added to 48 µl of a fresh PCR mix containing nested primers (forward
primer "b" [5'-ACGCCCCTCACTGACCGTTGCTATT-3'] [located between a and the insertion site] and c or d) and amplified for an additional
30 cycles: 94°C for 30 s, 63°C for 30 s, and 72°C for 40 s.
Quantification of Lysosome Distribution
Cultured blastocysts were incubated for 8 h at 37°C in ES cell media containing 1 mg/ml FITC-dextran (Sigma Chemical Co.), washed with PBS, and fixed in 2% paraformaldehyde, PBS for 10 min. The samples were observed using a confocal laser scanning microscope (model MRC-100; Bio-Rad Laboratories) in the photon counting mode to ensure that the fluorescein intensity reflects the amount of FITC-dextran uptake, and the data were quantified using National Institutes of Health (NIH) Image software.
Taxol Treatment of Cultured Blastocysts
Taxol was obtained from the National Cancer Institute (Bethesda, MD)
and was dissolved in dimethyl sulfoxide at a concentration of 3.5 mM and
stored at 20°C. Blastocysts cultured for 3 d were incubated with ES media containing either 5 or 20 µM taxol for 24 h. Similar results were obtained under both conditions.
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Results |
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Targeting of the Cytoplasmic Dynein Heavy Chain Gene
To disrupt the endogenous cDHC gene, a replacement-type targeting vector was constructed (Fig. 1 a). Genomic
PCR and Southern blot analysis revealed a targeting frequency of 4 in ~600 G418R clones. To assess the levels of
cDHC and DHC2 in wild-type (cDHC+/+) and heterozygous mutant (cDHC+/) ES cells, we performed immunoblotting of crude extracts prepared from wild-type ES
cells, heterozygous mutant ES cells, and other cell lines
(NRK and Cos cells) using antibodies specific for cDHC and DHC2 (Vaisberg et al., 1996
). In wild-type ES cells,
cDHC is expressed at similar or slightly higher levels compared with other cell lines (NRK and Cos cells) (Fig. 1 d,
left, lanes 1-3). On the contrary, the amount of DHC2 in
wild-type ES cells is much lower than that in Cos cells,
which express high amounts of DHC2, and at similar levels
to that in NRK cells, which express the protein much
lower than Cos cells (Fig. 1 d, right, lanes 1-3) (see also
Vaisberg et al., 1996
). There was no apparent compensatory increase in DHC2 in cDHC+/
ES cells (Fig. 1 d, right,
lanes 3 and 4). In cDHC+/
ES cells, the amount of cDHC
was reduced to approximately one half that of cDHC+/+
ES cells (Fig. 1 d, left, lanes 4 and 5), and proteins of aberrant sizes were not detected with this antibody (Fig. 1 d,
left, lanes 6 and 7), suggesting that the allele generated by
this gene targeting experiment is likely to be null. Chimeric male mice generated from injections with three different cell lines transmitted the ES cell genome through
the germline. Approximately 50% of these agouti pups
were found to be heterozygous for the mutant cDHC allele when Southern blot analysis was carried out (data not
shown). These mice were indistinguishable from their
wild-type littermates and displayed no discernible abnormalities. To determine the phenotype of homozygous mutant mice, we interbred heterozygous animals and genotyped litters, but we could not identify cDHC
/
animals
either at birth or after weaning. When DNA isolated from fetuses at different gestation times from 8.5 to 12.5 dpc
(Fig. 1 b) was analyzed by genomic Southern blot or by
PCR, we could not identify any cDHC
/
among morphologically normal fetuses. Instead, we found much smaller
decidua that contained completely resorbed embryos. This observation indicated that cDHC
/
embryos seemed to
develop until the blastocyst stage and were able to hatch
and be implanted, but they died before 8.5 dpc. Therefore,
we genotyped blastocysts after heterozygote interbreedings. As shown in Fig. 1 c, cDHC
/
blastocysts were identified, and they appeared to be indistinguishable from control
(cDHC+/
or cDHC+/+) blastocysts at the light microscopic level (Fig. 2, a and b). To check for the presence of
cDHC mRNA, we performed reverse transcriptase PCR
using total RNA from homozygote blastocyst cultures and
control blastocyst cultures as templates. Contrary to our
expectations, we found PCR products from both homozygotes and controls. Hence, we conclude that detectable
amounts of mRNA of maternal origin still remain even at
this developmental stage.
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Growth Abnormalities of cDHC/
Cells
To assess possible defects in cDHC/
cells, we cultured
blastocysts and examined them by light microscopy. On
the third day of culture, both mutant and control blastocysts had completed adhesion onto the surface of coverslips and produced trophoblast giant cell outgrowths (Fig.
2, c and d). However, the ICM appeared smaller in size in
most of the mutant blastocysts compared with control
blastocysts. After longer periods of blastocyst culture, we
found cDHC
/
cells incapable of growing in vitro (Fig. 2,
f and h). After 9 d in culture (6 d after attachment onto the
substrate surface) (Fig. 2, g and h), all of the cDHC
/
blastocysts had failed to grow, and most of the trophoblast
cells had died while control cells were proliferating vigorously (Fig. 2 g). To examine if there was any mitotic phenotype, we took photographs of four additional cultured
cDHC
/
blastocysts every day. However, we were not able
to find any mitotic figures nor an increase in cell number.
The Golgi Complex in cDHC/
Cells Were Highly
Fragmented and Distributed throughout the Cytoplasm
When we examined the distribution of the Golgi complex,
significant changes were found in cDHC/
cells compared with control cells. In cDHC
/
cells, the Golgi complex stained either by antisera against GM130 (cis-Golgi
marker) or by CTR433 monoclonal antibody (medial-Golgi marker) appeared to be highly vesiculated and distributed throughout the cytoplasm (Fig. 3 b), which was in
marked contrast to the case for the control cells, in which
it appeared as large flattened cisternae located around the
nucleus (Fig. 3 a). To further investigate the morphological changes in cDHC
/
cells, we examined control and
cDHC
/
cells by electron microscopy. The rough endoplasmic reticulum, mitochondria, and the nuclei of cDHC
/
cells were indistinguishable from those of control (cDHC+/
or cDHC+/+) cells (Fig. 3, c and d). However, a gross
change in the Golgi complex was observed in cDHC
/
cells (Fig. 3 d). First, the Golgi complex (arrows) was not
located near the nucleus as observed in the control cells
but distributed throughout the cytoplasm as observed by
confocal microscopy. Second, the Golgi complex was
much smaller in size, but it consisted of stacks of several
small flattened cisternae (Fig. 3 e) similar to the Golgi
complex observed in control cells.
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The Distributions of Endosomes/Lysosomes
and the Dynactin Components Were Also Altered in
cDHC/
Cells
When endosomes and lysosomes were labeled by uptake
of extracellular FITC-labeled dextran, they were found to
be distributed around the nuclei in control cells (Fig. 4, a
and c). However, in cDHC/
cells, they were rather uniformly distributed or often remained close to the cell periphery (Fig. 4, b and d, arrows). To quantify the distribution of these organelles, the intensity of FITC-dextran was
measured by confocal microscopy. The ratio of the FITC
intensity in the inner one third against the intensity in the
outer two thirds of the cytoplasm (inner/outer) was found
to be significantly lower in the cDHC
/
cells (0.81 ± 0.31 [mean ± SD]; n = 11) than in the control cells (2.96 ± 1.48; n = 11) (P < 0.001; Student's t test). We obtained similar results when lysosomes were labeled by anti-LAMP2 antibody (Fig. 4, e and f). Moreover FITC-dextran uptake was reduced in cDHC
/
cells. When we
quantified the intensity of endocytosed FITC-dextran by
cDHC
/
cells and control cells cultured on the same coverslip, there was a statistically significant difference in intensity between cDHC
/
cells (12.7 ± 6.84 arbitrary
units; n = 10) and control cells (20.0 ± 6.55 arbitrary units;
n = 21) (P < 0.01; Student's t test). In contrast, the distributions of the endoplasmic reticulum network (Fig. 4, g
and h) and mitochondria (data not shown) in cDHC
/
cells were not significantly different from those in the control cells. The dynactin complex, a protein complex known
to bind cytoplasmic dyneins, is considered to serve as a
linker between membranous organelles and cytoplasmic
dyneins (Schroer, 1994
). To examine their distribution in
the absence of cDHC, we used monoclonal antibodies
against Arp1(45A) and p150Glued (150B) to stain cDHC
/
cells and control cells (Fig. 4, i-l). While both dynactin
components are localized to centrosomally dominant
punctate structures in control cells as described previously
(Gill et al., 1991
) (Fig. 4, i and k), they are localized to
punctate structures distributed throughout the cytoplasm
in cDHC
/
cells, although some centrosomal staining still
remained (Fig. 4, j and l). To assess the amount of cytoplasmic dynein heavy chain, immunostaining using antibodies that specifically react with cDHC was performed
(Vaisberg et al., 1996
). Though control cultured blastocysts showed prominent staining as described in a previous
paper (Vaisberg et al., 1996
), marked reduction in the
level of staining was observed in cDHC
/
cultured blastocysts (Fig. 4, m and n).
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Association of the Golgi Fragments and Endosomes/Lysosomes with Microtubules
Next we examined the spatial relationship between the
Golgi fragments, lysosomes, and microtubules by double
or triple immunofluorescence analysis of cDHC/
cells
(Fig. 5, a-f). The Golgi "fragments" were located quite close to microtubules (Fig. 5, a and b), which was clearly
shown in the cytoplasmic region below the nucleus, where
a relatively small number of microtubules were observed
(Fig. 5 c). To demonstrate the association of the Golgi
fragments with microtubules more clearly, we performed
two additional experiments. First, cDHC
/
cells were incubated with the microtubule-stabilizing agent taxol. Taxol
is known to have the unique ability to promote the formation of discrete microtubule bundles in cells. In the presence
of taxol, microtubule bundles were formed in cDHC
/
cells (Fig. 6 a) as well as in cDHC+/
and cDHC+/+ cells
(data not shown). The Golgi fragments (arrows) in
cDHC
/
cells were found to be associated with these bundles (Fig. 6 a). In the cellular region where microtubule
bundles were not found, the density of microtubules was
sometimes so low that each microtubule filament could
easily be distinguished. In this region the Golgi fragments
(arrows) were also associated with these microtubules (Fig. 6 b), even after stronger fixation protocol (2% paraformaldehyde plus 0.2% glutaraldehyde). These Golgi
fragments were not associated with the ER fragments and
ER protrusions (Fig. 6 c, arrowheads). Second, we examined cDHC
/
cells under electron microscope at higher
magnification (Fig. 3 e). We found microtubules (arrowheads) frequently associated with the small Golgi stacks
(arrows). Lysosomes were also found to be located close
to microtubules (Fig. 5 d). Since Golgi fragments and lysosomes were similarly distributed throughout the cytoplasm and were both associated with microtubules, and since
previous studies have shown the active transport of materials between the TGN and endosome/lysosomes (Goda
and Pfeffer, 1989
; Griffiths, 1989
), we would like to determine whether the Golgi fragments and endosomes/lysosomes are located in close proximity or not. We compared
them with respect to their distribution and found that the
distributions of the Golgi fragments and endosomes/lysosomes were clearly different (Fig. 5, e and f).
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Discussion |
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We examined the role of cDHC in the formation and distribution of organelles for the first time by gene targeting,
and the results were surprising in that only one species
among various DHCs (Tanaka et al., 1995; Vaisberg et al.,
1996
) is essential for the movement of various organelles
in the centrosomal orientation and in cell proliferation.
Regarding cell proliferation, our results are in agreement
with those of Drosophila cDHC mutant analysis (Gepner
et al., 1996
), suggesting that cDHC is essential in higher eukaryotes, while in yeast (Eshel et al., 1993
; Li et al.,
1993
) and fungi (Plamann et al., 1994
; Xiang et al., 1994
), it
is dispensable.
It is intriguing that, without cDHC, the Golgi complex
breaks apart into numerous fragments and disperses throughout the cytoplasm. It seems that, in the steady state, the
Golgi complex is in dynamic equilibrium between the anterograde force mediated by kinesin superfamily motors
and the retrograde force mediated by cytoplasmic dyneins
rather than simply being associated with microtubules near their minus end (or centrosomes). On reduction in
the concentration of cDHC, this equilibrium favors the anterograde force, causing the Golgi complex to be fragmented and dispersed throughout the cytoplasm. Fragmentation of the Golgi complex is easily recognized during mitosis or upon treatment of cells with a microtubule-depolymerizing agent. Even in the fragmented state,
the Golgi complex can serve its function (Cole et al., 1996)
and maintain its polarity (Shima et al., 1997
). Therefore,
fragmentation caused by a reduction in the concentration
of cDHC likely reflects a physiological phenomenon rather
than being simply caused by a mechanical force.
To date, fragmentation of the Golgi during mitosis has
been considered to result from a lack of cytoplasmic microtubules due to microtubule depolymerization (Wehland et al., 1983; Warren, 1993
). Since the microtubule network was not substantially affected in cDHC
/
cells in
this study, we consider Golgi fragmentation to have resulted from the loss of activity of cytoplasmic dynein. Since cDHC is thought to be important in mitosis and is localized to the mitotic spindle and kinetochores (Pfarr et al.,
1990
; Steuer et al., 1990
), translocation of cDHC from the
cytoplasm to the nucleus might occur before the formation
of the mitotic spindle, which may cause a reduction in the
level of cDHC activity in the cytoplasm and thus lead to
fragmentation of the Golgi complex. Alternatively, the inactivation of cDHC in the cytoplasm may occur by some
unknown mechanism such as phosphorylation/dephosphorylation and thus lead to the same results.
Previous studies have shown that injection of anti-DHC2 leads to dispersion of the Golgi complex (Vaisberg
et al., 1996). As suggested by depletion studies indicated
by this paper and the paper of Vaisberg et al., cDHC and
DHC2 appear to have similar functions in maintenance of
the Golgi complex. However, as shown in the study of
Vaisberg et al., the localization of DHC1 (cDHC in this
paper) and DHC2 is clearly different. Recently, Vaisberg et al. indicated that DHC2 is localized to the ER-Golgi intermediate compartment (Vaisberg, E.A., P.M. Grissom,
S.R. Gill, T.A. Schroer, and J.R. McIntosh. 1996. Mol.
Biol. Cell. 7:403a). Since the intermediate compartment
and Golgi complex actively exchange materials and are
known to have similar morphology and distributions under various conditions (e.g., in the presence of brefeldin A
or nocodazole, under temperature change) (Lippincott-Schwartz et al., 1990
; Cole et al., 1996
), it is conceivable
that, under physiological conditions, the two species of dynein heavy chains are localized to different compartments
that reside somewhere between the intermediate compartment and Golgi complex, and that depletion of either molecule leads to a similar phenotype (i.e., dispersion of the
Golgi complex). This can explain the results of recent dynamitin overexpression studies (Burkhardt et al., 1997
;
Presley et al., 1997
) because its overexpression is likely to
affect the interactions between both species of cytoplasmic
dyneins and their target organelles.
It is also surprising that, even in cDHC/
cells, the
Golgi fragments and endosomes/lysosomes are attached to
microtubules. This association raises the possibility that other
molecules such as the dynactin complex (Hirokawa, 1993
;
Holzbaur and Vallee, 1994
; Schroer, 1994
; Hirokawa
1998
) or other anterograde/retrograde motors (Hirokawa,
1996
) might mediate attachment of these organelles to microtubules. In particular, dynactin, known as a binding
protein of cytoplasmic dynein (Gill et al., 1991
; Paschal et al.,
1993
), has intriguing characteristics because one of its
complexes, p150Glued, has its own microtubule-binding domain (Waterman-Storer et al., 1993
) and cytoplasmic dynein depleted of p150Glued is inactive in promoting organelle movement in vitro (Gill et al., 1991
). Therefore, it
is quite likely that in the absence of cDHC, the Golgi complex and endosomes/lysosomes are attached via p150Glued
to microtubules. Previous analyses of the amino acid sequence of p150Glued revealed a motif that is homologous to
a microtubule-binding motif in the endosome-microtubule linker protein CLIP170 (Pierre et al., 1992
). Thus, for
endosomes/lysosomes, CLIP170, together with p150Glued,
may link these organelles to microtubules through this
common microtubule-binding domain.
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Footnotes |
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Received for publication 2 September 1997 and in revised form 5 February 1998.
1. Abbreviations in this paper: cDHC, cytoplasmic dynein heavy chain; dpc, days postcoitum; ES, embryonic stem; ICM, inner cell mass; RT, room temperature.We would like to thank Dr. T. Noda (Cancer Institute, Tokyo, Japan) for his advice and generous gifts of various plasmids. We would also like to thank: Drs. M. Fujiwara (University of Tokyo School of Medicine), S. Matsuyama, and S. Tanaka (University of Tokyo School of Agricultural Sciences) for their advice on animal care; E.A. Vaisberg and J.R. McIntosh (University of Colorado Department of Molecular, Cellular, and Developmental Biology, Boulder, CO), M. Bornens (Institut Curie, Paris, France), N. Nakamura, G. Warren (Imperial Cancer Research Fund Cell Biology Laboratory, London, UK), and T. Schroer (Johns Hopkins University, Baltimore, MD) for their generous gifts of antibodies; and R. Sato-Harada, Y. Okada, Y. Yonekawa, Z. Zhang, H. Sato, C. Zhao, and other members of Hirokawa's lab for help and discussions. The anti-LAMP-2 antibody ABL-93 was obtained from the Developmental Studies Hybridoma Bank maintained by the Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine and the Department of Biological Sciences, University of Iowa (Iowa City, IA) under contract NO1-HD-2-3144 from the National Institue of Child Health and Human Development.
This work was supported by a Special Grant-in-Aid for Center of Excellence from the Japan Ministry of Education, Science and Culture to N. Hirokawa.
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