1 Epithelial Pathobiology Research Unit, Department of Pathology and Laboratory
Medicine, Emory University, Whitehead Biomedical Research Building, Atlanta,
GA 30322, USA
2 Max Planck Institute of Molecular Cell Biology and Genetics,
Pfotenhauerstrasse 108, 01307 Dresden, Germany
3 INSERM Unite 452, IFR 50, Faculté de Medecine, 28 Avenue de Valombrose,
F-06107, Nice, France
* Author for correspondence (e-mail: ahopkin{at}emory.edu)
Accepted 2 December 2002
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Epithelium, Rho GTPases, Tight junction, Paracellular permeability, F-actin
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The Rho family of small GTPases, comprising Rho, Rac and Cdc42, are
critical regulators of F-actin dynamics
(Hall, 1998;
Nobes and Hall, 1995
;
Ridley and Hall, 1992
).
Activation of Rho family members requires guanosine exchange factors (GEFs),
which catalyze the exchange of GDP for GTP. Conformational changes then allow
the GTPases to interact with multiple effector molecules involved in actin
cytoskeletal control (Aspenstrom,
1999
; Hall, 1990
).
Rho activity cycles are rapidly reversible, and are terminated upon hydrolysis
of GTP by GTPase-activating proteins.
The biological consequences of Rho GTPase activation have been extensively
explored in mesenchymal cells (Mackay et
al., 1997; Nobes and Hall,
1995
; Ridley and Hall,
1992
; Ridley et al.,
1992
). How Rho GTPases govern F-actin dynamics in non-mesenchymal
cells is less clear. Considering the close apposition between the
perijunctional F-actin ring and the epithelial TJ complex, it is conceivable
that Rho proteins, through cytoskeletal modification, could affect epithelial
barrier function. Several insights have been offered, utilizing diverse
pharmacological and molecular tools to interfere with Rho protein function.
Our initial investigations utilized a modified cell-permeant chimeric toxin
consisting of the Clostridium botulinum toxin C3 transferase (to
inhibit RhoA activity through ADP-ribosylation of Asp41), and the
receptor-binding domain of Diphtheria toxin to facilitate
internalization (Nusrat et al.,
1995
). In this system, the barrier function of T84 intestinal
epithelial cells was compromised with reductions in transepithelial resistance
(TER), enhancements in paracellular permeability and redistribution of ZO-1
and occludin away from the TJ membrane
(Nusrat et al., 1995
). Related
work has focused upon the impact of dominant-active and dominant-negative
small GTPase mutants on polarity and barrier function in Madin-Darby canine
kidney (MDCK) epithelial cells, and again demonstrated that antagonism of RhoA
function can adversely affect TJ structure/function
(Jou et al., 1998
).
Conversely, dominant-active RhoA mutants reportedly protect TJs during ATP
depletion (Gopalakrishnan et al.,
1998
). Mechanisms whereby TJs are influenced by GTPases have
recently been reviewed (Hopkins et al.,
2000
). Our study provides novel insights into the regulation of TJ
structure by activated Rho GTPases.
An array of bacterial toxins modulates Rho GTPase function
(Aktories, 1997;
Boquet, 1999
;
Boquet et al., 1999
;
Fiorentini et al., 1998a
;
Gyles, 1992
;
Lerm et al., 2000
;
Schmidt and Aktories, 1998
;
von Eichel-Streiber et al.,
1996
). Interference with the cytoskeleton is a common pathogenic
mechanism, exemplified by membrane ruffling events or `cup and pedestal'
formation during respectively Salmonella and Escherichia
invasion (Lesser et al.,
2000
). However, since cytoskeletal rearrangements during
Salmonella but not Escherichia invasion appear under Rho
control (Ben-Ami et al., 1998
),
Rho-independent mechanisms probably exist during distinct forms of bacterial
invasion.
Cytotoxic necrotizing factor (CNF-1), a toxin derived from necrotizing
Escherichia coli, has been implicated in the pathogenesis of
prostatitis (Andreu et al.,
1997), urinary tract infections
(Blanco et al., 1995
) and
others (Sears and Kaper,
1996
). Its mechanism of action involves deamidation of Gln63 of
Rho or Gln61 of Rac/Cdc42, resulting in constitutive activation of GTPase
signaling via inhibition of GTP hydrolysis
(Flatau et al., 1997
;
Lerm et al., 1999a
;
Schmidt et al., 1997
). CNF-1
was initially identified as a toxin that stimulated the formation of giant
multinucleated cells (Caprioli et al.,
1983
; Caprioli et al.,
1984
). Bacterial strains secreting CNF-1 are reportedly cytopathic
to epithelial cells (De Rycke et al.,
1997
), however this might reflect the influence of additional
virulence factors other than CNF-1 (Island
et al., 1998
; Island et al.,
1999
). Evidence actually suggests that CNF-1 protects against
apoptosis (Fiorentini et al.,
1997
; Fiorentini et al.,
1998b
). Cytoskeletal effects attributed to purified CNF-1 toxin
include stimulation of phagocytotic behavior
(Falzano et al., 1993
),
bacterial internalization (Kazmierczak et
al., 2001
) and stress fiber aggregation in epithelial/mesenchymal
cells (Gerhard et al., 1998
;
Hofman et al., 1998
). However,
little is known about the effects of CNF-1 on apical F-actin in epithelial
cells, including that in the perijunctional ring that is affiliated with TJs.
Conflicting reports have suggested both increased
(Hasegawa et al., 1999
) and
decreased (Gerhard et al.,
1998
) epithelial barrier function following constitutive Rho
activation. Our study conducted an in-depth investigation of CNF-1 effects on
epithelial TJ structure, barrier function and polarized organization of
F-actin/associated proteins. The intestinal epithelial cell line T84 was
chosen as a model for our studies
(Dharmsathaphorn et al., 1984
;
Dharmsathaphorn and Madara,
1990
; Madara and
Dharmsathaphorn, 1985
) because its high transepithelial resistance
to passive ion flow and well-ordered F-actin/TJ structures make it a stringent
model for mechanistic examination of alterations in barrier function. Emphasis
was placed on morphological localization of F-actin pools and associated TJ
proteins before and after CNF-1 treatment, in an attempt to resolve
ambiguities regarding the contribution of Rho proteins to the genesis and
maintenance of barrier function in vitro. Our study also presents novel data
showing toxin-induced internalization of TJ proteins in caveole and
early/recycling endosomes, suggesting a mechanism for fast re-establishment of
barrier function upon CNF-1 removal.
![]() |
Materials and Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
GTPase activation assays
To test for RhoA activation in T84 intestinal epithelial cells upon CNF-1
treatment, a rhotekin-binding assay was performed as previously described
(Kranenburg et al., 1999;
Sagi et al., 2001
;
Seasholtz et al., 2001
).
Briefly, T84 monolayers grown to confluence on 5 cm2 polycarbonate
transwells (Costar) were treated with either CNF-1 (1 nM) or a corresponding
volume of vehicle for 24 hours, and harvested into Rho lysis buffer (25 mM
HEPES pH 7.4, 125 mM NaCl, 1% Igepal CA-630, 10 mM MgCl2, 1 mM
EDTA, 2% glycerol, and protease inhibitors PMSF 250 µM, leupeptin 5
µg/ml, chymostatin 10 µg/ml, pepstatin 0.25 µg/ml, aprotinin 2
µg/ml). Following a brief centrifugation to remove cell debris, lysates
from control and CNF-treated cells containing equivalent protein
concentrations were rotated for 45 minutes with 40 µL slurry of a GST
fusion protein composed of the Rho-binding domain of the specific Rho effector
rhotekin coupled to agarose beads (Upstate Biotechnology, Lake Placid, NY).
Addition of GTP
S to additional whole cell lysates served as a positive
control for Rho activation. Beads were collected by centrifugation and washed
three times with lysis buffer. Beads were resuspended in 2 µL 1M
dithiothreitol and 48 µL non-reducing sample buffer, boiled for 5 minutes
and subjected to SDS-PAGE on a 12% Tris-HCl gel. Whole cell lysates from both
control and CNF-treated cells were run in parallel to determine baseline
levels of total RhoA protein. Separated proteins were transferred to
nitrocellulose and western blotted with a monoclonal antibody to RhoA (Santa
Cruz Biotechnology, Santa Cruz, CA). Using related methods, the activation
status of Rac and Cdc42 was also tested in T84 monolayers following CNF-1
treatment. In this case, the Rac effector PAK-1 conjugated to agarose beads
was used to pulldown activated Rac from control and CNF-treated monolayers,
using a Rac activation assay kit (Upstate Biotechnology). Positive controls
consisted of T84 whole cell lysates treated with GTP
S to irreversibly
activate Rac. As described above for Rho, activated Rac from the pulldown
experiments and total Rac from whole cell lysates was detected by SDS-PAGE and
western blot analysis using a monoclonal antibody to Rac.
Since PAK-1 is also an effector for activated Cdc42, the same nitrocellulose membranes were then stripped in ß-mercaptoethanol-based buffer and reblotted with a monoclonal antibody to Cdc42 (Upstate Biotechnology).
Paracellular permeability assays
T84 monolayers on permeable supports were exposed apically or basolaterally
to CNF-1 or vehicle for 6, 24 or 48 hours. Paracellular permeability to
fluorosceinated dextran (FD-3; MW 3000) was assessed according to previously
published methods (Sanders et al.,
1995). Briefly, TER was measured, monolayers were washed in Hanks
balanced salt solution/10 mM HEPES (HBSS+) and then equilibrated at
37°C for 10 minutes on an orbital shaker. Monolayers were loaded apically
with 1 mg/ml FD-3 (Molecular Probes, Eugene, OR) at time=zero. Basolateral
samples were taken at t=0 and 120 minutes, and fluorescence intensity was
analyzed on a CytoFluor 2350 Fluorescence Measurement System (Millipore,
Cambridge, MA). FD-3 concentrations transported were extrapolated from a
standard curve (generated by diluting known concentrations of fluorescent
tracer), and expressed as µM FD-3 transported/cm2/hour.
Numerical values from individual experiments were pooled and expressed as mean
± standard error of the mean (s.e.m.) throughout. Control and test
values at each time point of CNF/vehicle treatment were compared by two-tailed
unpaired Student's t-tests, with statistical significance assumed at
P values<0.05.
Immunofluorescence of TJ/adherens junction (AJ) proteins, F-actin and
actin-binding proteins
T84 monolayers exposed to CNF-1 or vehicle for 24 hours were washed, fixed
in absolute ethanol (20 minutes, -20°C) and blocked in 5% normal goat
serum [1 hour, room temperature (RT)]. Monolayers were incubated in humidity
chambers for 1 hour with primary antibodies to human occludin, ZO-1, (Zymed
Labs, San Francisco, CA), JAM-1 (C. A. Parkos, Emory University, Atlanta, GA),
E-cadherin (HECD-1 hybridoma supernatant; A. S. Yap, University of Queensland,
Australia), ß-catenin, villin, paxillin, caveolin-1, lysosomal-associated
membrane protein-1 (LAMP-1; Transduction Laboratories, Lexington, KY),
transferrin receptor (Molecular Probes), early endosomal antigen-1 (EEA-1) or
Rab11 (Santa Cruz Biotechnology). Monolayers were washed, probed with FITC- or
rhodamine-conjugated goat anti-mouse/-rabbit IgG (1 hour, RT; Jackson
Immunoresearch Labs, West Grove, PA) and mounted on phosphate-buffered
saline/glycerol/p-phenylenediamine, 1:1:0.01 (v/v/v). Monolayers
double-labeled for p-MLC and ZO-1 were blocked in 3% BSA and co-incubated with
goat anti-human p-MLC (Santa Cruz Biotechnology) and rabbit anti-human ZO-1
antibodies. Donkey anti-goat and anti-rabbit antibodies conjugated to
fluorescent red or green Alexa dyes (Molecular Probes) were used for detection
of the respective primary antibodies. For immunolocalization of filamentous
actin (F-actin), monolayers were fixed in 3.7% paraformaldehyde (10 minutes,
RT), permeabilized in 0.5% Triton X-100 (30 minutes, RT), incubated with
rhodamine-conjugated phalloidin (Molecular Probes; 1 hour, RT), washed and
mounted as above. Monolayers were visualized on an LSM510 confocal microscope
(Zeiss Microimaging, Thornwood, NY). Images shown are representative of at
least six experiments, with multiple images taken per slide.
Immunogold electron microscopy
T84 monolayers treated basolaterally with CNF-1 or vehicle for 24 hours
were washed in HBSS+, fixed in 3.7% paraformaldehyde for 20 minutes
at room temperature and processed for immunogold electron microscopy as
previously described (Nusrat et al.,
2000). In short, sections were first labeled with polyclonal
antibodies to occludin and detected by protein A coupled to 15 nm gold
particles. After fixation and blocking, sections were labeled with polyclonal
caveolin-1 antibodies and detected by protein A coupled to 10 nm gold
particles. Primary antibodies were omitted from negative control samples. The
presence of occludin and caveolin-1 at the TJ was quantitated in control and
CNF-1-treated cells. Previously, we had calculated that 90% of the immunogold
labeling for occludin was found in the uppermost 250 nm of the basolateral
plasma membrane where the apical plasma membranes of two neighboring cells
meet (Nusrat et al., 2000
).
Therefore, numbers of gold particles in this area were counted. Statistical
differences between the two occludin groups were tested with a Welch-test
since the variances of the two groups were not equal; and statistical
differences between the two caveolin-1 groups were tested with a Student's
t-test.
Immunoblotting for TJ/AJ proteins in epithelial cells
Confluent T84 monolayers on 5 cm2 permeable supports were
incubated basolaterally with CNF-1 or vehicle for 24 hours and washed in
HBSS+. Monolayers were scraped into Relax buffer (KCl 100 mM, NaCl
3 mM, MgCl2 3.5 mM, HEPES pH 7.4 10 mM) containing 1% Triton X-100
with protease inhibitors (as above) and phosphatase inhibitors (sodium
fluoride 25 mM, sodium orthovanadate 10 mM). Cell suspensions were dounced 20
times with a Dounce homogenizer, and the nuclei removed by low-speed
centrifugation (1500 g, 5 minutes, 4°C). Equivalent
concentrations of post-nuclear lysate proteins (10 g/lane) from control
and CNF-treated monolayers were subjected to SDS-PAGE and immunoblotted with
antibodies to TJ, AJ and actin-binding proteins (suppliers as above) and actin
(Sigma). In order to determine whether CNF-1 caused tyrosine phosphorylation
of paxillin, we treated monolayers for 24 hours with CNF-1 or vehicle,
prepared lysates and performed SDS-PAGE as above, and immunoblotted with an
antibody specific for phospho-paxillin (on Tyr118; Santa Cruz
Biotechnology).
Calcium switch assay
T84 monolayers were transiently exposed to ethylene
glycol-bis(ß-aminoethyl ether)N,N,N',N'-tetraacetic acid
(EGTA; 2 mM, 20 minutes, 37°C) in calcium- and magnesium-free HBSS with 10
mM HEPES (HBSS-) to chelate extracellular calcium and disrupt
intercellular junctions (Liu et al.,
2000; Parkos et al.,
1995
). After washing, monolayers were allowed to recover in
complete cell culture media (containing calcium) in the presence of CNF-1 or
vehicle control. TER was monitored during recovery as an index of TJ function.
Additionally, monolayers were fixed and immunostained for occludin and
E-cadherin (as described above) immediately after EGTA treatment, and
following 9 hours recovery in the presence of CNF-1 or vehicle.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
CNF-1 enhances paracellular permeability across intestinal epithelial
monolayers
T84 intestinal epithelial cells were grown as polarized monolayers on
permeable supports, which at confluence displayed high stable TER to passive
ion flow (>1000 .cm2). As shown in the concentration
response curve, basolateral exposure of T84 cells to CNF-1 (0.005-5 nM;
plotted on a log scale) for 24 hours evoked significant reductions in TER
(Fig. 2). Maximal reductions
were observed with 1 and 5 nM CNF-1, hence 1 nM was chosen for subsequent
experiments. The bioactive effect of CNF-1 was polarized to the basolateral
surface, since apical exposure to toxin did not significantly alter TER over
control levels (data not shown).
|
To complement our electrophysiological observations documenting CNF-induced
reductions in TER and to confirm enhanced paracellular permeability, we
examined the flux of a macromolecule across T84 monolayers exposed to CNF-1
(Fig. 2; n=3-5
experiments). FD-3 was chosen as a marker of paracellular transport
(Sanders et al., 1995). T84
epithelial monolayers were treated for 6, 24 or 48 hours with CNF-1 or
vehicle, and flux assays performed over the subsequent 2-hour period. Control
monolayers (black bars) exposed to vehicle exhibited low levels of FD-3 flux
from the apical into the basolateral compartment, which were similar across
the three time points measured. FD-3 fluxes across monolayers treated apically
with CNF-1 (light gray bars) were statistically indistinguishable from
controls at the same time points. However, monolayers treated basolaterally
with CNF-1 (dark gray bars) showed significant enhancements in paracellular
flux of FD-3. Permeability enhancements were dependent upon the length of
exposure to the toxin, with flux following 24 or 48 hours incubation with
CNF-1 being significantly different to control fluxes at the same time points
(P<0.01).
CNF-1 induces redistribution of key TJ proteins involved in
epithelial barrier function
Since the epithelial TJ protein complex is a major regulator of transport
through the paracellular route, we examined the effect of CNF-1 on the
immunolocalization of candidate TJ proteins
(Fig. 3). T84 intestinal
epithelial monolayers incubated either apically or basolaterally with CNF-1 (1
nM, 24 hours) or vehicle control were immunostained with antibodies to
occludin, ZO-1 and JAM and examined by confocal microscopy. Occludin
distribution in control T84 epithelial monolayers
(Fig. 3a) mirrored that in
monolayers incubated apically with CNF-1
(Fig. 3b); namely a
characteristic cobblestone pattern shown in en face confocal micrographs
captured at the level of the TJ. Monolayers incubated basolaterally with CNF-1
(Fig. 3c) displayed occludin
reorganization in the TJ plane, manifested as decreased occludin localization
at the TJ membrane (arrow) with potential internalization (#). Total occludin
protein levels (Fig. 3j) were
similar in cell lyastes from both control (lane 1) and CNF-treated monolayers
(lane 2).
|
The morphological distribution of another TJ protein, ZO-1, a cytoplasmic plaque protein that affiliates with the underlying F-actin cytoskeleton, was also examined. ZO-1 staining in control monolayers (exposed to vehicle alone; Fig. 3d) resembled that of occludin, being confined to distinctive TJ ring formations. No substantial alterations were visible upon apical exposure to CNF-1 toxin (Fig. 3e); however basolateral exposure to toxin for the same time period (Fig. 3f) stimulated dramatic reorganization of ZO-1 away from the TJ plane. Like occludin in cells treated basolaterally with CNF-1, there was partial or, in some cases, complete loss of the ring structure that is morphologically characteristic of these proteins in en face confocal images. All images presented are representative of at least four independent experiments, and all morphological alterations were similar or intensified upon prolonged basolateral incubation with CNF-1 (48 hours). Morphological reorganization events were not accompanied by alterations in the total ZO-1 protein levels in control versus CNF-treated monolayers (Fig. 3k).
The ring pattern of JAM-1 immunolocalization in en face images of control
monolayers (Fig. 3g) resembled
that of occludin and ZO-1, with sharp rings indicative of localization at the
distinct lateral TJ membrane. Some additional staining was observed along the
lateral membrane in the xz or vertical plane (not shown). JAM
distribution in TJs of monolayers treated apically with CNF-1 for 24 hours
(Fig. 3h) was virtually
identical to that in control monolayers. By contrast, monolayers exposed
basolaterally to CNF-1 for 24 hours (Fig.
3i) displayed some loss of JAM definition at the membrane. The
`blurred' rings probably represent some JAM redistribution away from the TJ
membrane to a cytoplasmic region just below the lateral membrane. This could
be important in the loss of barrier function since the presence of JAM at the
TJ membrane is essential for correct re-localization of occludin during TJ
assembly (Liu et al., 2000).
Western blot data (Fig. 3l) did
not reveal any differences in total JAM protein levels between control and
CNF-treated monolayers.
CNF-1 induces internalization of occludin and caveolin-1 in
caveolar/recycling endosomal structures
By immunogold electron microscopy, we have previously shown that occludin
predominantly (>90%) localizes to the apical-most 250 nm of the basolateral
plasma membrane (Nusrat et al.,
2000). In control epithelial cells, occludin (pseudo-colored in
red) displays this pattern (Fig.
4Aa, arrow); however, upon basolateral treatment of the cells with
CNF-1 for 24 hours, some occludin disappeared from this TJ domain into an
intracellular compartment (Fig.
4Ab, #). A percentage of internalized occludin moved into
membranous structures ultrastructurally resembling endosomes/caveolae
(Fig. 4Ac), and co-localized
with caveolin-1, a caveolar scaffolding protein (particles colored in black).
We quantitated the redistribution of occludin and caveolin-1 from TJs after
CNF-1 treatment (Fig. 4B, occludin in n=25 TJs; caveolin-1 in n=10 TJs for both
control and CNF-treated cells). In control cells, we found 6.8±0.5
occludin gold particles per TJ (mean±s.e.m.) while, in CNF-treated
cells, we only found 1.9±0.3 occludin gold particles per TJ domain.
This approximately 3.5-fold difference was statistically significant
(P<0.00001). Analogous to our findings with occludin, caveolin-1
was also redistributed away from the TJ following CNF-1 incubation. We
observed 6.7±0.6 caveolin-1 gold particles per TJ in control cells, but
only 2.4±0.4 caveolin-1 gold particles (P<0.0001) per TJ of
CNF-treated cells. Similar trends were seen for ZO-1 internalization (data not
shown).
|
In order to address more precisely the nature of the structures into which occludin was internalized upon CNF-1 treatment (1 nM, 24 hours), we performed double-immunolabeling/confocal microscopy for occludin and a range of endosomal/caveolar markers. Occludin in control cells colocalized with a TJ-associated pool of caveolin-1 (Fig. 5a2, arrowhead) in ring structures distinct from the large pool of caveolin-1 highlighting the apical brush border. Upon treatment with CNF-1, internalization of occludin (Fig. 5b1, arrows) and caveolin-1 (Fig. 5b2, arrowheads) was observed. Areas of co-localization between internalized occludin and caveolin-1 are shown in Fig. 5b3 (#). Occludin staining following CNF-1 incubation was also examined in the context of the early endosomal markers EEA-1 and the transferrin receptor. The transferrin receptor exhibited diffuse cytoplasmic staining in control cells (Fig. 5c2), and no overlap with occludin (Fig. 5c3) was observed. Following CNF-1 incubation and characteristic occludin internalization (Fig. 5d1, arrow), the transferrin receptor was observed to condense into areas also suggestive of internalization sites (Fig. 5d2, arrowhead). However, although neighbors, these internalization sites seemed morphologically distinct from each other (Fig. 5d3, #). EEA-1 in control cells (Fig. 5e2) displayed a diffuse cytoplasmic staining pattern similar to that of the transferrin receptor, in addition to a distinct membranous pool in rings (Fig. 5e2, arrow) that co-localized with those of occludin (Fig. 5e1, occludin alone; Fig. 5e3, composite of occludin and EEA-1). Internalization of occludin induced by exposure to CNF-1 (Fig. 5f1, arrows) occurred in compartments similar to those of internalized EEA-1 (Fig. 5f2, arrowheads). Morphological overlap of some of these internalized structures (Fig. 5f3, #) suggests that at least a pool of occludin can be internalized in EEA-1-positive early endosomes following CNF-1 treatment.
|
We next explored the possibility that this sub-pool of early endosomes
could be recycling endosomes, which target their cargo back to the membrane
rather than onwards to late endosomes/lysosomes
(Casanova et al., 1999;
Ullrich et al., 1996
). Thus,
we stained for occludin and the recycling endosomal marker Rab11 in control
cells and those treated with CNF-1. Rab11 positivity in control cells
(Fig. 5g2) seemed to be
confined to sub-membranous structures resembling vesicles. No co-localization
with the characteristic TJ rings of occludin was observed in control cells
(Fig. 5g1, occludin alone;
Fig. 5g3, occludin/Rab11
composite). However, following treatment with CNF-1, internalization of
occludin (Fig. 5h1, arrow) and
condensations of Rab11 (Fig.
5h2, arrowhead) partially overlapped with each other
(Fig. 5h3, #). This suggests
that at least a pool of internalized occludin might be recycled back to the
membrane in Rab11-positive recycling endosomes.
|
Since our biochemical experiments showed that occludin was not degraded following CNF-1 treatment, we double-stained monolayers for occludin and the late endosomal marker LAMP-1. Anti-LAMP-1 stained control T84 cells (Fig. 5i2) in a characteristic ring pattern reminiscent of, but not coincident with, occludin (Fig. 5i1). In the composite image (Fig. 5i3), it can be seen that both these proteins are in the same plane but are mainly adjacent to each other rather than in an overlapping pattern (#). Incubation with CNF-1 induced fragmentation of LAMP-1 rings, with apparent internalization in submembranous vesicles (Fig. 5j2). Absolutely no overlap was observed between areas of internalized occludin (Fig. 5j1, arrows; Fig. 5j3, #) and LAMP-1 positivity, suggesting that occludin internalized after CNF-1 treatment is not designated for a lysosomal degradative pathway. In addition, there was a lack of co-localization between internalized occludin and clathrin following incubation with CNF-1 in our system (data not shown).
CNF-1 does not abolish the localization of AJ proteins in ring
structures at cell-cell borders
Immunolocalization of E-cadherin and ß-catenin was performed in order
to evaluate the influence of CNF-1 on AJs. As shown in
Fig. 6, in control cells both
E-cadherin (Fig. 6a) and
ß-catenin (Fig. 6c) were
visualized in a ring pattern by en face confocal imaging, consistent with
their localization in AJs. Minor changes in both E-cadherin
(Fig. 6b) and ß-catenin
(Fig. 6d) distribution were
observed following basolateral incubation with CNF-1 for 48 hours, mainly a
slight increase in staining intensity at cell-cell borders, or potentially a
subtle diffusion of the same away from the membrane. However, the localization
of both E-cadherin and ß-catenin in ring structures was essentially
preserved despite CNF-1 treatment, in contrast to ring structures of TJ
proteins that were severely disrupted under the same conditions. Expression of
total cellular E-cadherin and ß-catenin was not influenced by CNF-1
treatment for various time periods, as determined by western blot analysis
(Fig. 6e,f respectively; lanes
1-3 control, lanes 4-6 CNF-1).
|
CNF-1 alters the localization of a TJ-associated pool of p-MLC
TJ proteins affiliate with the underlying perijunctional F-actin ring and
this affiliation is important in the regulation of TJ function
(Madara, 1987). In addition,
MLC phosphorylation has also been shown to regulate TJ function
(Turner and Madara, 1995
).
Therefore, we investigated whether epithelial exposure to CNF-1 could alter
the distribution of p-MLC in TJs. There was extensive co-localization between
the ring structures of ZO-1 and a pool of p-MLC at the level of the TJ in
control T84 epithelial cells (Fig.
7a-c). This co-localization was mirrored in monolayers treated
apically for 24 hours with CNF-1 (Fig.
7d-f). However, upon basolateral treatment with the toxin, severe
disruption in the perijunctional staining pattern of not only ZO-1
(Fig. 7g) but also p-MLC
(Fig. 7h) occurred. Almost
complete fragmentation of the ring structures of both proteins was observed at
the level of the TJ, although punctate fragments of ZO-1 and p-MLC that
remained in the TJ plane still exhibited some co-localization
(Fig. 7i).
|
CNF-1 induces polarized reorganization of F-actin and select
actin-binding proteins
Since TJs have previously been shown to affiliate with F-actin
(Madara, 1987;
Madara et al., 1987
;
Madara et al., 1988
), and
since actin-myosin contraction is an important regulator of paracellular
permeability (Turner and Madara,
1995
), we examined in detail the effects of CNF-1 treatment on the
organization of polarized pools of F-actin and associated binding proteins in
epithelial cells. F-actin architecture in T84 monolayers was highlighted by
rhodamine-phalloidin staining and confocal microscopy following 6 hours
exposure to either CNF-1 (1 nM) or vehicle
(Fig. 8a-1).
Fig. 8a represents en face
imaging at the level of the apical membrane. F-actin in this plane is
organized in characteristic perijunctional rings (arrowhead) and, in the
apical brush border, as fine dot-like staining representing microvillous actin
cores (*). This pattern was unaltered following apical incubation
with CNF-1 toxin (Fig. 8b). While perijunctional F-actin ring staining was preserved (or perhaps even
enhanced) in response to basolateral treatment with CNF-1
(Fig. 8c), a striking abolition
of microvillous actin staining was observed (arrow). This was complemented by
alterations in the staining patterns of villin, a major actin-regulating
protein enriched in apical epithelial brush borders. In control monolayers
(Fig. 8d) and those treated
apically with CNF-1 (Fig. 8e),
en face images revealed identical villin staining patterns in F-actin-rich
microvillous cores. Villin staining intensity in the apical plane was
significantly diminished following basolateral treatment with CNF-1
(Fig. 8f), suggesting brush
border effacement.
|
At the basal pole of both control (Fig.
8g) and apically treated (Fig.
8h) monolayers, F-actin stress fibers were observed as a meshwork
of filaments. In contrast to the effacement of brush border F-actin,
basolateral exposure to CNF-1 (Fig.
8i) induced aggregation of stress fibers into prominent bundles
with a `cabled' appearance. Enhancements in both stress fiber number and
aggregation have been previously observed in non-polarized cells such as
fibroblasts transfected with dominant active RhoA
(Ridley and Hall, 1992). This
is a hallmark feature confirming the specific activation of RhoA. Since stress
fiber changes were observed following CNF-1 incubation, we also examined the
distribution of an actin-binding protein in the same region of the cells. In
control monolayers, the actin-binding protein paxillin was seen as small
plaques consistent with its distribution in focal cell-matrix contacts
(Fig. 8j). Monolayers treated
apically with CNF-1 displayed a similar staining pattern for paxillin
(Fig. 8k). However, basolateral
incubation with CNF-1 (Fig. 8l)
induced a small increase in paxillin plaques. All morphological alterations in
F-actin/actin-binding proteins induced by CNF-1 occurred independently of
biochemical alterations in protein levels at all sampled time points, as shown
by western blot data (Fig.
8m-o). Actin levels in control
(Fig. 8m, lanes 1-3) versus
CNF-treated (lanes 4-6) lysates appeared similar, as did villin levels under
the same conditions (Fig. 8n,
lanes 1-3 control, lanes 4-6 CNF-1). A slight increase in the molecular mass
of paxillin was noticed in lysates from CNF-treated monolayers at all time
points shown (Fig. 8o, lanes
1-3 control, lanes 4-6 CNF-1). When lysates from control and CNF-treated (1
nM, 24 hours) monolayers were blotted with an antibody specific for
tyrosine-phosphorylated paxillin (Fig.
8p), a substantial increase in phospho-paxillin was detected in
CNF-treated samples. We also observed increased tyrosine phosphorylation of
paxillin in CNF-treated cells following immunoprecipitation with anti-paxillin
and immunoblotting with anti-phosphopaxillin (data not shown). Tyrosine
phosphorylation of paxillin has previously been observed in other systems
following Rho GTPase activation (Imamura
et al., 2000
; Sinnett-Smith et
al., 2001
).
CNF-1 impairs intercellular junction assembly
Transient depletion of extracellular calcium is a well-validated model for
studying the assembly of intercellular junctional contacts. Since CNF-1
influenced TJs in stable polarized epithelial cells, we examined the influence
of Rho GTPase activation by CNF-1 on assembly of intercellular contacts and
recovery of barrier function. As shown
(Fig. 9A), the baseline TER of
monolayers subsequently incubated with vehicle (closed circles) or CNF-1 (open
circles) were similarly high before calcium depletion (#). Following EGTA
treatment, TER predictably dropped to below 100 .cm2,
whereupon monolayers were washed and allowed to recover in calcium-containing
media in the presence or absence of CNF-1 (1 nM). Control monolayers recovered
high TER values of
700
.cm2 by 6 hours following
repletion of extracellular calcium. This corresponds to the presence of
assembled TJs in the monolayer (Liu et
al., 2000
). By contrast, monolayers exposed to CNF-1 during the
recovery period failed to recover TER (162±14
.cm2 24
hours after EGTA washout and calcium repletion). To answer the question of
whether this related to inhibitory effects of CNF-1 on nascent AJs as well as
TJs, we examined E-cadherin and occludin distribution in monolayers after EGTA
treatment and during recovery in the presence of CNF-1 or vehicle. Following
EGTA treatment (Fig. 9B), both
occludin (a) and E-cadherin (b) were localized intracellularly rather than at
their respective junctions. Redistribution of both these proteins back to the
lateral membrane was evident in control cells 9 hours later (c,d). However,
occludin did not correctly redistribute back to the membrane in monolayers
exposed to CNF-1 during recovery (e). It was instead observed in punctate dots
that may mark tricellular corners. E-cadherin movement back to the membrane in
CNF-treated cells (f) resembled that in control cells, further indicating
preferential disruption of the TJ over the AJ by CNF-1. These findings
illustrate that overactivation of Rho GTPases impairs not only the barrier
function of mature TJs, but is also detrimental for the assembly of nascent
TJs and the acquisition of barrier properties.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The functional consequences of Rho, Rac and Cdc42 activation by CNF-1 in
our model included the induction of parallel reductions in TER and
enhancements in paracellular permeability. This response was polarized to
basolateral incubation with CNF-1. The C terminus of CNF reportedly harbors
the cell-binding domain (Lemichez et al.,
1997), mediating clathrin-independent endocytosis requiring
transient membrane acidification (Contamin
et al., 2000
).
Reported effects of the activation of Rho GTPases by CNF-1 on epithelial
TER have to date been ambiguous. Interference with signaling could cause: (1)
increased TER; (2) decreased TER; or (3) no change in TER. One report suggests
that CNF-1 does not affect TER (Hofman et
al., 1998), however our results support observations of
CNF-induced reductions in TER in another well-polarized epithelial cell line,
Caco-2 (Gerhard et al., 1998
).
Since CNF-1 simultaneously activates Rho, Rac and Cdc42, it is difficult in
this model to dissect out the specific contributions of individual GTPases to
observed changes in barrier function. However, some information can be gained
by using specific pharmacological inhibitors of the RhoA pathway in
combination with CNF-1. We used the RhoA inhibitor C3 toxin to examine whether
inhibition of RhoA in T84 cells could protect against the barrier-disruptive
effects of CNF-1 (data not shown). Pre-incubation of the cells with low
concentrations of C3 caused a reproducible drop in TER, which was not
prevented by application of CNF-1 (data not shown). Similarly, simultaneous
incubation of cells with CNF-1 and C3 evoked profound disruptions in TER (data
not shown). We also tried to block the effects of CNF-1 on epithelial TER
using the Rho kinase inhibitor Y-27632. However, since this by itself also
compromises epithelial barrier function
(Walsh et al., 2001
), we did
not observe any protective effect against the influence of CNF-1 (data not
shown). Thus, in our model, the effects of CNF-1 were not blocked by
inhibitors of RhoA or Rho kinase, verifying that RhoA is not the sole factor
responsible for reductions in TER and barrier function. Given that all
compounds (CNF-1, C3, Y-27632) adversely affect the barrier function of
epithelial cells, this supports the notion that a delicate balance of Rho
activity/quiescence must be maintained in order to preserve optimal barrier
function.
A more straightforward approach towards understanding the contributions of
individual Rho GTPases to epithelial barrier function involves the
over-expression of constitutively active/dominant-negative mutants of each
GTPase. Stable expression of dominant-active Rho reportedly increases renal
epithelial TER (Hasegawa et al.,
1999). It is possible that dominant-active Rho `swamps' a cell
with signaling GTPase, potentially engaging effectors not normally activated
physiologically, and thereby eliciting unusual TER phenomena. Additionally, in
low-resistance monolayers such as those used in the study in question
(Hasegawa et al., 1999
), TER
is a less-sensitive index of barrier function than the assessment of
paracellular flux. Since paracellular permeability increased during
dominant-active Rho expression (Hasegawa
et al., 1999
), this corroborates our results indicating
compromised epithelial barrier function during Rho GTPase activation.
Inducible expression systems that allow precise manipulation of mutant
expression levels are a further improvement on stable expression systems where
the mutant transgene is continuously `on'. Using an elegant system, convincing
evidence for an involvement of RhoA in disruption of epithelial barrier
function has been obtained with tetracycline-repressible dominant-active Rho
transfected into MDCK cells (Jou et al.,
1998
). In this system, dominant-active Rho mutant induction
correlated with inability to form high TER
(Jou et al., 1998
). Thus, data
by Jou et al., 1998
complements our use of CNF-1 to manipulate pharmacologically the activity of
Rho GTPases (Jou et al.,
1998
). Additionally, our study attempted to characterize further
the likely reasons for the disturbance of epithelial barrier function, in the
context of alterations in TJ proteins and F-actin organization.
Several lines of evidence suggest that CNF-induced perturbations of barrier
function are non-cytotoxic (data not shown). First, TER values did not drop
below 300
.cm2, discounting frank cytotoxicity. Second,
cell death was not observed in CNF-treated monolayers stained with
hematoxylin/eosin. Finally, mitochondrial bioreduction of a tetrazolium salt
was unimpaired in CNF-treated monolayers relative to control monolayers.
Additionally, it has been reported that CNF-1 might in fact protect against
apoptosis by enhancing cell-matrix adherence and cell spreading
(Fiorentini et al., 1997
;
Fiorentini et al., 1998b
),
possibly related to enhanced F-actin stress fiber aggregation mediated by
activated Rho (Ridley and Hall,
1992
). There is also evidence that CNF-1 exerts an additional
anti-apoptotic influence by increasing the expression of proteins in the Bcl-2
family (Fiorentini et al.,
1998c
).
To explore reasons for epithelial permeability defects induced upon
activation of Rho GTPases, we began by examining the status of key proteins
that localize in the TJ and exert regulatory control over the paracellular
pathway. Significant redistribution of TJ proteins occludin, ZO-1 and JAM was
observed following basolateral treatment with CNF-1. Similar effects were also
observed for claudin-1 (data not shown). Occludin and ZO-1 immunolocalization
in characteristic TJ `ring structures' was severely disrupted, resulting in
almost complete lack of continuity. JAM localization too was disturbed but,
unlike occludin and ZO-1, did not virtually `disappear' from the plane of the
TJ membrane in response to basolateral treatment with CNF-1. Instead, broad,
blurred rings of JAM were evident in en face confocal images, indicating some
JAM redistribution below the TJ membrane in contrast to its former presence in
a sharply localized ring at the TJ itself. It is intriguing to speculate why
the disruption of JAM is not as drastic as that of occludin and ZO-1. This
might relate to a necessity for JAM localization at the TJ membrane for
correct assembly of the TJ protein complex
(Liu et al., 2000), otherwise
loss of this protein could prove detrimental for the capability of T84
monolayers to reestablish barrier function after transient insult by toxins
such as CNF-1.
Our observations extend reports of irregular ZO-1 staining in MDCK cells
transiently induced to express dominant-active RhoA
(Jou et al., 1998;
Hasegawa et al., 1999
).
However, continuous expression of the same construct reportedly does not
affect (Takaishi et al., 1997
)
or even enhances (Gopalakrishnan et al.,
1998
) ZO-1 localization at MDCK cell contact sites. The
limitations of inducible versus continuous expression systems have been
mentioned regarding inconsistencies between TER/permeability data. ZO-1
localization was reportedly unaffected in T84 cells exposed to CNF-1 for 10-16
hours (Hofman et al., 1998
).
However, control ZO-1 staining in this model was uncharacteristically diffuse,
probably representing antibody specificity issues. Thus, nonspecific ZO-1
staining could obscure the detection of differences between control and
CNF-treated monolayers. Our studies provide evidence that many proteins move
away from the TJ upon overactivation of Rho GTPases, potentially explaining
CNF-induced deficits in epithelial barrier function. We have also observed
similar effects using an alternative intestinal epithelial model, the Caco-2
cell line, which is also a highly validated model for the study of TJs and
barrier function.
In an attempt to explain why dramatic CNF-induced reductions in TJ protein immunostaining were not accompanied by biochemical loss of protein, we performed immunogold electron microscopy on T84 monolayers in order to define better the precise localization of TJ proteins following CNF-1 treatment. In control cells treated only with vehicle, occludin labeling was confined to areas of cell-cell contact, mainly in the uppermost 250 nm of the basolateral plasma membrane. Following exposure to CNF-1, some occludin remained at this site but a significant amount was internalized in endosomal/caveolar-like membranous structures, evidenced by co-localization of TJ proteins with caveolin-1 (also significantly internalized away from the TJ). Similar trends were observed for ZO-1 internalization and co-localization with caveolin-1 after CNF-1 treatment (data not shown). This is the first demonstration of TJ protein internalization in such structures following constitutive activation of Rho GTPases, and probably explains why profound reductions in TJ protein immunostaining were not accompanied by corresponding reductions in total TJ protein levels.
In order better to address the cellular destination of TJ proteins
internalized following incubation with CNF-1, we double-stained monolayers for
internalized occludin and a range of caveolar/endosomal markers. Strong
co-localization was observed with the caveolar marker caveolin-1. A sub-pool
of internalized occludin also co-localized with EEA-1-positive early endosomes
and Rab11-positive recycling endosomes, but did not co-localize with late
endosomes/lysosomes marked by LAMP-1. This provides novel evidence that TJ
proteins can be internalized into endosomal compartments after toxin
treatment. Dominant-active Rac1 and Cdc42 have been shown to affect endocytic
trafficking in epithelial cells (Jou et
al., 2000; Rojas et al.,
2001
), but little is known about the fate of TJ proteins following
insult to the epithelium. Internalization of TJ proteins in caveolae or
recycling endosomes is likely to have important functional consequences for
cell survival. Proteins internalized in this process (sometimes termed
potocytosis) appear to be protected from lysosomal degradation pathways unlike
proteins internalized in classical clathrin-coated pits
(Anderson, 1998
;
Mineo and Anderson, 2001
).
Thus, effective recycling of TJ proteins back to the membrane could occur in
the absence of new protein synthesis, allowing faster re-establishment of
barrier function. Since it is not clear exactly how much of each protein must
remain at the TJ in order to maintain adequate barrier function, it must be
speculated that any loss could adversely affect barrier properties to the
extent observed in our functional assays following CNF-1 treatment.
Interestingly, we have previously documented similar intracellular
sequestration of TJ proteins following Rho protein inhibition with C.
difficile toxins (Nusrat et al.,
2001
). Taken together, this illustrates that disruption of Rho
GTPase function, whether excessive stimulation or indeed inhibition, adversely
affects TJ structure and inevitably function.
Another important component of the TJ functional unit relates to its
affiliations with cellular actin-myosin contractile machinery in the
perijunctional F-actin ring. Inhibitors of MLC kinase have been shown to
inhibit increases in paracellular permeability induced upon activation of the
sodium-glucose transporter SGLT-1, which is physiologically important for
enhancing nutrient uptake (Turner and
Madara, 1995). Thus, the localization and phosphorylation status
of MLC is likely to play an important role in governing paracellular
permeability. In our study, basolateral treatment with CNF-1 significantly
attenuated the presence of a pool of p-MLC co-localizing with ZO-1 at the TJ.
However, CNF-induced enhancements in paracellular permeability could not be
prevented using the MLC kinase inhibitor ML-7 (data not shown). This might
reflect a two-way signaling mechanism, where CNF-1 mediates its effects by
influencing TJ proteins that in turn modulate the underlying F-actin
cytoskeleton. By contrast, certain physiological signals such as
sodium-glucose cotransport act primarily by influencing F-actin/MLC, which
also affects TJ function. Thus, phosphorylation of MLC could be more important
as a physiological regulator of paracellular permeability
(Turner and Madara, 1995
) than
as a pathophysiological target of bacterial toxins such as CNF-1.
Additionally, since the presence of p-MLC at the TJ is crucial for contraction
of the perijunctional F-actin ring, this could suggest that the primary
permeability deficits induced by CNF-1 are not mediated by alterations in
perijunctional ring contraction, but rather by another mechanism that might be
related. This may prove an interesting area for further study, in the context
of differences in the regulation of paracellular permeability between
physiological stimuli such as glucose uptake and pathophysiological stimuli
such as bacterial toxins.
In comparison to the profound effects of CNF-1 at the epithelial TJ, an
interesting feature of our model is that the underlying adherens junction (AJ)
was minimally disrupted. Some thickening or diffusion of E-cadherin and
ß-catenin was observed at cell-cell borders; however, the characteristic
`ring structure' patterns of both these proteins was largely preserved.
Relative selectivity for TJs over AJs has also been observed upon direct
inhibition of RhoA (Nusrat et al.,
1995; Nusrat et al.,
2001
). Adhesive control at the AJ probably relates to Rac over Rho
activity, since dominant-active Rac increased E-cadherin and ß-catenin
protein levels in MDCK cells (Takaishi et
al., 1997
). Therefore, the subtle alterations observed in AJ
protein localization in our model might reflect Rac activation by CNF-1.
However, CNF-induced morphological changes are most striking at the level of
the TJ, a site where Rho exerts major structural and functional influence
(Gopalakrishnan et al., 1998
;
Hirase et al., 2001
;
Jou et al., 1998
;
Nusrat et al., 1995
;
Nusrat et al., 2001
;
Walsh et al., 2001
). In light
of the ability of CNF-1 to induce simultaneous activation of Rho, Rac and
Cdc42, cross-talk between these three GTPases is likely to play a major role
in the complex and intricate regulation of barrier function in epithelial
cells.
Since the TJ has an intimate association with the F-actin cytoskeleton
(Madara, 1987;
Madara et al., 1987
;
Madara et al., 1988
), we
sought to explore further the role of Rho GTPase activation in regulating
F-actin structures in polarized epithelial cells. Perijunctional F-actin was
minimally affected in our system upon treatment with CNF-1, although slight
increases in F-actin staining intensity at cell-cell borders were observed.
This corroborates a report that CNF-1 stimulates F-actin accumulation at HEp-2
cell borders (Fiorentini et al.,
1988
) and might in fact reflect Rac rather than Rho activity,
since dominant-active Rac stimulated F-actin accumulation at MDCK junctions
(Takaishi et al., 1997
). The
most striking feature of basolateral exposure to CNF-1 was the almost complete
loss of apical (microvillous) F-actin and its binding protein villin. Thus, as
suggested, CNF-1 might efface apical microvilli
(Hofman et al., 1998
), but
also disrupt the continuity between adjacent F-actin pools in the microvilli,
perijunctional ring and the terminal web that could potentially destabilize TJ
complex `scaffolding'. However, as discussed in the context of reduced p-MLC
localization at the TJ following incubation with CNF-1, contraction-based
events might be secondary to mislocalization of TJ proteins as a mechanism for
disrupting barrier function.
An interesting feature of the effects of CNF-1 on our model was polarized
restructuring of F-actin. Thus, while apical F-actin was diminished, we
observed prominent F-actin cables at the level of basal stress fibers, with
some reorganization of paxillin, a focal adhesion protein that links actin to
the extracellular matrix (Turner,
2000). Increased stress fiber density has been reported upon
exposure to CNF-1 (Gerhard et al.,
1998
; Hofman et al.,
1998
) or activated Rho
(Hasegawa et al., 1999
;
Takaishi et al., 1997
). This
mimics Rho-stimulated stress fiber formation in mesenchymal cells
(Ridley and Hall, 1992
).
Increased basal F-actin might protect against injury to both epithelial
(Barth et al., 1999
) and
endothelial (Vouret-Craviari et al.,
1999
) monolayers. Our findings represent restructuring of F-actin
cytoskeletal structures, since total cellular levels of actin, villin and
paxillin did not change in control versus CNF-treated monolayers. However,
increased levels of tyrosine-phosphorylated paxillin were detected in
CNF-treated samples relative to controls. This complements existing data
showing enhanced tyrosine phosphorylation of paxillin in hepatoma cells
treated with lysophosphatidic acid (LPA) by a RhoA/Rho kinase pathway
(Imamura et al., 2000
).
Inhibition of Rho kinase has also been shown to prevent increases in tyrosine
phosphorylation of paxillin observed upon bombesin treatment of Swiss 3T3
cells (Sinnett-Smith et al.,
2001
). However, in our system, the tyrosine kinase inhibitor
genistein did not reverse any of the functional effects of CNF-1 such as
reductions in TER or induced rearrangements in TJ protein localization (data
not shown). Thus, tyrosine phosphorylation of paxillin upon Rho activation
probably reflects polarized regulatory control of the basal F-actin network at
the cell-matrix interface.
Rho GTPase regulation of TJ/AJ assembly was also analyzed using calcium
switch assays (Liu et al.,
2000). CNF-1 severely impaired TER recovery after transient
calcium depletion. Recovery was similarly impaired in a related system, MDCK
cells exposed to the ligand for a G-protein-coupled receptor activating RhoA
(Hasegawa et al., 1999
).
Conflicting evidence was obtained by the same group using dominant-active RhoA
transfectants, suggesting utilization of additional Rho effectors or perhaps
negative regulation after swamping cells with actively signaling GTPase. In
our system, CNF-induced impairment of recovery following calcium switch
related to a direct effect on nascent TJs rather than AJs. Thus, we observed
re-assembly of E-cadherin into ring-structures at the level of the AJ, whereas
movement of occludin back to the TJ membrane was severely impaired in
CNF-treated cells. Taken together with evidence of severe perturbations in TJs
of confluent cells with mature junctions, this further illustrates the
selectivity of CNF-1 for the TJ over the AJ.
While CNF-1 constitutively activates all Rho family members (Rho, Rac and
Cdc42) (Lerm et al., 1999b),
stress fiber assembly observed in our cells is highly consistent with RhoA
activation (Ridley and Hall,
1992
). Permeability alterations seem compatible with enhanced RhoA
activity as reported to date, based on pharmacological studies and
dominant-active RhoA mutants. However, it is not possible from our study to
estimate the relative contributions of each activated GTPase to the functional
changes observed. The low levels of RhoA activation observed in our assays in
response to CNF-1 treatment may be sufficient to alter barrier function,
particularly if Rho is concentrated in a highly enriched microenvironment at
the cell surface as has been described previously
(Michaely et al., 1999
). High
levels of Rac or Cdc42 activation might also be important in altering barrier
function or, alternatively, other complementary cellular processes not
directly investigated here. Additionally, cooperation between all three
GTPases cannot be discounted as a mechanism for CNF-induced alterations in
barrier function, since Cdc42 activation in fibroblasts has been reported to
initiate sequential activation of Rac and Rho
(Nobes and Hall, 1995
).
Examination of specific effectors involved in Rho GTPase regulation of
permeability is outside the scope of this study. However, multiple targets are
potentially involved (Aspenstrom,
1999). Recent evidence suggests a role for Rho kinase (p160ROCK),
which regulates actin-myosin contractility through multiple effects on myosin
and its regulatory enzymes (Amano et al.,
1996
; Fukata et al.,
1998
; Kimura et al.,
1996
). Inhibition of Rho-induced permeability enhancements in
endothelial/epithelial cells was recently shown with both dominant-negative
Rho and a Rho kinase inhibitor (Hirase et
al., 2001
). Recent evidence from our laboratory has also
implicated Rho kinase in regulating epithelial barrier function
(Walsh et al., 2001
). However,
the Rho kinase inhibitor Y-27632 did not block the effects of CNF-1 on TER or
TJ protein immunolocalization in our model (data not shown). This does not
rule out the possibility that Rho kinase is involved, but merely underlines
the difficulties in interpreting data obtained with two compounds both of
which decrease barrier function.
In conclusion, our studies have demonstrated compromised intestinal
epithelial barrier function upon constitutive activation of Rho GTPases with
E. coli CNF-1. We have presented novel evidence that barrier deficits
are probably accounted for by movement of occludin and ZO-1 away from the TJ
into membranous structures resembling caveolae/endosomes. Internalized TJ
proteins might cycle through caveolin-containing rafts, and early and
recycling endosomes, but appear to evade a degradative pathway involving late
endosomes/lysosomes. Reports of TJ protein internalization in epithelial cells
treated with clostridial toxins to inhibit Rho activity
(Nusrat et al., 2001)
illustrate the complexity of Rho GTPase-mediated regulation of TJ
structure/function. Epithelial cells in vivo probably exert careful control
over levels of activity/quiescence of Rho GTPase family members in order to
preserve barrier function. While the role of CNF-1 as a virulence factor is
controversial, it is possible that, by decreasing epithelial barrier function,
the toxin facilitates access of luminal bacteria and proven virulence factors
(such as hemolysin) to the sub-epithelial compartment. This could account for
clinical correlations between the detection of CNF-1 and other virulence
factors in diverse diseases (Andreu et al.,
1997
; Blanco et al.,
1995
; Caprioli et al.,
1987
; Elliott et al.,
1998
; Yuri et al.,
1998
). Furthermore, our study clearly supports a role of Rho
GTPases in not only the maintenance of established TJs and paracellular
permeability, but also the assembly of intercellular associations and recovery
of barrier function.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Aktories, K. (1997). Bacterial toxins that
target Rho proteins. J. Clin. Invest.
99,827
-829.
Amano, M., Ito, M., Kimura, K., Fukata, Y., Chihara, K., Nakano,
T., Matsuura, Y. and Kaibuchi, K. (1996). Phosphorylation and
activation of myosin by Rho-associated kinase (Rho-kinase). J.
Biol. Chem. 271,20246
-20249.
Anderson, R. G. (1998). The caveolae membrane system. Annu Rev. Biochem. 67,199 -225.[CrossRef][Medline]
Andreu, A., Stapleton, A. E., Fennell, C., Lockman, H. A., Xercavins, M., Fernandez, F. and Stamm, W. E. (1997). Urovirulence determinants in Escherichia coli strains causing prostatitis [published erratum appears in J. Infect Dis. 1997 Nov;176(5):1416]. J. Infect. Dise. 176,464 -469.
Aspenstrom, P. (1999). Effectors for the Rho GTPases. Curr. Opin. Cell Biol. 11, 95-102.[CrossRef][Medline]
Barth, H., Olenik, C., Sehr, P., Schmidt, G., Aktories, K. and
Meyer, D. K. (1999). Neosynthesis and activation of Rho by
Escherichia coli cytotoxic necrotizing factor (CNF1) reverse
cytopathic effects of ADP-ribosylated Rho. J. Biol.
Chem. 274,27407
-27414.
Ben-Ami, G., Ozeri, V., Hanski, E., Hofmann, F., Aktories, K.,
Hahn, K. M., Bokoch, G. M. and Rosenshine, I. (1998). Agents
that inhibit Rho, Rac, and Cdc42 do not block formation of actin pedestals in
HeLa cells infected with enteropathogenic Escherichia coli. Infect.
Immun. 66,1755
-1758.
Blanco, M., Blanco, J., Blanco, J. E., Alonso, M. P., Abalia, I., Rodriguez, E., Bilbao, J. R. and Umaran, A. (1995). [Virulence factors and 0 serogroups of Escherichia coli as a cause of community-acquired urinary infections]. Enferm. Infecc. Microbiol. Clin. 13,236 -241 [Spanish].[Medline]
Boquet, P. (1999). Bacterial toxins inhibiting
or activating small GTP-binding proteins. Ann. N. Y. Acad.
Sci. 886,83
-90.
Boquet, P., Sansonetti, P. J. and Tran Van Nhieu, G. (1999). Rho GTP-binding proteins as targets for microbial pathogens. Prog. Mol. Subcell. Biol. 22,183 -199.[Medline]
Caprioli, A., Falbo, V., Roda, L. G., Ruggeri, F. M. and Zona, C. (1983). Partial purification and characterization of an Escherichia coli toxic factor that induces morphological cell alterations. Infect. Immun. 39,1300 -1306.[Medline]
Caprioli, A., Donelli, G., Falbo, V., Possenti, R., Roda, L. G., Roscetti, G. and Ruggeri, F. M. (1984). A cell division-active protein from E. coli. Biochem. Biophys. Res. Commun. 118,587 -593.[Medline]
Caprioli, A., Falbo, V., Ruggeri, F. M., Baldassarri, L., Bisicchia, R., Ippolito, G., Romoli, E. and Donelli, G. (1987). Cytotoxic necrotizing factor production by hemolytic strains of Escherichia coli causing extraintestinal infections. J. Clin. Microbiol. 25,146 -149.[Medline]
Casanova, J. E., Wang, X., Kumar, R., Bhartur, S. G., Navarre,
J., Woodrum, J. E., Altschuler, Y., Ray, G. S. and Goldenring, J. R.
(1999). Association of Rab25 and Rab11a with the apical recycling
system of polarized Madin-Darby canine kidney cells. Mol. Biol.
Cell 10,47
-61.
Contamin, S., Galmiche, A., Doye, A., Flatau, G., Benmerah, A.
and Boquet, P. (2000). The p21 Rho-activating toxin cytotoxic
necrotizing factor 1 is endocytosed by a clathrin-independent mechanism and
enters the cytosol by an acidic-dependent membrane translocation step.
Mol. Biol. Cell 11,1775
-1187.
De Rycke, J., Nougayrede, J. P., Oswald, E. and Mazars, P. (1997). Interaction of Escherichia coli producing cytotoxic necrotizing factor with HeLa epithelial cells. Adv. Exp. Med. Biol. 412,363 -366.[Medline]
Denker, B. M. and Nigam, S. K. (1998). Molecular structure and assembly of the tight junction. Am. J. Physiol. 274,F1 -9.[Medline]
Dharmsathaphorn, K. and Madara, J. L. (1990). Established intestinal cell lines as model systems for electrolyte transport studies. Methods Enzymol. 192,354 -389.[Medline]
Dharmsathaphorn, K., McRoberts, J. A., Mandel, K. G., Tisdale,
L. D. and Masui, H. (1984). A human colonic tumor cell line
that maintains vectorial electrolyte transport. Am. J.
Physiol. 246,G204
-208.
Elliott, S. J., Srinivas, S., Albert, M. J., Alam, K.,
Robins-Browne, R. M., Gunzburg, S. T., Mee, B. J. and Chang, B. J.
(1998). Characterization of the roles of hemolysin and other
toxins in enteropathy caused by alpha-hemolytic Escherichia coli
linked to human diarrhea. Infect. Immun.
66,2040
-2051.
Falzano, L., Fiorentini, C., Donelli, G., Michel, E., Kocks, C., Cossart, P., Cabanie, L., Oswald, E. and Boquet, P. (1993). Induction of phagocytic behaviour in human epithelial cells by Escherichia coli cytotoxic necrotizing factor type 1. Mol. Microbiol. 9,1247 -1254.[Medline]
Fiorentini, C., Arancia, G., Caprioli, A., Falbo, V., Ruggeri, F. M. and Donelli, G. (1988). Cytoskeletal changes induced in HEp-2 cells by the cytotoxic necrotizing factor of Escherichia coli.Toxicon 26,1047 -1056.[Medline]
Fiorentini, C., Fabbri, A., Matarrese, P., Falzano, L., Boquet, P. and Malorni, W. (1997). Hinderance of apoptosis and phagocytic behaviour induced by Escherichia coli cytotoxic necrotizing factor 1: two related activities in epithelial cells. Biochem. Biophys. Res. Commun. 241,341 -346.[CrossRef][Medline]
Fiorentini, C., Gauthier, M., Donelli, G. and Boquet, P. (1998a). Bacterial toxins and the Rho GTP-binding protein: what microbes teach us about cell regulation. Cell Death Differ. 5,720 -728.[CrossRef][Medline]
Fiorentini, C., Matarrese, P., Straface, E., Falzano, L., Donelli, G., Boquet, P. and Malorni, W. (1998b). Rho-dependent cell spreading activated by E. coli cytotoxic necrotizing factor 1 hinders apoptosis in epithelial cells. Cell Death Differ. 5,921 -929.[CrossRef][Medline]
Fiorentini, C., Matarrese, P., Straface, E., Falzano, L., Fabbri, A., Donelli, G., Cossarizza, A., Boquet, P. and Malorni, W. (1998c). Toxin-induced activation of Rho GTP-binding protein increases Bcl-2 expression and influences mitochondrial homeostasis. Exp. Cell Res. 242,341 -350.[CrossRef][Medline]
Flatau, G., Lemichez, E., Gauthier, M., Chardin, P., Paris, S., Fiorentini, C. and Boquet, P. (1997). Toxin-induced activation of the G protein p21 Rho by deamidation of glutamine. Nature 387,729 -733.[CrossRef][Medline]
Fukata, Y., Kimura, K., Oshiro, N., Saya, H., Matsuura, Y. and
Kaibuchi, K. (1998). Association of the myosin-binding
subunit of myosin phosphatase and moesin: dual regulation of moesin
phosphorylation by Rho-associated kinase and myosin phosphatase. J.
Cell Biol. 141,409
-418.
Gerhard, R., Schmidt, G., Hofmann, F. and Aktories, K.
(1998). Activation of Rho GTPases by Escherichia coli
cytotoxic necrotizing factor 1 increases intestinal permeability in Caco-2
cells. Infect. Immun.
66,5125
-5131.
Gopalakrishnan, S., Raman, N., Atkinson, S. J. and Marrs, J. A. (1998). Rho GTPase signaling regulates tight junction assembly and protects tight junctions during ATP depletion. Am. J. Physiol. 275,C798 -809.[Medline]
Gyles, C. L. (1992). Escherichia coli cytotoxins and enterotoxins. Can. J. Microbiol. 38,734 -746.[Medline]
Hall, A. (1998). Rho GTPases and the actin
cytoskeleton. Science
279,509
-514.
Hall, A. (1990). The cellular functions of small GTP-binding proteins. Science 249,635 -640.[Medline]
Hasegawa, H., Fujita, H., Katoh, H., Aoki, J., Nakamura, K.,
Ichikawa, A. and Negishi, M. (1999). Opposite regulation of
transepithelial electrical resistance and paracellular permeability by Rho in
Madin-Darby canine kidney cells. J. Biol. Chem.
274,20982
-20988.
Hirase, T., Kawashima, S., Wong, E. Y., Ueyama, T., Rikitake,
Y., Tsukita, S., Yokoyama, M. and Staddon, J. M. (2001).
Regulation of tight junction permeability and occludin phosphorylation by
RhoA-p160ROCK-dependent and -independent mechanisms. J. Biol.
Chem. 276,10423
-10431.
Hofman, P., Flatau, G., Selva, E., Gauthier, M., le Negrate, G.,
Fiorentini, C., Rossi, B. and Boquet, P. (1998).
Escherichia coli cytotoxic necrotizing factor 1 effaces microvilli
and decreases transmigration of polymorphonuclear leukocytes in intestinal T84
epithelial cell monolayers. Infect. Immun.
66,2494
-2500.
Hopkins, A. M., Li, D., Mrsny, R. J., Walsh, S. V. and Nusrat, A. (2000). Modulation of tight junction function by G protein-coupled events. Adv. Drug Deliv. Rev. 41,329 -340.[CrossRef][Medline]
Imamura, F., Mukai, M., Ayaki, M. and Akedo, H. (2000). Y-27632, an inhibitor of rho-associated protein kinase, suppresses tumor cell invasion via regulation of focal adhesion and focal adhesion kinase. Jpn J. Cancer Res. 91,811 -816.[Medline]
Island, M. D., Cui, X., Foxman, B., Marrs, C. F., Stamm, W. E.,
Stapleton, A. E. and Warren, J. W. (1998). Cytotoxicity of
hemolytic, cytotoxic necrotizing factor 1-positive and -negative
Escherichia coli to human T24 bladder cells. Infect.
Immun. 66,3384
-3389.
Island, M. D., Cui, X. and Warren, J. W.
(1999). Effect of Escherichia coli cytotoxic necrotizing
factor 1 on repair of human bladder cell monolayers in vitro.
Infect. Immun. 67,3657
-3661.
Jou, T. S. and Nelson, W. J. (1998). Effects of
regulated expression of mutant RhoA and Rac1 small GTPases on the development
of epithelial (MDCK) cell polarity. J. Cell Biol.
142,85
-100.
Jou, T. S., Schneeberger, E. E. and Nelson, W. J.
(1998). Structural and functional regulation of tight junctions
by RhoA and Rac1 small GTPases. J. Cell Biol.
142,101
-115.
Jou, T. S., Leung, S. M., Fung, L. M., Ruiz, W. G., Nelson, W.
J. and Apodaca, G. (2000). Selective alterations in
biosynthetic and endocytic protein traffic in Madin-Darby canine kidney
epithelial cells expressing mutants of the small GTPase Rac1. Mol.
Biol. Cell 11,287
-304.
Kazmierczak, B. I., Jou, T. S., Mostov, K. and Engel, J. N. (2001). Rho GTPase activity modulates Pseudomonas aeruginosa internalization by epithelial cells. Cell. Microbiol. 3,85 -98.[CrossRef][Medline]
Kimura, K., Ito, M., Amano, M., Chihara, K., Fukata, Y., Nakafuku, M., Yamamori, B., Feng, J., Nakano, T., Okawa, K. et al. (1996). Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273,245 -248.[Abstract]
Kranenburg, O., Poland, M., van Horck, F. P., Drechsel, D.,
Hall, A. and Moolenaar, W. H. (1999). Activation of RhoA by
lysophosphatidic acid and Galpha 12/13 subunits in neuronal cells: induction
of neurite retraction. Mol. Biol. Cell
10,1851
-1857.
Lemichez, E., Flatau, G., Bruzzone, M., Boquet, P. and Gauthier, M. (1997). Molecular localization of the Escherichia coli cytotoxic necrotizing factor CNF1 cell-binding and catalytic domains. Mol. Microbiol. 24,1061 -1070.[CrossRef][Medline]
Lerm, M., Schmidt, G., Goehring, U. M., Schirmer, J. and
Aktories, K. (1999a). Identification of the region of rho
involved in substrate recognition by Escherichia coli cytotoxic
necrotizing factor 1 (CNF1). J. Biol. Chem.
274,28999
-29004.
Lerm, M., Selzer, J., Hoffmeyer, A., Rapp, U. R., Aktories, K.
and Schmidt, G. (1999b). Deamidation of Cdc42 and Rac by
Escherichia coli cytotoxic necrotizing factor 1: activation of c-Jun
N-terminal kinase in HeLa cells. Infect. Immun.
67,496
-503.
Lerm, M., Schmidt, G. and Aktories, K. (2000). Bacterial protein toxins targeting Rho GTPases. FEMS Microbiol. Lett. 188,1 -6.[CrossRef][Medline]
Lesser, C. F., Scherer, C. A. and Miller, S. I. (2000). Rac, ruffle and rho: orchestration of Salmonella invasion. Trends Microbiol. 8, 151-152.[CrossRef][Medline]
Liu, Y., Nusrat, A., Schnell, F. J., Reaves, T. A., Walsh, S.
V., Pochet, M. P. and Parkos, C. A. (2000). Human junction
adhesion molecule regulates tight junction resealing in epithelia.
J. Cell Sci. 113,2363
-2374.
Mackay, D. J., Esch, F., Furthmayr, H. and Hall, A.
(1997). Rho- and rac-dependent assembly of focal adhesion
complexes and actin filaments in permeabilized fibroblasts: an essential role
for ezrin/radixin/moesin proteins. J. Cell Biol.
138,927
-938.
Madara, J. L. (1987). Intestinal absorptive
cell tight junctions are linked to cytoskeleton. Am. J.
Physiol. 253,C171
-175.
Madara, J. L. (1998). Regulation of the movement of solutes across tight junctions. Ann. Rev. Physiol. 60,143 -159.[CrossRef][Medline]
Madara, J. L. and Dharmsathaphorn, K. (1985). Occluding junction structure-function relationships in a cultured epithelial monolayer. J. Cell Biol. 101,2124 -2133.[Abstract]
Madara, J. L., Moore, R. and Carlson, S.
(1987). Alteration of intestinal tight junction structure and
permeability by cytoskeletal contraction. Am. J.
Physiol. 253,C854
-861.
Madara, J. L., Stafford, J., Barenberg, D. and Carlson, S.
(1988). Functional coupling of tight junctions and microfilaments
in T84 monolayers. Am. J. Physiol.
254,G416
-423.
Michaely, P. A., Mineo, C., Ying, Y. S. and Anderson, R. G.
(1999). Polarized distribution of endogenous Rac1 and RhoA at the
cell surface. J. Biol. Chem.
274,21430
-21436.
Mineo, C. and Anderson, R. G. (2001). Potocytosis. Robert Feulgen Lecture. Histochem. Cell Biol. 116,109 -118.[Medline]
Nobes, C. D. and Hall, A. (1995). Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81,53 -62.[Medline]
Nusrat, A., Giry, M., Turner, J. R., Colgan, S. P., Parkos, C. A., Carnes, D., Lemichez, E., Boquet, P. and Madara, J. L. (1995). Rho protein regulates tight junctions and perijunctional actin organization in polarized epithelia. Proc. Natl. Acad. Sci. USA 92,10629 -10633.[Abstract]
Nusrat, A., Parkos, C. A., Verkade, P., Foley, C. S., Liang, T.
W., Innis-Whitehouse, W., Eastburn, K. K. and Madara, J. L.
(2000). Tight junctions are membrane microdomains. J.
Cell Sci. 113,1771
-1781.
Nusrat, A., von Eichel-Streiber, C., Turner, J. R., Verkade, P.,
Madara, J. L. and Parkos, C. A. (2001). Clostridium
difficile toxins disrupt epithelial barrier function by altering membrane
microdomain localization of tight junction proteins. Infect.
Immun. 69,1329
-1336.
Parkos, C. A., Colgan, S. P., Bacarra, A. E., Nusrat, A.,
Delp-Archer, C., Carlson, S., Su, D. H. and Madara, J. L.
(1995). Intestinal epithelia (T84) possess basolateral ligands
for CD11b/CD18-mediated neutrophil adherence. Am. J.
Physiol. 268,C472
-479.
Ridley, A. J. and Hall, A. (1992). The small GTP-binding protein rho regulates the assembly of focal adhesions and actin stress fibers in response to growth factors. Cell 70,389 -399.[Medline]
Ridley, A. J., Paterson, H. F., Johnston, C. L., Diekmann, D. and Hall, A. (1992). The small GTP-binding protein rac regulates growth factor-induced membrane ruffling. Cell 70,401 -410.[Medline]
Rojas, R., Ruiz, W. G., Leung, S. M., Jou, T. S. and Apodaca,
G. (2001). Cdc42-dependent modulation of tight junctions and
membrane protein traffic in polarized Madin-Darby canine kidney cells.
Mol. Biol. Cell. 12,2257
-2274.
Sagi, S. A., Seasholtz, T. M., Kobiashvili, M., Wilson, B. A.,
Toksoz, D. and Brown, J. H. (2001). Physical and functional
interactions of Galphaq with Rho and its exchange factors. J. Biol.
Chem. 276,15445
-15452.
Sanders, S. E., Madara, J. L., McGuirk, D. K., Gelman, D. S. and Colgan, S. P. (1995). Assessment of inflammatory events in epithelial permeability: a rapid screening method using fluorescein dextrans. Epithelial Cell Biol. 4,25 -34.[Medline]
Schmidt, G., Sehr, P., Wilm, M., Selzer, J., Mann, M. and Aktories, K. (1997). Gln 63 of Rho is deamidated by Escherichia coli cytotoxic necrotizing factor-1. Nature 387,725 -729.[CrossRef][Medline]
Schmidt, G. and Aktories, K. (1998). Bacterial cytotoxins target Rho GTPases. Naturwissenschaften 85,253 -261.[CrossRef][Medline]
Sears, C. L. and Kaper, J. B. (1996). Enteric
bacterial toxins: mechanisms of action and linkage to intestinal secretion.
Microbiol. Rev. 60,167
-215.
Seasholtz, T. M., Zhang, T., Morissette, M. R., Howes, A. L.,
Yang, A. H. and Brown, J. H. (2001). Increased expression and
activity of RhoA are associated with increased DNA synthesis and reduced
p27(Kip1) expression in the vasculature of hypertensive rats. Circ.
Res. 89,488
-495.
Sinnett-Smith, J., Lunn, J. A., Leopoldt, D. and Rozengurt, E. (2001). Y-27632, an inhibitor of Rho-associated kinases, prevents tyrosine phosphorylation of focal adhesion kinase and paxillin induced by bombesin: dissociation from tyrosine phosphorylation of p130(CAS). Exp. Cell Res. 266,292 -302.[CrossRef][Medline]
Takaishi, K., Sasaki, T., Kotani, H., Nishioka, H. and Takai,
Y. (1997). Regulation of cell-cell adhesion by Rac and Rho
small G proteins in MDCK cells. J. Cell Biol.
139,1047
-1059.
Troyanovsky, S. M. (1999). Mechanism of cell-cell adhesion complex assembly. Curr. Opin. Cell Biol. 5,561 -566.[CrossRef]
Turner, C. E. (2000). Paxillin interactions.
J. Cell Sci. 113,4139
-4140.
Turner, J. R. and Madara, J. L. (1995). Physiological regulation of intestinal epithelial tight junctions as a consequence of Na(+)-coupled nutrient transport. Gastroenterology 109,1391 -1396.[Medline]
Ullrich, O., Reinsch, S., Urbe, S., Zerial, M. and Parton, R. G. (1996). Rab11 regulates recycling through the pericentriolar recycling endosome. J. Cell Biol. 135,913 -924.[Abstract]
von Eichel-Streiber, C., Boquet, P., Sauerborn, M. and Thelestam, M. (1996). Large clostridial cytotoxins a family of glycosyltransferases modifying small GTP-binding proteins. Trends Microbiol. 4,375 -382.[CrossRef][Medline]
Vouret-Craviari, V., Grall, D., Flatau, G., Pouyssegur, J.,
Boquet, P. and Van Obberghen-Schilling, E. (1999). Effects of
cytotoxic necrotizing factor 1 and lethal toxin on actin cytoskeleton and
VE-cadherin localization in human endothelial cell monolayers.
Infect. Immun. 67,3002
-3008.
Walsh, S. V., Hopkins, A. M., Chen, J., Narumiya, S., Parkos, C. A. and Nusrat, A. (2001). Rho-kinase regulates tight junction function and is necessary for tight junction assembly in polarized intestinal epithelia. Gastroenterology 121,566 -579.[Medline]
Yuri, K., Nakata, K., Katae, H., Yamamoto, S. and Hasegawa, A. (1998). Distribution of uropathogenic virulence factors among Escherichia coli strains isolated from dogs and cats. J. Vet. Med. Sci. 60,287 -290.[CrossRef][Medline]