Host-Parasite Interactions Section, Laboratory of Intracellular Parasites, NIAID, NIH, Rocky Mountain Laboratories, Hamilton, MT 59840, USA
* Author for correspondence (e-mail: ted_hackstadt{at}nih.gov)
Accepted 3 June 2003
![]() |
Summary |
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Key words: Chlamydia, Cytoskeleton, Microtubules, Dynein, Dynactin
![]() |
Introduction |
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Within the first few hours postinfection, endocytosed EBs are trafficked to
the peri-Golgi region of the host cell where the inclusions remain for the
duration of chlamydial intracellular development
(Hackstadt, 1999). Recruitment
of the nascent inclusion to the peri-Golgi region appears to require an active
process on the part of the chlamydiae, given that inhibition of chlamydial
transcription or translation blocks their translocation
(Scidmore et al., 1996
).
C. trachomatis inclusions aggregate at a perinuclear or peri-Golgi
location that corresponds to the microtubule-organizing center (MTOC) within 6
hours postinfection (Clausen et al.,
1997
; Hackstadt et al.,
1996
). The minus-end-directed microtubule motor, dynein, has been
implicated in this process as dynein colocalizes with chlamydial early
inclusions and Na3VO4, a general inhibitor of tyrosine
kinases that also inhibits dynein, detrimentally affects chlamydial
development when cells are treated during the course of infection
(Clausen et al., 1997
). Thus,
like many cellular organelles, the chlamydial inclusion appears to be
trafficked intracellularly via interactions with the microtubular network.
Microtubules are polarized structures with a minus end anchored at the MTOC
and a plus end directed towards the periphery of the cell. Microtubules serve
as a scaffold for the transport and sorting of various cellular cargoes and
are involved in such diverse cellular functions as chromosome segregation,
organelle transport, and regulation of anterograde and retrograde trafficking
through the Golgi complex. Transport along microtubules is mediated by
ATP-dependent, microtubule-associated motor proteins. Kinesin superfamily
proteins comprise the major plus-end-directed motors, whereas dynein
superfamily proteins are the minus-end-directed motors. An accessory protein
complex called dynactin is thought to be required for cargo binding to the
dynein motor complex. Dynactin is a large multisubunit complex consisting of
p150(Glued), p62, p50 dynamitin, actin-related protein 1
(Arp1) and actin-capping protein
(Hirokawa, 1998;
Waterman-Storer et al., 1997
).
Overexpression of p50 dynamitin inhibits the ability of dynein to interact
with its cargo and results in the disruption of minus-end movement along
microtubules (Burkhardt et al.,
1997
; Valetti et al.,
1999
).
Many bacterial and viral pathogens exploit the microtubular network to
access specific sites within the host cell
(Alonso et al., 2001;
Kim et al., 2001
;
Ploubidou et al., 2000
;
Ye et al., 2000
). In this
study, we show that C. trachomatis modifies the inclusion membrane to
recruit dynein and selected components of the dynactin complex for migration
along microtubules in a fashion similar to host vesicular trafficking.
However, chlamydial recruitment of dynein appears to be mechanistically
distinct as migration to the MTOC is not disrupted by overexpression of the
p50 dynactin subunit of dynamitin. These findings suggest that chlamydiae
circumvent the necessity for an intact dynein-dynactin motor complex in a
unique manner in which a chlamydial protein supplants a requirement for at
least the dynamitin component of dynactin.
![]() |
Materials and Methods |
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For live imaging, C. trachomatis L2 EBs were intrinsically labeled
with the fluorescent dye 5-(and
-6)-([(4-chloromethyl)benzoyl]-amino)tetramethylrhodamine (CMTMR) as
previously described (Boleti et al.,
2000; Carabeo et al.,
2002
). Briefly, HeLa cells were infected with serovar L2 EBs at a
multiplicity of infection (MOI) of 1 and incubated for an additional 24 hours
with 25 µg of CMTMR cell tracker/ml added to the culture medium at 12 hours
postinfection. EBs were then harvested by Renografin density gradient
centrifugation. The infectivity of CMTMR-treated EBs was evaluated by
inclusion-forming assay and was found to be unaffected by the labeling
procedure (Carabeo et al.,
2002
).
For microcopy studies Cos-7 cells were seeded on 25 mm #1 borosilicate coverslips at a density of 3x105 cells per coverslip and cultivated overnight at 37°C in RPMI medium supplemented with 10% fetal bovine serum (FBS) and 10 µg/ml gentamicin (Invitrogen, Carlsbad, CA) under 5% CO2.
All infections were carried out similarly unless otherwise noted. Cells
were incubated with C. trachomatis EBs at an MOI of 50 in Hanks
Balanced Salts Solution (HBSS) (Invitrogen, Carlsbad, CA) for 10 minutes,
after which the inoculum was removed and the coverslips were washed twice with
HBSS with 100 µg/ml heparin (Pharmacia, Peapack, NJ), and once with HBSS
without heparin. The HBSS was then replaced with RPMI media containing 10% FBS
and 10 µg/ml gentamicin. Infections were allowed to proceed for appropriate
times.
Transfections and microinjections
Cos-7 cells were seeded on 12 mm glass coverslips in 24-well plates to
obtain a monolayer of approximately 50% confluency. Transfections of plasmid
constructs were performed using Lipofectamine 2000 (Invitrogen) according to
the manufacturer's directions. The transfection mixture was prepared as
follows: 0.8 µg of DNA was diluted in 50 µl of Optimem serum-free media
(Invitrogen) and added to a solution of 50 µl of Optimem with 3 µl
lippofectamine 2000. After a 20 minute incubation at room temperature, the
complexes were added to one well of a 24-well plate containing 500 µl of
RPMI with 10% FBS. Expression vectors used were enhanced green fluorescent
protein (EGFP)-C1, EYFP-Golgi (BD Biosciences, San Jose, CA) and GFP-dynamitin
(Valetti et al., 1999).
Expression from the transfected vectors was allowed to proceed for 24 to 30
hours before experimentation.
Microinjection of antibodies was performed using an automated
microinjection system as described previously
(Heinzen and Hackstadt, 1997).
Microinjected antibodies used in these experiments were: mouse mAb to dynein
intermediate chain dic74.1 (Covance); mouse mAb to dynein intermediate chain
dic70.1 (Abcam, Cambridge, UK); mouse mAb to kinesin (Covance); and mouse mAb
to Rickettsia rickettsii (Heinzen
et al., 1999
). All antibodies were purified using protein G
sepharose columns (Pharmacia, Peapack, NJ) according to the manufacturers'
directions, followed by buffer exchange and concentration using microspin
concentrators (Millipore, Bedford, MA). The antibodies were concentrated to 10
µg/ml in microinjection buffer (48 mM K2HPO4, 14
mMNaH2PO4, 4.5 mM KH2PO4, pH 7.2).
Before the injections, cells on coverslips were removed from 24-well plates
and placed in 100 cm diameter Petri dishes containing 10 ml of fresh RPMI plus
10% FBS. Microinjection of the cytoplasm of cells was done with a
Micromanipulator 5171 and a Transjector 5426 plus (Eppendorf, Hamburg,
Germany). Femtotips (Eppendorf) were backfilled with 3 µl samples by using
Microloaders (Eppendorf). Injections were monitored by epifluorescence
illumination on a Nikon TE 300 inverted microscope equipped with a Polychrome
I polychromatic illumination system (Applied Scientific Instrumentation,
Eugene, OR.). Oregon green dextran was used for monitoring injections
(Molecular Probes, Eugene, OR) and was added to the antibodies at a final
concentration of 0.5 mg/ml. All injections were conducted at room temperature
and the delivery pressure and injection duration for most experiments was 1.4
lb/in2 and 0.5 seconds, respectively. This resulted in an estimated
delivery volume of approximately 0.1 pl
(Minaschek et al., 1989
).
Following injection, cells were washed once with RPMI plus 10% FBS, and fresh
medium was added. Ten to fifteen minutes after injection, the coverslips were
infected with C. trachomatis L2 or used in endocytic trafficking
experiments. The injected antibodies were detected using AlexaFluor
488-conjugated goat anti-mouse IgG secondary antibodies.
Endocytic markers
For endocytosis assays, Cos-7 cells transfected with appropriate constructs
or microinjected with various antibodies were incubated with 25 µg/ml
AlexaFluor 594-transferrin (Tf) or 5 µg/ml AlexaFluor 594-cholera toxin B
(Molecular Probes) in Optimem serum-free media (Invitrogen) for 5 hours at
37°C. Cells were quickly rinsed in HBSS before fixing with freshly
prepared 4% paraformaldehyde in PBS for 10 minutes at room temperature.
Immunofluorescence staining
For fluorescent antibody staining, cells were fixed with cold methanol for
10 minutes. The cells were washed three times with PBS. Antibodies used in
these experiments were mouse mAb to dynein intermediate chain dic70.1 diluted
1:100 (Abcam), mouse mAb to p150(Glued) 2.5 µg/ml (BD
Biosciences, San Jose, CA) and mouse mAb to ß-tubulin diluted 1:300
(Sigma). Primary antibodies were incubated on cells for 3 hours followed by
three washes with PBS. To visualize the primary antibodies the cells were
incubated with AlexaFluor 488-conjugated goat anti-mouse IgG 4 µg/ml. To
simultaneously visualize chlamydiae, cells were also stained using a rabbit
polyclonal Ab to C. trachomatis serovar L2 at a dilution of 1:1000.
This antibody was detected using AlexaFluor 594-conjugated goat anti-rabbit
IgG secondary antibody 4 µg/ml. For cytochalasin D experiments, cells were
fixed with 4% paraformaldehyde for 10 minutes and permeabilized with 0.05%
triton-X 100 and stained using FITC-phalloidin 10 U/ml (Molecular Probes). All
fluorescence images were obtained with a Zeiss Axiovert LSM 510 confocal
microscope. Projections were constructed using the ImageJ image software
(written by Wayne Rasband at the U.S. National Institutes of Health and
available by anonymous FTP from zippy.nimh.nih.gov).
Quantification of fluorescent microscopy
Quantification of peri-Golgi fluorescence was conducted using ImageJ
software. Regions of interest were chosen that represented the peri-Golgi
region in individual cells from the micrographs. The fluorescence intensity of
this region was represented as a percentage of the total fluorescence signal
of the entire cell.
Live-cell confocal microscopy
For live-cell imaging, cells were subcultured into 35 mm glass bottom Petri
dishes (Mat Tek Corp, Ashland, MA). The dishes were placed in a
temperature-controlled chamber and maintained at 37°C throughout the
experiment. To observe normal trafficking of host cell vesicles, cells were
labeled with fluorescent
N-[7-(4-nitrobenzo-2-oxa-1,3-diazole)]-6-aminocaproyl-D-erythro-sphingosine
(C6-NBD-Cer) (Molecular Probes). C6-NBD-Cer was
complexed with 0.034% defatted bovine serum albumin (dfBSA) in HBSS, as
described previously (Pagano and Martin,
1988), to yield complexes equal to a concentration of 5 µM in
both dfBSA and C6-NBD-Cer. Cells were incubated with the
dfBSA/NBD-Cer complex at 37°C for 15 minutes, washed with HBSS and
incubated for 2 hours in HBSS with 0.34% dfBSA to back-exchange excess probe
from the plasma membrane. These cells were simultaneously infected with
CMTMR-labeled L2 EBs. The infection was allowed to proceed for 2 hours before
viewing. Images were acquired on a Zeiss Axiovert LSM 510 confocal microscope
and a 64x, 1.4 NA lens with a computer-controlled 488 nm argon laser to
excite NBD or a 564 nm krypton laser to excite CMTMR. Images were collected at
15-second intervals using the Zeiss LSM 510 software.
![]() |
Results |
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|
To determine whether chlamydial aggregation at the MTOC is dependent on microtubules, the microtubule network was disrupted with nocodazole. Cos-7 cells were infected with L2 EBs for 1 hour to allow for entry before treatment with nocodazole. Five hours after infection cells were observed by indirect immunofluorescence for both tubulin and EBs. The nocodazole treatment completely disrupted the microtubule network and the chlamydiae remained dispersed throughout the cytoplasm of infected cells (Fig. 2A,B). This effect was specific to microtubules; disrupting the Golgi apparatus with brefeldin A (Fig. 2C,D) or depolymerizing the actin cytoskeleton with cytochalasin D (Fig. 2E,F) had no effect on aggregation.
|
Chlamydial migration is delayed due to a requirement for chlamydial
protein synthesis
Approximately 4-5 hours is required for most inclusions to aggregate at the
MTOC. The speed of host vesicle movement along microtubules is about 0.5
µm/second (Garcia-Mata et al.,
1999
; Schroer,
2000
; Toomre et al.,
1999
); thus, the time it takes for perinuclear delivery of
chlamydiae could be due to either slow movement along microtubules or a delay
in the initiation of migration. Chlamydial modification of the inclusion
membrane occurs very soon after infection
(Hackstadt, 1999
). A
requirement for chlamydial protein synthesis suggests that modification of the
nascent inclusion by chlamydiae is necessary to initiate migration along
microtubules and could explain this observed lag in perinuclear delivery. To
confirm a role for chlamydial protein synthesis in Cos-7 cells, the cells were
infected in the presence of chloramphenicol (50 µg/ml), an inhibitor of
prokaryotic translation, or rifampicin (150 µg/ml), a specific inhibitor of
prokaryotic transcription. Treatment of infected cells with chloramphenicol
completely inhibited perinuclear localization of chlamydiae
(Fig. 3). Treatment with
rifampicin also inhibited perinuclear aggregation
(Fig. 3). Chlamydial
modification of the inclusion membrane is therefore necessary for aggregation
and migration.
|
Rapid movement along microtubules
Although the necessity for chlamydial protein synthesis accounts for the
observed lag in perinuclear delivery it does not indicate if
Chlamydia are capable of rapid travel on microtubules. To address
this question, we synchronized chlamydial migration using nocodazole to
inhibit migration and allowed the chlamydiae time to modify the inclusion.
This was followed by a washout of nocodazole to re-initiate migration. Cos-7
cells were infected with L2 for 30 minutes at 4°C before the addition of
nocodazole. Infected cells were incubated in the presence of nocodazole for 4
hours, after which time the medium was replaced with fresh media without drug.
The cells were subsequently fixed and stained for both microtubules and C.
trachomatis after either 15 minutes or 1 hour additional incubation. By
15 minutes the microtubule network was partially restored and many cells had
obvious MTOCs and some microtubules. In these cells, many of the inclusions
were already aggregated at the reorganizing MTOC
(Fig. 4). By 1 hour after
nocodazole washout, the microtubules were more organized and the majority of
the chlamydiae were aggregated at the MTOC. Occasionally, multiple MTOCs
reformed in a single cell, some at sites distal to the nucleus. The chlamydiae
could be found aggregated at each of these sites
(Fig. 4).
|
To further characterize chlamydial migration we compared the trafficking of
host vesicles with that of chlamydiae. Cos-7 cells were infected with C.
trachomatis L2 EBs prelabeled with the fluorescent dye CMTMR to enable
simultaneous visualization of chlamydial migration and host vesicular
trafficking. The infected cells were then incubated with the fluorescent
lipid, C6-NBD-ceramide. C6-NBD-ceramide is taken up by
cells and converted into sphingomyelin, which brightly stains the Golgi
apparatus, and Golgi-derived vesicles. Sphingomyelin-containing vesicles can
then be observed trafficking bidirectionally between the Golgi and plasma
membrane, a process that is microtubule dependent
(Koval and Pagano, 1989).
Viable infected cells were observed for 3 hours postinfection. Time-lapse
confocal microscopy revealed that chlamydial migration is indeed very
asynchronous, with only a few chlamydiae migrating per cell during
visualization (about 30 minutes). Most of the chlamydiae were either already
at the MTOC or still in the cell periphery. The migrating CMTMR-labeled
chlamydiae displayed patterns of movement similar to the NBD-stained vesicles
(Fig. 5). In general, the
movement was similar in speed, averaging 0.1±0.01 µm/second
(n=3) for C. trachomatis and 0.11±0.01 µm/second
(n=3) for vesicles containing C6-NBD ceramide. This
movement was characterized by bursts of fast motion followed by stalls with
occasional direction changes. Fig.
5 shows time-lapse panels of a CMTMR-labeled EB traveling parallel
to and similar in velocity to a C6-NBD-ceramide-labeled vesicle.
These data suggest that inclusions, once modified by the chlamydiae, are
capable of rapid migration at rates comparable to those of host-cell
vesicles.
|
Chlamydial inclusions colocalize with dynein
Dynein has previously been shown to be associated with early chlamydial
inclusions (Clausen et al.,
1997). To confirm this in Cos-7 cells, cultures were infected with
C. trachomatis L2 and fixed and stained with the dic70.1 monoclonal
antibody at 5 hours and 24 hours postinfection. Confocal micrographs revealed
that dynein localized along microtubules and to the nascent inclusions in the
perinuclear region (Fig. 6).
When the migration of the chlamydial nascent inclusions was inhibited by
nocodazole treatment, dynein was still recruited to the chlamydial inclusion
(Fig. 6). This recruitment
could be inhibited by treatment with chloramphenicol, indicating that
chlamydial protein synthesis was necessary for this interaction (data not
shown). Colocalization of dynein with the chlamydial inclusion was not
restricted to early time-points as dynein staining of the mature chlamydial
inclusions remained apparent at 24 hours postinfection
(Fig. 6). At this time the
multiple individual inclusions had fused and the inclusion had taken on its
characteristic spherical structure with a spacious, fluid-filled center rimmed
by chlamydial RBs.
|
Chlamydial microtubule migration is independent of the dynactin
complex
To investigate chlamydial interactions with the dynein-dynactin system, we
transiently transfected Cos-7 cells with a plasmid encoding GFP-dynamitin.
Overexpression of dynamitin causes dissociation of the dynactin complex, thus
decoupling dynein-binding and cargo-anchoring functions and effectively acting
as a dominant negative for dynactin activity
(Echeverri et al., 1996;
Wittmann and Hyman, 1999
).
Cells overexpressing dynamitin were infected with C. trachomatis L2
for 5 hours before fixing and staining for chlamydiae and microtubules.
Dynamitin overexpression had no effect on chlamydial aggregation at the MTOC
(Fig. 7). In many
dynamitin-transfected cells the MTOC was less organized than in untransfected
cells, but in each case the chlamydiae were aggregated at a perinuclear
site.
|
To verify that the GFP-dynamitin construct was indeed acting as a dominant
negative for the dynactin-dynein pathway, we tested the effect of
overexpression on trafficking of two model proteins, transferrin (Tf) and
cholera toxin subunit B (CTX), trafficked by microtubule-dependent pathways.
Tf binds to the Tf receptor on the cell surface and is delivered to late
endosomes in the peri-Golgi region. This trafficking is inhibited by
overexpression of dynamitin (Valetti et
al., 1999). Alexa-594-conjugated Tf was incubated with either GFP
or GFP-dynamitin-transfected cells for 5 hours. The cells were then fixed and
observed by confocal microscopy. Cells expressing GFP-dynamitin had little to
no Tf signal in the perinuclear region, and most of the signal was distributed
throughout the periphery. This is in contrast to untransfected cells, which
had significant signal in the perinuclear region
(Fig. 7). The Tf signal in
cells expressing GFP only were indistinguishable from untransfected cells
(data not shown).
The trafficking of CTX to the Golgi involves binding to the lipid GM1, a
ganglioside on the cell surface, followed by internalization and delivery by
vesicle trafficking directly to the Golgi without interacting with
transferrin-containing late endosomes
(Nichols et al., 2001;
Nichols, 2002
). This pathway
is distinct from that of transferrin but dynein-dynactin dependent. We
investigated the effect of dynamitin overexpression on the delivery of CTX to
the Golgi. Cos-7 cells were transfected with plasmids expressing either GFP or
GFP-dynamitin for 24 hours before incubating the cells for 5 hours with
Alexa-594 CTX. Cells expressing GFP-dynamitin did not accumulate CTX in the
perinuclear region, whereas untransfected cells on the same coverslip had
substantial CTX signal in a distinct perinuclear pattern
(Fig. 7). The localization of
CTX in GFP-expressing cells was identical to neighboring untransfected cells
(data not shown).
The differential effect of GFP-dynamitin overexpression on the intracellular trafficking of Tf and CTX compared with C. trachomatis EBs was quantified by measuring the percent fluorescent signal in the perinuclear region compared with the total signal in the cell. Cells transfected with GFP-dynamitin were subsequently incubated with Alexa 594-Tf or Alexa 594-CTX, or infected with C. trachomatis L2 for 5 hours. Cells incubated with Tf or CTX were fixed and observed directly and cells infected with C. trachomatis were fixed and the bacteria were visualized by indirect immunofluorescence. Dual color micrographs were taken of both transfected cells (bright GFP signal) and untransfected cells (no GFP signal) on the same coverslips. In untransfected cells, about 65% of the fluorescence signal for Tf, CTX and chlamydial nascent inclusions was perinuclear by 5 hours after treatment (68%±3, n=14; 62%±3, n=16; and 67%±5, n=34, respectively). By contrast, cells that expressed GFP-dynamitin had very limited amounts of Tf and CTX signal in the perinuclear region (10%±2, n=15 for Tf and 7%±2; n=13 for CTX). However, chlamydial inclusions localized no differently in transfected versus untransfected cells (71%±3, n=31 and 67%±5, n=34, respectively). These data suggest that chlamydial perinuclear migration may be mechanistically distinct from currently described systems.
Chlamydial perinuclear aggregation is dependent on dynein
Because chlamydial aggregation was unaffected by dynamitin overexpression,
we asked whether chlamydiae require dynein for their minus-end-directed
microtubule motor activity. For these experiments we microinjected monoclonal
antibodies against the dynein intermediate chain to determine any effect on
chlamydial intracellular trafficking. The monoclonal antibody dic74.1 was
microinjected into Cos-7 cells followed by infection with C.
trachomatis L2. The cells were incubated for 5 hours to allow the
chlamydiae to initiate development and translocate before fixing and staining
for chlamydiae and the microinjected antibody. Cells that had been
microinjected showed a chlamydial aggregation phenotype that was
indistinguishable from nocodazole-treated cells, in that there was no
perinuclear aggregation, and the nascent inclusions were scattered throughout
the cytoplasm (Fig. 8). This
was, to some extent, dependent on antibody concentration as cells that stained
weakly for microinjected antibody showed an intermediate phenotype with many
aggregated and unaggregated nascent inclusions. We confirmed the requirement
for dynein using a second antibody dic70.1 that also recognizes the dynein
intermediate chain. Microinjection of the dic70.1 antibody showed an identical
phenotype (data not shown). To confirm the specificity of the antibody, two
irrelevant antibodies, against either R. rickettsii or the major
plus-end-directed microtubule motor, kinesin, were also microinjected. Neither
of these antibodies had any effect on chlamydial perinuclear aggregation
(Fig. 8). Taken together, these
data show that dynein is recruited to the chlamydial inclusion and is
necessary for migration and that this recruitment is dependent on expression
of a chlamydial protein or proteins.
|
The p150(Glued) subunit of dynactin colocalizes
with the chlamydial nascent inclusions
Because overexpression of the p50 dynamitin subunit does not affect
chlamydial intracellular aggregation but dynein is recruited and required for
this movement, we asked whether p150(Glued), the largest
subunit of the dynactin complex, colocalized to the chlamydial inclusion.
Indirect immunofluorescence showed that the p150(Glued)
subunit of dynactin is recruited to the chlamydial inclusion
(Fig. 9). Like dynein,
p150(Glued) also colocalized with nascent inclusions in
nocodazole-treated cells (Fig.
9). This recruitment is also dependent on chlamydial protein
synthesis as recruitment is inhibited by chloramphenicol (data not shown).
|
Overexpression of the p50 dynamitin subunit of dynactin disrupts the
interaction between the motor complex, dynein, and the activating and
cargo-binding complex, dynactin (Kamal and
Goldstein, 2002; Karcher et
al., 2002
). Because migration of the nascent inclusion was
unaffected by overexpression of GFP-dynamitin, we asked whether this
overexpression was able to disrupt the ability of the chlamydial inclusion to
recruit p150(Glued). Cos-7 cells were transfected with
GFP-dynamitin for 24 hours before infection with C. trachomatis. The
infection was allowed to proceed for 5 hours before fixation and staining. The
cells were observed by three-color confocal microscopy for GFP signal, as well
as immunofluorescence staining for chlamydiae and
p150(Glued). The p150(Glued)
colocalized with the chlamydial inclusions in the cells that expressed
GFP-dynamitin in the same fashion as untransfected cells
(Fig. 9). The overexpression of
GFP-dynamitin did not remove p150(Glued) from the
inclusion, suggesting that this interaction is independent of p50
dynamitin.
![]() |
Discussion |
---|
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---|
The exploitation of the host cell microtubule network is not unique to
chlamydiae. Several large DNA viruses including adenovirus, African swine
fever virus, vaccinia virus and herpes viruses use the dynein-dynactin
machinery for intracellular migration
(Alonso et al., 2001;
Dohner et al., 2002
;
Leopold et al., 2000
;
Ploubidou et al., 2000
). At
least one other intracellular bacterium, Orientia tsutsugamushi, also
uses this system for intracellular migration
(Kim et al., 2001
). However,
each of these examples differ from chlamydiae in that their migration is
disrupted by the overexpression of the dynactin subunit p50 dynamitin. Indeed,
overexpression of this dynactin subunit inhibits all reported
dynein-dynactin-dependent processes in host cells. We confirmed that
overexpression of p50 dynamitin inhibits the perinuclear accumulation of
transferrin as well as the delivery of cholera toxin subunit B to the Golgi
apparatus. The overexpression of p50 dynamitin has been reported to inhibit
many diverse host functions that rely on minus-directed motor activity such
as: endocytic vesicle trafficking, the organization of the Golgi complex, the
formation of perinuclear aggresomes, reorientation of the MTOC after injury to
fibroblasts, axonal transport in motor neurons, and the assembly and
organization of the intermediate filament network
(Echeverri et al., 1996
;
Garcia-Mata et al., 1999
;
Helfand et al., 2002
;
LaMonte et al., 2002
;
Palazzo et al., 2001
;
Valetti et al., 1999
).
An intact dynactin complex is believed to be a necessary component in
dynein-mediated minus-end-directed microtubule-mediated transport.
Overexpression of the dynactin subunit p50 dynamitin dissociates dynactin into
two subcomplexes: a dynein-binding subunit containing
p150(Glued) and a cargo-binding subunit that contains Arp1
(Burkhardt et al., 1997).
Unlike most intracellular pathogens, interactions of chlamydial inclusion with
microtubules appear unique in that dynamitin overexpression fails to disrupt
trafficking of endocytosed EBs to the MTOC. Although transport of the nascent
chlamydial inclusion is not sensitive to the overexpression of p50 dynamitin,
the dynactin subunit p150(Glued) is recruited to the
inclusion, suggesting that the dynactin complex may have multiple and
distinguishable functions. Our results suggest that
p150(Glued) may be required for dynein activation or
processivity but that the dynamitin subunits and possibly other subunits
required for cargo binding are dispensable for interaction with C.
trachomatis inclusions. In the model by Hirokawa
(Hirokawa, 1998
), the
chlamydial inclusion may supply the cargo binding activity in the form of
chlamydial proteins (Fig. 10).
This model is supported by the report that Arp1 but not
p150(Glued) is removed from microtubules by p50 dynamitin
overexpression (Vaughan et al.,
1999
), suggesting that the functional interaction of dynein and
p150(Glued) may not require the entire dynactin complex.
An alternative model might be that chlamydial proteins may stabilize the
interactions of the cargo-binding and dynein-binding subunits of dynactin to
the extent that it is no longer susceptible to dissociation by overexpression
of dynamitin. A detailed characterization of the components of dynein-dynactin
complex associated with chlamydial inclusions and the chlamydial proteins
involved will be required to distinguish between these models. In either case,
the novel interactions of the chlamydial inclusion with the dynactin complex
may reveal some interesting biology of the early events in chlamydial
development but may also help elucidate mechanisms by which the dynactin
complex and dynein interact.
|
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Acknowledgments |
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