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*
Institut für Prophylaxe und Epidemiologie der Kreislaufkrankheiten, Universität München, and
Max-von-Pettenkofer Institut für Medizinische Mikrobiologie, Munich, Germany; and
Division of Signal Transduction, Nara Institute of Science and Technology, Ikoma, Japan
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PMT is an intracellularly acting toxin that enters cells via a poorly
defined endocytotic mechanism and activates a variety of signal
transduction pathways (5). By activation of a phospholipase Cß
(PLCß) isoenzyme, PMT increases the formation of diacylglycerol and
inositol-1,4,5-trisphosphate, which activate protein kinase C and
mobilize intracellular Ca2+, respectively (6, 7, 8). In
addition, PMT treatment of fibroblasts induced tyrosine phosphorylation
of several proteins, including focal adhesion kinase and paxillin
(9). It has been suggested that activation of the small GTPase
Rho mediates PMT-stimulated tyrosine phosphorylation of focal adhesion
kinase and paxillin as well as formation of stress fibers and focal
adhesion sites in fibroblasts (9). The Ras-related GTPase Rho has been
implicated in the organization of the actin cytoskeleton (10). A
variety of target proteins have been identified that specifically
interact with and are stimulated by GTP-bound Rho and thus mediate
downstream Rho functions (11, 12). Among the Rho targets is a 160-kDa
protein called Rho kinase/ROK
(13, 14, 15) that phosphorylates the
myosin-binding subunit of PP1 and thereby inhibits phosphatase activity
(16). In fibroblasts, microinjection of the Rho binding domain (RBD)
and the pleckstrin homology (PH) domain of Rho kinase blocked
lysophosphatidic acid-stimulated stress fiber and focal adhesion
formation, indicating that Rho kinase might be the target protein
important for actomyosin-dependent contractile events (17).
In the present study we investigated the mechanisms by which the bacterial product PMT increases endothelial permeability and the role of the GTPase Rho therein. Our data suggest that PMT activates Rho/Rho kinase, which inactivates PP1 and thus increases MLC phosphorylation. We propose that the resulting cell retraction then causes increased endothelial permeability. This mechanism could contribute to the observed vascular effects of PMT.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Okadaic acid, KT5926, tautomycin and BAPTA-acetoxymethyl ester were purchased from Calbiochem (Bad Soden, Germany). All other materials not specifically indicated were obtained from Sigma (Deisenhofen, Germany).
Cell culture
HUVEC were obtained and cultured as described previously (18). Briefly, cells harvested from umbilical cords were plated onto collagen-coated (24 h; 100 µg/ml collagen G; Biochrom, Berlin, Germany) plastic culture flasks and cultured in endothelial growth medium (Promo Cell, Heidelberg, Germany), containing endothelial cell growth supplement/heparin and 10% FCS. For all experiments, cells were plated at a density of 2 x 104 cells/cm2 and grown to confluence for 10 days, with medium changes every 2 or 3 days. Confluent monolayers were stimulated for the indicated time periods with rPMT (Sigma).
Measurement of endothelial permeability
Horseradish peroxidase (HRP) diffusion through HUVEC monolayers was determined as described previously with some modifications (19). Briefly, cells were plated (2 x 104 cells/cm2) on collagen-coated polyethylene terephtalate cell culture inserts (3-µm pore size; Becton Dickinson), which were set into 24-well Falcon Companion TC plates (Becton Dickinson) and cultured for 10 days with medium changes in the upper compartment every 2 days. For PMT stimulation, medium in the upper compartment was replaced with 500 µl of culture medium containing PMT. After 60 min of stimulation, 500 µl of medium was added to the lower compartment, and the medium in the upper compartment was replaced with fresh medium containing HRP (0.34 mg/ml; IV-A-type; 44,000 Mr; Sigma). For controls, PMT was omitted, but otherwise cells were treated identically. After 1 min, 60 µl of medium was collected from the lower compartment and mixed with 860 µl of reaction buffer (50 mM NaH2PO4, 5 mM guaiacol) and 100 µl of freshly made H2O2 solution (0.6 mM in H2O). The reaction was allowed to proceed for 15 min at room temperature, and absorbance was measured at 470 nm.
Immunofluorescence
For fluorescence staining, HUVEC were plated (2 x 104 cells/cm2) on Eppendorf Cellocate glass coverslips (Eppendorf, Hamburg, Germany) coated with 100 µg/ml collagen G (20°C, 24 h) and grown to confluence for 10 days. To label filamentous actin, cells were fixed for 10 min with 3.7% formaldehyde in PBS containing 1 mM Ca2+ and 1 mM Mg2+, permeabilized for 5 min in cold acetone (-20°C), and air-dried. Coverslips were then incubated for 20 min with rhodamine phalloidin (Molecular Probes, Oregon, OR; 1/20 in PBS) and mounted in Mowiol (Calbiochem) containing 0.2% p-phenylenediamine (Sigma) as anti-fading agent. All steps were performed at room temperature with three washes in PBS/2% BSA between Ab incubations. Fluorescence microscopy was performed with a Leica RBM 3 fluorescence microscope (Bemsheim, Germany), and microphotographs were recorded on Kodak T-Max 400 films (Eastman Kodak, Rochester, NY).
Recombinant proteins
Recombinant C3 transferase, the RBD, and the PH domain of Rho kinase were expressed as glutathione S-transferase fusion proteins in Escherichia coli and purified on glutathione-Sepharose beads as previously described (17, 20). The fusion proteins were cleaved by thrombin, the thrombin was removed by incubation with p-aminobenzamidine beads, and thereafter proteins were concentrated and dialyzed against microinjection buffer (see below). Purity and complete removal of thrombin were checked by SDS-PAGE and Coomassie staining. Protein concentrations were determined with the BCA Protein Assay Kit (Pierce, Rockford, IL) using BSA as standard. As tested by SDS-PAGE and Coomassie staining, protein preparations showed essentially only one band.
Microinjection
Microinjection was performed with an Eppendorf Transjector 5246
and an Eppendorf Micromanipulator 5171. Cells were plated and cultured
on Cellocate coverslips (Eppendorf) as described above. The RBD from
Rho kinase or the PH domain from Rho kinase was diluted with
microinjection buffer (150 mM NaCl, 50 mM Tris, and 5 mM
MgCl2, pH 7.5) and injected at concentrations of 0.64
µg/µl (RBD) and 0.96 µg/µl (PH) into the cytoplasm of HUVEC.
The PP1 catalytic subunit (-isoform; Calbiochem) was diluted with
phosphatase buffer (100 mM K+ glutamate and 39 mM
K+ citrate, pH 7.3) and was injected at a concentration of
200 U/ml. Control injections conducted with microinjection or
phosphatase buffer, respectively, did not produce any significant
effect on cell morphology or actin organization. The microinjected
volume was about 1–3 x 10-15 L/cell. Injected cells
were identified by coinjecting rat IgG (5 mg/ml) followed by staining
with FITC-conjugated goat anti-rat IgG (Dianova, Hamburg, Germany).
For each experiment at least 100 cells were injected and examined by
fluorescence microscopy.
MLC phosphorylation
MLC phosphorylation was analyzed by urea-PAGE separation of the mono- and diphosphorylated MLC forms as described in detail previously (21, 22). HUVEC grown in 100-mm diameter dishes were stimulated with thrombin as indicated, and the reaction was terminated by addition of 1.5 ml of ice-cold 10% TCA. Cells were scraped and then centrifuged for 20 min at 14,000 x g at 4°C. Supernatants were discarded, and pellets were washed with ddH2O to remove TCA and resolved in 1.5 ml of sample buffer (6.7 M urea, 20 mM Tris, 22 mM glycine, and 10 mM DTT, pH 9.0). Samples were applied to urea-gel electrophoresis (top gel, 3.5% acrylamide; bottom gel, 10% acrylamide, 20 mM Tris, and 22 mM glycine) and were run at 9 mA for approximately 45 min until the marker dye had come out of the bottom gel. Proteins were then electroblotted onto polyvinylidene difluoride membranes at 25 V for 1.5 h. Membranes were incubated overnight with anti-MLC Ab IgM (1/100 in Tris-buffered saline (TBS) containing 0.3% Tween 20; Sigma), washed three times, incubated for 1 h with biotinylated anti-mouse IgM Ab (1/500 in TBS; Amersham, Arlington Heights, IL), washed three times, and then incubated for 30 min with HRP-streptavidin conjugate (1/1,000 in TBS; Amersham). Membranes were developed with Luminol solution (Pierce) and exposed to Kodak X-OMAT films. The stochiometry of MLC phosphorylation (moles of phosphate per moles of MLC) was determined by densitometric quantitation of the unphosphorylated and phosphorylated (faster migrating) protein bands that reacted with anti-MLC Ab using a Sharp XL-325 densitometer and Pharmacia Image Master software and was calculated using the formula P1 + 2 x P2/P0 + P1 + P2, where P0, P1, and P2 are un-, mono-, and diphosphorylated MLC, respectively.
Preparation of myosin-enriched cell fractions
Myosin-enriched fractions of HUVEC were prepared as described previously (23). Briefly, HUVEC were plated on collagen-coated 100-mm diameter plates (Falcon) and cultivated for 10 days. Monolayers were washed twice with ice-cold PBS (Sigma) and 200 µl of homogenization buffer (50 mM Tris-aminomethane (pH 7.5), 0.1 mM EDTA, 28 mM ß-ME, leupeptin, pepstatin, Pefabloc (Boehringer Mannheim, Mannheim, Germany), and aprotinin, 1 µg/ml each) was added. Plates were incubated at -80°C, scraped with a rubber policeman, and homogenized by passing the suspension 5–10 times through a syringe. Homogenates were then treated with high salt buffer (0.6 M NaCl and 0.1% Tween 20, containing 1 µg/ml of leupeptin, pepstatin, aprotinin, and Pefablock) for 1 h at 4°C and subsequently centrifuged at 4500 x g for 30 min at 4°C. The supernatant was diluted 10-fold with assay buffer (50 mM Tris, 0.1 mM EDTA, and 28 mM ß-ME, pH 7.0) and centrifuged for 30 min at 10,000 x g at 4°C. The resulting pellet was resolved in 10 µl of high salt buffer. This myosin-enriched cell fraction contains mostly PP1 and essentially no PP2 activity (23). By Western blot using anti-PP1 Ab we confirmed that equal amounts of PP1 were present in myosin-enriched samples.
Measurement of myosin-associated phosphatase activity
For measuring myosin-associated phosphatase activity in
myosin-enriched cell fractions we used the Protein Phosphatase Assay
System (Life Technologies, Gaithersburg, MD) according to the
instructions of the manufacturer. This assay system is based on the
method described by Cohen (24). Briefly, phosphorylase b (0.1 mM) was
in vitro phosphorylated by phosphorylase kinase (0.1 mg/ml) in the
presence of [
-32P]ATP (5 mCi/ml) in phosphorylation
buffer (250 mM Tris-HCl (pH 8.2), 16.7 mM MgCl2, 1.67 mM
ATP, 0.83 mM CaCl2, and 133 mM 2-ME) for 1 h at
30°C. The reaction was stopped with 90% ammoniumpersulfate solution
(4°C), kept on ice for 1 h, and subsequently centrifuged at
12,000 x g for 10 min. The protein pellet was
resuspended and subsequently washed four times in ammoniumpersulfate
solution (45% saturated). Proteins were then concentrated to a final
concentration of 3 mg/ml using Amicon Centricon-30R concentrators
(Beverley, MA), and phosphatase quantified by measuring release of
radioactivity from [-32P]phosphorylase a. For this
purpose myosin-enriched fractions were diluted with 30 µl of assay
buffer (50 mM Tris, 0.1 mM EDTA, 28 mM ß-ME, and 6.25 mM caffeine, pH
7.0) and mixed with 20 µl of radioactive phosphatase substrate. The
reaction was allowed to proceed for 10 min at 30°C and was
stopped with ice-cold 20% TCA. Samples were incubated on ice for 10
min and were centrifuged at 12,000 x g for 3 min. The
radioactivity released in the supernatant was measured using a Wallac
1410 liquid scintillation counter (Gaithersburg, MD).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Inflammatory mediators such as thrombin or histamine stimulate
actomyosin-dependent endothelial cell retraction, which causes an
increase in endothelial permeability (25). As an in vitro assay for a
PMT-induced increase in endothelial permeability we measured diffusion
of HRP through tightly confluent HUVEC monolayers. The results
presented in Fig. 1
demonstrate that
monolayers show a low transendothelial diffusion of HRP.
Stimulation with PMT (1 h, 40 ng/ml) increased diffusion up to 10-fold
(Fig. 1). To investigate the role of Rho in the PMT-induced increase in
HRP diffusion we pretreated cells with the specific Rho inhibitor C3
transferase from Clostridium botulinum. C3 transferase
ADP-ribosylates Asn41 in the Rho effector region and
thus inactivates Rho specifically (10). As evident from Fig. 1, the
PMT-induced increase in permeability could be almost completely
abolished by C3 transferase pretreatment (24 h, 5 µg/ml). We
previously found that under these conditions C3 ADP-ribosylates
70–80% of total RhoA in HUVEC (8). Our results suggest that Rho
mediates the PMT-induced increase in endothelial permeability. It has
been demonstrated that Rho can inactivate PP1 via its target Rho kinase
(16). If this mechanism is relevant in our system, the C3 transferase
could prevent a Rho/Rho kinase-induced inhibition of PP1 activity, and
consequently, the PMT effect in C3-treated cells would be restored by
external inhibition of PP1. In fact, in endothelial cells pretreated
with C3 transferase the effect of PMT could largely be restored by the
PP1 inhibitor tautomycin (15 min, 6 µM; Fig. 1). Together these data
suggest that PMT increases endothelial permeability through activation
of Rho and inhibition of myosin-bound MLC phosphatase (PP1 M) activity.
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To demonstrate directly that PMT induces shape changes in
endothelial cells we performed phase contrast microscopy and stained
filamentous actin with rhodamine phalloidin followed by
immunofluorescence microscopy. In control monolayers, actin was mainly
localized in a peripheral ring delineating the cell borders (Fig. 2B). In the C3
transferase-treated monolayer, we occasionally found giant cells
suggestive of a maximally spread phenotype (Fig. 2C), but
cell shapes were otherwise not different from controls (compare Fig. 2, A and C). Actin was slightly thinned out along
the cell borders in the C3-treated cells, but its continuity was well
preserved (compare Fig. 2, B and D). Stimulation
with PMT (1 h, 40 ng/ml) induced prominent actin stress fibers and
intercellular gaps (Fig. 2, E and F). However,
when cells were pretreated with C3 (24 h, 5 µg/ml), the PMT-induced
actin rearrangements were essentially abolished, indicating involvement
of Rho (Fig. 2H). We also noticed the development of a large
number of giant cells in the C3-pretreated and PMT-stimulated cells
(Fig. 2G). We have no explanation for this phenomenon at
present, but PMT and circumstantially C3 could activate a signal
pathway promoting cell spreading, such as activation of Rac (10, 11).
To test whether interaction of Rho with Rho kinase is necessary for
PMT-induced stress fiber formation, we microinjected the rRBD or the PH
domain of Rho kinase. Isolated RBD and PH domain have been used to
inhibit interaction of active Rho with endogenous Rho kinase (17).
Microinjection of both the RBD and the PH domain of Rho kinase
completely abolished PMT-induced actin rearrangements (Fig. 3, A–D), indicating that PMT
stimulates Rho/Rho kinase interaction. Rho kinase has been shown to
phosphorylate and inactivate PP1. To investigate the contribution of
PP1 to the PMT-stimulated actin rearrangements we microinjected the
constitutively active catalytic domain of PP1 (Fig. 3, E and F) in the PMT-stimulated cells. In fact,
PMT-induced stress fiber formation was completely abolished by
microinjection of PP1. These results are consistent with the idea that
PMT decreases PP1 activity through Rho kinase in endothelial cells. To
test whether MLCK activity is also required for PMT-induced
reorganization of the cytoskeleton we pretreated cells with the
selective MLCK inhibitor KT5926. As indicated in Fig. 3G, actin rearrangements were prevented by KT5926. We
conclude from these data that at least a basal MLCK activity is
required for the PMT-induced actin rearrangement.
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We measured okadaic acid-insensitive release of
32PO4 from [32P]phosphorylase b
in myosin fractions of HUVEC to clarify whether PMT, in fact,
inactivates PP1 M (23, 24). As shown in Fig. 4A, PMT treatment (1–24 h, 40 ng/ml)
produced an effective (
50%) and long-term (up to 24 h)
inhibition of PP1 M activity. In HUVEC pretreated with C3 transferase
(5 µg/ml, 24 h), the PMT-induced decrease in PP1 M activity was
essentially abolished (Fig. 4A). We conclude that PMT
inactivates PP1 M in HUVEC by a Rho-dependent mechanism.
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B). Taken together these data suggest that PMT inactivates
PP1 M via Rho/Rho kinase, and this causes an increase in MLC
phosphorylation. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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depicts the
presumptive signal pathway by which PMT induces cell retraction and
increased endothelial permeability. The inhibition of PP1 by Rho/Rho
kinase induces, in concert with Ca2+-dependent MLCK
activity, an increase in MLC phosphorylation that together with actin
drives contraction.
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In contrast to CNF and dermonecrotizing toxin from B.
bronchiseptica, PMT does not seem to directly activate Rho in
various cell types (6, 9), yet according to our findings and the
results of others (9) it stimulates a signal pathway involving Rho. A
study in Xenopus oocytes revealed that PMT stimulates Gq,
which couples to and activates PLCß1 and increases inositol
1,4,5-trisphosphate (IP3) in this manner. In that study it
was speculated, but not biochemically demonstrated, that PMT directly
modified and activated Gq (6). Furthermore, in that study
microinjection of anti-RhoA and RhoB Abs did not block the PMT
effect, suggesting that Rho is not involved in PMT-induced
IP3 production. It thus remains to be tested whether
Gq/PLCß or their product, IP3, can activate Rho in
endothelial cells or whether Rho activation occurs independent of this
pathway. Interestingly, it has been reported that Rho can be stimulated
by PLC activity (6, 30). In contrast, other reports suggest that Rho
can stimulate PLC-mediated inositol metabolism in N1E-115
neuroblastoma cells (31). Alternate signals that could be responsible
for Rho activation by PMT are activation of tyrosine kinases (9, 32, 33).
Besides through cell retraction mediators such as thrombin are thought to increase endothelial permeability by disassembling vascular endothelial-cadherin-based adherens junctions (34). Although our results clearly identify the Rho/Rho kinase-mediated shape changes as a major regulator of the PMT-stimulated increase in endothelial permeability, we cannot exclude that disintegration of adherens junctions is also involved. It therefore remains to be investigated whether in addition to cell retraction, adherens junctions are disassembled by PMT, for example by tyrosine phosphorylation or protein kinase C-dependent dephosphorylation of the vascular endothelial-cadherin-associated catenins (34, 35).
In summary, we provide evidence that PMT activates the Rho/Rho kinase pathway and thus inactivates PP1, which then increases MLC phosphorylation. This mechanism could contribute to the pathophysiologically important disturbance of endothelial integrity brought about by PMT (and presumably other Rho-activating bacterial toxins) in human infection and inflammation.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Markus Essler (E-mail address: ) or Dr. Martin Aepfelbacher (E-mail address: ), Institut für Prophylaxe und Epidemiologie der Kreislaufkrankheiten, Universität München, Pettenkoferstr 9, 80336 Munich, Germany.
3 Abbreviations used in this paper: PMT, Pasteurella multocida toxin; MLC, myosin light chain; MLCK, MLC kinase; PP1, MLC phosphatase; PLC, phospholipase C; PH, pleckstrin homology; RBD, Rho binding domain; HRP, horseradish peroxidase; C3, C3 transferase from Clostridium botulinum; TBS, Tris-buffered saline; PP1 M, myosin-bound PP1; IP3, inositol 1,4,5-trisphosphate.
Received for publication January 22, 1998. Accepted for publication July 9, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-coupled phospholipase C-ß1. J. Biol. Chem. 272:1268. is a member of a kinase family and is involved in the organisation of the cytoskeleton. Mol. Cell. Biol. 16:5313.[Abstract]
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