1 Department of Cell Biology, University of Texas Southwestern Medical Center,
Dallas, Texas 75235-9039, USA
2 Department of Biomedical Engineering, University of Tokyo, 7-3-1 Hongo,
Bunkyo-ku, Tokyo113-8655, Japan
3 Department of Nephrology and Endocrinology, University of Tokyo, 7-3-1 Hongo,
Bunkyo-ku, Tokyo113-8655, Japan
* Author for correspondence (e-mail: anders06{at}utsw.swmed.edu)
Accepted 30 October 2001
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Summary |
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Key words: Caveolae, Signal transduction, Polarization, Endothelium
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Introduction |
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A variety of studies now support the proposal that caveolae
compartmentalize signal transduction
(Anderson, 1998). Caveolae are
enriched in a variety of signal transducing molecules; complex signaling
events that use these molecules have been localized to this domain, and the
caveolin family of lipid binding proteins have been identified as molecules
involved in organizing the activity of various signaling molecules in caveolae
(Smart et al., 1999
). There
remains, however, the question of whether or not caveolae can spatially
organize signal transduction. Immunofluorescence localization of caveolin-1 in
fibroblasts and endothelial cells show that this caveolae marker protein
collects at the margins of cells and in patches that are aligned with
actin-rich stress fibers (Isshiki et al.,
1998
; Rothberg et al.,
1992
). Electron microscopic thin section and enface
(Rothberg et al., 1992
) images
of plasma membranes have confirmed that both invaginated and flat caveolae are
densely packed in specific regions of the membrane, but devoid from other
regions. GFP-tagged caveolin-1 has been used to monitor the distribution of
caveolin-1 in response to different growth conditions and at various stages of
the cell cycle. GFP-caveolin collects at sites of cell-cell contact in contact
inhibited cells (Volonte et al.,
1999
) and concentrates at the cleavage furrow during cytokinesis
(Kogo and Fujimoto, 2000
).
Assuming that GFP caveolin-1 is a reliable marker for caveolae, these studies
suggest that the distribution of caveolae on the cell surface is regulated. It
is not known if resident signaling molecules move with caveolae when they are
induced to relocate.
A signaling event that has been localized to caveolin-rich regions of the
endothelial cell surface is the ATP-stimulated initiation of Ca2+
waves (Isshiki et al., 1998).
Changes in intracellular Ca2+ concentration can be detected with
the confocal microscope using the Ca2+ sensing dye Indo-1. The
initial phase of Ca2+ release from ER was found to occur at the
edge of cells in juxtaposition to caveolin-rich areas of membrane. A wave of
Ca2+ release was then propagated from this site across the cell.
Interestingly, initiation of Ca2+ waves occurred only at a subset
of the caveolin-rich regions, suggesting that not all caveolae in a cell are
equipped to stimulate Ca2+ release from ER. These results suggest
that caveolae spatially organize ATP-induced wave initiation. If so, then
conditions that cause signal competent caveolae to relocate on the cell
surface should shift sites of Ca2+ wave initiation to these
regions. Here we demonstrate that when endothelial cells are exposed to shear
stress the caveolae accumulate in the region of the cell nearest the direction
of media flow (designated the `upstream edge' of the cell). Polarization of
caveolae is dependent on the amount of flow force and the time that it is
applied. Once polarized, G
q, the heterotrimeric G protein
that links ATP receptors to ER Ca2+ release, and sites of
Ca2+ wave initiation both co-localize with caveolae.
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Materials and Methods |
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Cell culture
Primary cultures of endothelial cells (ECs) were derived from the
descending thoracic aorta of a bovine fetus by brief collagenase digestion of
the intimal lining. Cells were grown in M199 supplemented with 20% FBS (v/v),
2 mM L-glutamine, 50 IU/ml of penicillin and 50 µg/ml of streptomycin
(designated, standard medium) and routinely passed by trypsinization in a
0.25% trypsin/1 mM EDTA solution before reaching confluence. Cumulative
population doublings (CPD) of ECs were calculated using a Coulter Counter
(Model ZM system, Coulter Electronics Ltd., Luton, UK). Sub-confluent ECs with
a CPD under 40 were used in each experiment.
Exposing endothelial cells to fluid shear stress
On day zero, 3.0x105 endothelial cells were seeded onto a
60 mm culture dish containing a 25.8x45x0.2 mm glass coverslip
(Matsunami, Osaka, Japan) and grown for 3 days in 5 ml of standard medium. The
coverslip was removed from the dish and placed in a flow-loading chamber
(Yasuhisa Biomechanics, Tokyo, Japan) for each experiment. A
parallel-plate-type flow chamber was used to load a laminar flow on ECs as
previously described (Ando et al.,
1993). Briefly, one side of the chamber was the coverslip
(25.8x45x0.2 mm) on which the cells were cultured. The other side
was a polymethacrylate plate. These two flat surfaces were held approximately
200 µm apart by a silicone rubber gasket. Culture medium maintained at
37°C and 5% CO2 was re-circulated through the chamber using a
silicone tube connected to a reservoir by a roller/tube pump (ATTO Co., Tokyo,
Japan). The amount of shear stress
(dyn/cm2) applied to the
cells was calculated using the formula
=6µQ/a2b, where
µ is the viscosity of the perfusate (0.0094 poise at 37°C), Q is the
volume flow (ml/s), and `a' (0.02 cm) and `b' (1.6 cm) are the cross-sectional
dimensions of the flow path.
Cell wounding
On day zero, 3.0x105 endothelial cells were seeded onto a
60 mm culture dish containing a 25.8x45x0.2 mm coverslip and grown
for 3 days in 5 ml of standard medium. One half of the cell monolayer was
sharply scraped with a sterilized cell scraper (Corning Inc, Corning, NY) and
the scraped cells were removed by washing the culture dish with fresh medium.
The cells were then allowed to grow for the indicated time and processed for
indirect immunofluorescence.
Indo-1 imaging of intracellular [Ca2+]i
Loading of cells with the Ca2+ indicator dye, image acquisition
and image processing was carried out as previously described
(Isshiki et al., 1998).
Indirect immunofluorescence and actin-staining
After the application of shear stress and the recording of
[Ca2+] images, the cells were washed three times with PBS, fixed
for 30 minutes at room temperature with 3% (w/v) paraformaldehyde in PBS and
processed for fluorescence localization of the indicated proteins as
previously described (Isshiki et al.,
1998). To co-localize caveolin-1 with other cellular proteins,
fixed cells were incubated in the presence of a 1/120 dilution of caveolin-1
pAb plus either 4 U/ml Texas Red-X phalloidin (actin), a 1/80 dilution of
vinculin mAb, a 1/50 dilution of
tubulin mAb or a 1/50 dilution of
AP-1/2 mAb. For co-localization of G
q/11, cells were stained
with mAb caveolin-1 (1/20 dilution) and pAb G
q/11 (1/50
dilution) and visualized with Alexa Fluor 488 goat anti-mouse IgG and Alexa
Fluor 568 goat anti-rabbit IgG, respectively.
Quantification of caveolin distribution
After the cells were exposed to the indicated amount of shear-stress, the
coverslip was removed from the chamber, washed with PBS, fixed with 3%
paraformaldehyde in PBS and processed for indirect immunofluorescence
localization of caveolin-1. Images of 300-500 cells were recorded and each
cell was divided into eight equal sectors (45° each) using the nucleus as
the pivot point for sectioning (Fig.
4). Each sector was designated as belonging to either region A, B,
C, D or E as shown in Fig. 4.
Each cell was then scored and assigned to one of these regions according to
where the heaviest caveolin staining occurred on the cell margins. Cells that
exhibited caveolin-1 staining in more than one region were not classified.
Approximately 10-15% of the cells were not classified.
|
Western blot analysis
Cells that had been subjected to shear stress as described above were
washed with ice-cold PBS and solubilized in 500 µl RIPA buffer (1% Nonidet
P-40, 20 mM Tris-HCl, pH 7.4, 0.15 M NaCl, 0.5% sodium deoxycholate, 2 mM
EDTA, 2 mM EGTA, 0.1% SDS, 0.2 mM Na2MoO4, 10 mM NaF, 1
mM Na3VO4, 1 mM phenylmethylsulfonyl fluoride, 5
µg/ml leupeptin, 5 µg/ml antipain, 5 µg/ml pepstatin A, 0.2 unit/ml
aprotinin). Lysates were centrifuged at 26,000 g for 30
minutes. The supernatants were mixed with SDS sample buffer and separated
using a 15% PAGE. Gels were transferred to ImmobilonTM
polyvinylidene difluoride membranes (Millipore, Bedford, MA). Membranes were
blocked in TBS (20 mM Tris-HCL, 137 mM NaCl, pH 7.6) plus 5% nonfat milk and
0.1% Tween-20, and then incubated for 1 hour in the presence of caveolin pAb
diluted in blocking solution. Membranes were washed in PBS and incubated with
anti-rabbit IgG horseradish peroxidase-conjugated antibody. The blots were
developed using Enhanced Chemiluminescence kit (Amersham, Piscateway, NJ) and
analyzed by a GS363 Molecular Imager System (Bio-Rad, Hercules, CA).
Detection of mRNA
PT/PCR was performed to quantify the mRNA levels of caveolin as previously
described (Ando et al., 1994).
The cDNA samples were co-amplified by PCR with primer pairs for caveolin-1 and
glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The primer pairs for
caveolin-1 were 5'-CAACAAGGCTATGGCAGAGG-3' and
5'-CGTAGATGGAATAGACACGGC-3', and for GAPDH were
5'-GGAAGCTCGTCATCAATGG-3' and
5'-AGGAGGCATTGCTGACAATC-3'. 10 µl of amplified product was
sampled every other cycle, and separated on a 5% polyacrylamide gel. To
quantify the PCR products, the radioactivity of each band was measured with a
GS363 Molecular Imager, and plotted against the number of PCR cycles on a
semi-logarithmic scale. From the sigmoid curve, the cycle in which the
operating range of the PCR was linear was selected and the ratio of
radioactivity caveolin to GAPDH in that cycle was designated as the relative
amount of caveolin mRNA.
EM techniques
Endothelial cell cultures were prepared as described above except that a
10.5x22 mm Thermanox coverslip (Electron Microscopy Sciences, Ft
Washington, PA) was substituted for the glass coverslip. Cells were exposed to
20 dynes/cm2 for 24 hours in a flow chamber customized for the
smaller coverslip. The coverslip was removed and marked to indicate the
orientation of laminar flow. The cells were fixed with 2% glutaraldehyde in
0.1 M phosphate buffer pH 7.6 for 30 minutes at room temperature followed by
1% OsO4 in 0.1 M phosphate buffer for 1 hour at room temperature.
Multiple coverslips with the same orientation were stacked together and
embedded in Epon 812. The embedded coverslip stacks were cut out of the Epon,
marked and then oriented in flat embedding molds so that the coverslips were
parallel with the bottom of the mold. The molds were infiltrated with Epon and
polymerized. These blocks were removed from the mold, positioned so that
sections would be cut perpendicular to the coverslip and oriented with respect
to the direction of laminar flow. These sections were mounted on slot grids
and viewed with a JEOL 1200 electron microscope.
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Results |
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Sheer stress induces redistribution of caveolae
Endothelial cells in situ are continually exposed to laminar shear stress,
which is a mechanical activity that has been found to stimulate multiple
signaling pathways (Davies,
1995; Traub and Berk,
1998
). We used immunofluorescence to determine whether shear
stress, like cell migration, also caused the rearrangement of caveolin-1
patches on the cell surface (Fig.
2). Cells grown on coverslips were either examined directly
(unstressed) or placed in a parallel-plate flow chamber and subjected to a
laminar shear stress of 20 dynes/µm2 for 24 hours
(Fig. 2, stressed, arrows
indicate direction of flow). The cells were fixed and processed for indirect
immunofluorescence co-localization of caveolin-1, actin and vinculin. The
normal, patchy distribution of caveolin-1 was evident in unstressed cells, and
these patches could be found in all regions of the cell
(Fig. 2A). In stressed cells,
by contrast, virtually all of the caveolin-1 staining was confined to the
upstream edge of the cell (Fig.
2E, white arrows). Some cells had long extensions that protruded
from the upstream portion of the cell in the direction of fluid flow
(Fig. 2, yellow asterisk). The
entire length of these extensions was stained heavily with the caveolin-1 pAb.
The stressed cells appeared to have an increased number of actin stress fibers
(compare Fig. 2B with F). The
number of vinculin positive spots was not changed (compare
Fig. 2C with G) in shear
stressed cells.
|
In separate experiments (Fig. 3), we compared the distribution of caveolin-1 (Fig. 3A,E) with either actin (Fig. 3B), tubulin (Fig. 3C) or clathrin AP-1/2 (Fig. 3F) in shear stressed cells. Whereas the caveolin-1 patches redistributed to the upstream edge of cells (white arrows), there was no change in the distribution of either AP-1/2 or tubulin. The number of stress fibers was increased and in many cells they tended to be oriented parallel to the direction of fluid flow. The polarization of caveolin-1 appeared to occur before stress fiber reorganization (data not shown).
|
The redistribution of the caveolin-1 patches was dependent on both the
strength of the shear stress and the time of exposure to stress
(Fig. 4). Cells were either
incubated in the presence of an increasing shear stress force for 44 hours
(Fig. 4, right panel) or
exposed to a constant force (20 dynes/cm2) for increasing times up
to 68 hours (Fig. 4, left
panel). The number of cells that had the majority of the caveolin-1 staining
in either region A, B, C, D, or E was tabulated
(Fig. 4, top-left). At the
beginning of the experiment, equal numbers of cells were found with caveolin-1
patches in the five regions. With either increasing time or increasing shear
stress force, however, the number of cells with caveolin-1 patches in region A
increased. The increase in the number of region A cells was matched by a
corresponding decline in cells with caveolin-1 in regions C, D or E. Little
change was seen in the number of cells with caveolin-1 patches in region B.
After 68 hours of incubation, >80% of the cells were found to have
caveolin-1 patches in region A. As little as 1.5 dynes/cm2 caused
an increase in the number of cells with caveolin-1 patches in region A, but it
took 20 dynes/cm2 to cause the number of cells with patches in this
region to reach a maximum (58% of the cells).
A change in the distribution of caveolin-1 patches suggests that these
conditions cause a redistribution of caveolae. Numerous studies have shown
that caveolin-1 is a marker for caveolae in endothelial cells
(Schnitzer et al., 1995).
Nevertheless, we used thin section electron microscopy to confirm that the
relocation of caveolae accounted for the redistribution of caveolin-1
staining. Cells were grown on plastic coverslips and subjected to 20
dynes/cm2 of stress for 24 hours. The coverslip was fixed and
positioned during embedding so that individual cells oriented in the direction
of media flow could be viewed throughout their entire length.
Fig. 5A shows at low
magnification a longitudinal view of a typical cell with the upstream region
of the cell towards the right (arrow indicates direction of flow). A higher
magnification of this region of the cell shows an anastomosing network of
flask-shaped, tubular and vesicular caveolae
(Fig. 5B, arrows). Many of the
caveolae vesicles were tightly associated with smooth ER
(Fig. 5B, asterisk).
Progressing downstream of this region, the number of recognizable caveolae
declined. Very few caveolae could be identified in the body of the cell or at
its downstream edge (data not shown). Therefore, caveolin-1 staining at the
trailing edge of shear-stressed cells corresponds to sites of massive caveolae
accumulation. Since 20 dynes/cm2 of shear stress for up to 24 hours
did not change the level of either caveolin-1 protein
(Fig. 6A) or mRNA
(Fig. 6B), the caveolae that
have accumulated at the upstream region probably came from other locations in
the cell.
|
|
Signal transduction from caveolae-rich regions of shear stressed
cells
Previously we have shown that agonists such as ATP and bradykinin stimulate
Ca2+ wave initiation in caveolin-rich regions of the cell
(Isshiki et al., 1998). These
results suggest that caveolae contain machinery that links G-protein-coupled
receptors to the release of Ca2+ from ER stores. A key molecular
component of this machinery is the heterotrimeric G protein subunit
G
q G
q has been localized to caveolae
(Oh and Schnitzer, 2001
), so
we used the same antibody to determine the distribution of
G
q in shear-stressed cells
(Fig. 7). As reported
previously, we found good colocalization
(Fig. 7). of caveolin-1
(Fig. 7A) and
G
q (Fig. 7B)
in unstressed cells, although not all of the caveolin-rich areas were positive
for G
q. The application of 20 dynes/cm2 of fluid
shear to the cells in the direction of the arrow caused the polarization of
caveolin-1 to the upstream edge of the cell
(Fig. 7D). Large portions of
these caveolin-1 patches were also positive for G
q
(Fig. 7E,F). Importantly, not
all of the caveolin-1 patches were positive for G
q,
suggesting that not all caveolae are equipped with this signaling
molecule.
|
We next examined whether or not Ca2+ wave initiation migrated with caveolae to the upstream edge of shear stressed cells (Fig. 8). Unstressed cells (A-D) were analyzed first. Endothelial cells were grown on coverslips for 2 days before being loaded with the Ca2+ indicator dye Indo-1 AM. Ca2+ waves were initiated by stimulating cells with 0.5 µM ATP. Fig. 8D shows images of cells taken at 0.38 second intervals after the addition of ATP. Within 10 seconds after ATP stimulation, focal sites of Ca2+ release were seen (Fig. 8, white arrows). Ca2+ release spread from these sites throughout the cell. Following the Ca2+ imaging phase of the experiment, the cells were fixed and processed for immunofluorescence detection of caveolin-1 (Fig. 8A-C). Each site of Ca2+ wave initiation corresponded to a caveolin-1 patch (Fig. 8A,D, compare arrows). Not all caveolin-rich regions, however, were associated with sites of Ca2+ release.
|
Sites of Ca2+ wave initiation were next recorded in cells that had been exposed to a laminar shear stress of 20 dynes/cm2 for 24 hours (Fig. 8E-H). Cells were loaded with Indo-1 AM and stimulated with 2 µM ATP before taking images of the cells at 0.38 second intervals (H). Ca2+ wave initiation occurred exclusively at the upstream edge of cells (H, arrow). Importantly, the initial site coincided with a cell extension that was oriented upstream of the cell. With time, Ca2+ release spread progressively towards the downstream end of the cell. Staining of cells with caveolin-1 pAb showed that the upstream edges and the extension were covered with caveolin-1. We conclude that sites of Ca2+ wave initiation follow the caveolae as they become polarized on the cell surface in response to shear stress.
In addition to repositioning sites of Ca2+ wave initiation, shear stress also caused a marked desensitization to ATP (Fig. 9). Sequential addition of increasing concentrations of ATP to either stressed (bottom panel) or unstressed (upper panel) cells showed that it took a tenfold higher concentration of ATP to initiate a Ca2+ wave in stressed cells than it did in unstressed cells.
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Discussion |
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Our observations stand in contrast to a recent report that growth
factor-stimulated chemotaxis causes the relocation of raft markers to the
leading edge of MCF-7 adenocarcinoma cells
(Manes et al., 1999). This
study did not determine if caveolin-1 moved to the leading edge, so it is not
clear whether they were observing the behavior of caveolae or non-caveolae
rafts. The possibility remains, therefore, that caveolae move to the trailing
end of a migrating cell when it is moving in response to a mechanical stimulus
and to the leading end when migrating in response to a chemical stimulus.
Alternatively, the polarization of caveolae during cell migration is cell-type
specific.
Several hours were required to achieve complete polarization of caveolae in
migrating endothelial cells. However, Kogo and Fujimoto saw migration of
GFP-caveolin-1 to the cleavage furrow within minutes after cells enter
cytokinesis (Kogo and Fujimoto,
2000). GFP-caveolin-1 also accumulates at sites of cell-cell
contact during contact inhibited cell growth
(Volonte et al., 1999
).
Assuming that GFP-caveolin-1 is a reliable marker for caveolae in these cells,
then caveolae must be mobile and be able to assume rapidly different spatial
arrangements in response to specific stimuli. The finding that a number of
different conditions cause caveolae relocation suggests that this is a general
mechanism cells use for spatially organizing specific activities at the cell
surface.
An important future area of investigation is to determine the mechanism of
caveolae polarization. One possibility is that recycling caveolae vesicles
collect at specific regions of the cell in response to the different stimuli.
Another is that caveolae behave like actin patches in yeast cells
Waddle et al., 1996) and move
in the plane of the membrane to the trailing edge of the cell. Finally,
caveolae might not move at all but accumulate at the end of a migrating cell
as the result of differential movement of other organelles. A number of
standard inhibitors such as staurosporin, herbimycin A, C3 toxin and the
MEK-1/2 inhibitor PD98059 had no affect on caveolae polarization (data not
shown).
Previous studies have shown that Ca2+ wave formation in the
non-migrating endothelial cell occurs at caveolin-rich regions of the cell
surface (Isshiki et al.,
1998). Importantly, not all caveolin-rich regions in the same cell
are competent to initiate waves. Tests to probe the functionality of these
regions in individual cells showed that wave formation occurred at the same
caveolin-1 patch in cells repeatedly exposed to the same stimuli or
sequentially exposed to different stimuli. Therefore, only subsets of caveolae
appear to contain the signaling machinery that regulates Ca2+
release from the ER. This is in agreement with our observation that not all
caveolin-1 patches were positive for G
q. The simultaneous
relocation of G
q, sites of Ca2+ wave formation
and caveolae to the trailing edge of migrating cells is strong evidence that
caveolae can function as containers and carry signaling machinery to specific
locations in the cell.
We cannot rule out the possibility that Ca2+ waves originate
from non-caveolae membrane that has accumulated at the trailing edge of
migrating cells. Nevertheless, we found that numerous invaginated caveolae and
caveolae vesicles at the trailing edge were in close apposition to elements of
smooth ER (Fig. 5, asterisk).
These images give the impression that caveolae specifically interact with ER.
Caveolae can deliver molecules to the ER
(Benlimame et al., 1998), so it
is possible that caveolae/ER contact sites are locations where Ca2+
release is stimulated (Isshiki and
Anderson, 1999
).
Genetic studies of budding and fission yeast have identified a variety of
molecules that control cell polarity in these organisms
(Chant, 1999). Interestingly,
the same molecules seem to be important for both budding and fission. Four of
the critical regulatory molecules are the MAP kinase Mpk1, Rho1, PKC1 and
phosphoinositides. Functional homologues of these molecules, such as ERK1 and
ERK2) (Liu et al., 1997
), Rho
A (Michaely et al., 1999
),
multiple PKC isoforms (Mineo et al.,
1998
) and phosphoinositides
(Pike and Casey, 1996
), are
concentrated in caveolae. Caveolae also contain G-proteincoupled receptors
(Feron et al., 1997
), are
enriched in ERM proteins (Michaely et al.,
1999
) that recruit Rho A
(Tsukita et al., 1994
), and
contain various receptor and non-receptor tyrosine kinases
(Ko et al., 1998
;
Liu et al., 1996
). If
migrating caveolae can carry these signaling molecules to different locations
in the cell, then they should be found wherever caveolae accumulate during
cell migration and cytokinesis. Immunofluorescence has shown that ERM proteins
collect both at the trailing edge of migrating cells
(Serrador et al., 1997
) and in
the cleavage furrow during cytokinesis
(Kosako et al., 1999
).
Transfected Rho A is found concentrated at the cleavage furrow and sites of
cell-cell contact (Kosako et al.,
1999
). Endogenous urho1, the sea urchin Rho homologue, has been
localized to the cleavage furrow of dividing eggs
(Nishimura et al., 1998
).
Finally, PKC isoforms translocate to the uropod of lymphocytes in response to
whole body hyperthermia (Wang et al.,
1999
). If caveolae are the vehicle for moving signaling molecules
to these locations, then they may play a crucial role in establishing cell
polarity.
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Acknowledgments |
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