Pasteurella multocida Toxin Increases Endothelial Permeability via Rho Kinase and Myosin Light Chain Phosphatase This information is current as of July 17, 2015.
Markus Essler, Karin Hermann, Mutsuki Amano, Kozo Kaibuchi, Jürgen Heesemann, Peter C. Weber and Martin Aepfelbacher J Immunol 1998; 161:5640-5646; ; http://www.jimmunol.org/content/161/10/5640
Subscriptions Permissions Email Alerts
This article cites 33 articles, 19 of which you can access for free at: http://www.jimmunol.org/content/161/10/5640.full#ref-list-1 Information about subscribing to The Journal of Immunology is online at: http://jimmunol.org/subscriptions Submit copyright permission requests at: http://www.aai.org/ji/copyright.html Receive free email-alerts when new articles cite this article. Sign up at: http://jimmunol.org/cgi/alerts/etoc
The Journal of Immunology is published twice each month by The American Association of Immunologists, Inc., 9650 Rockville Pike, Bethesda, MD 20814-3994. Copyright © 1998 by The American Association of Immunologists All rights reserved. Print ISSN: 0022-1767 Online ISSN: 1550-6606.
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015
References
Pasteurella multocida Toxin Increases Endothelial Permeability via Rho Kinase and Myosin Light Chain Phosphatase1 Markus Essler,2* Karin Hermann,* Mutsuki Amano,‡ Kozo Kaibuchi,‡ Ju¨rgen Heesemann,† Peter C. Weber,* and Martin Aepfelbacher2*†
I
nfections due to Pasteurella multocida are characterized by acute inflammatory symptoms of the skin, such as edema, and are often complicated by sepsis or dermonecrosis (1). P. multocida produces a toxin (PMT)3 that when intracutaneously injected into laboratory animals can induce all the symptoms of P. multocida infection, including edema, emigration of neutrophils, and increased attachment of thrombocytes to the vessel wall, suggesting an effect of PMT on the endothelium (2). It is well established that inflammatory mediators such as thrombin increase vascular permeability by inducing actomyosin-dependent endothelial cell retraction (3). Thrombin increases myosin light chain (MLC) phosphorylation both by activating Ca21/calmodulin-dependent MLC kinase (MLCK) and by deactivating MLC phosphatase (PP1) (4). The molecular mechanisms by which bacterial toxins such as PMT disturb endothelial integrity remain to be established. 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 *Institut fu¨r Prophylaxe und Epidemiologie der Kreislaufkrankheiten, Universita¨t Mu¨nchen, and †Max-von-Pettenkofer Institut fu¨r Medizinische Mikrobiologie, Munich, Germany; and ‡Division of Signal Transduction, Nara Institute of Science and Technology, Ikoma, Japan Received for publication January 22, 1998. Accepted for publication July 9, 1998. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. 1 This work was supported by Deutsche Forschungsgemeinschaft (Ae11/5-1 and SFB 413) and August Lenz Stiftung. This work is part of the doctoral thesis of K.H. at the University of Munich. 2 Address correspondence and reprint requests to Dr. Markus Essler (E-mail address:
[email protected]) or Dr. Martin Aepfelbacher (E-mail address:
[email protected]), Institut fu¨r Prophylaxe und Epidemiologie der Kreislaufkrankheiten, Universita¨t Mu¨nchen, Pettenkoferstr 9, 80336 Munich, Germany.
Cb (PLCb) isoenzyme, PMT increases the formation of diacylglycerol and inositol-1,4,5-trisphosphate, which activate protein kinase C and mobilize intracellular Ca21, 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/ROKa (13–15) 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.
Materials and Methods Materials
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. Copyright © 1998 by The American Association of Immunologists
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). 0022-1767/98/$02.00
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015
Pasteurella multocida toxin (PMT) has been shown to induce actin reorganization through activation of the GTPase Rho. Here we investigated the involvement of the Rho target proteins Rho kinase and myosin light chain (MLC) phosphatase in the PMTinduced increase in endothelial permeability and the underlying actin reorganization of endothelial cells. Stimulation of endothelial layers with PMT enhanced transendothelial permeability >10-fold, and this was abolished by pretreatment with the specific Rho inactivator C3 transferase from Clostridium botulinum. The PMT-induced increase in endothelial permeability was associated with 1) inactivation of MLC phosphatase, 2) an increase in MLC phosphorylation, and 3) endothelial cell retraction and actin stress fiber formation. PMT-stimulated actin reorganization could be prevented by 1) pretreatment of cells with C3 transferase, 2) microinjection of the Rho binding domain and the pleckstrin homology domain of Rho kinase, and 3) microinjection of constitutively active MLC phosphatase. Together, these results suggest that PMT activates Rho/Rho kinase, which inactivates MLC phosphatase. The resulting increase in MLC phosphorylation causes endothelial cell retraction and a rise in endothelial permeability. The Journal of Immunology, 1998, 161: 5640 –5646.
The Journal of Immunology
5641 MLC phosphorylation
HUVEC were obtained and cultured as described previously (18). Briefly, cells harvested from umbilical cords were plated onto collagen-coated (24 h; 100 mg/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 3 104 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).
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 3 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 HRPstreptavidin 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 1 2 3 P2/P0 1 P1 1 P2, where P0, P1, and P2 are un-, mono-, and diphosphorylated MLC, respectively.
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 3 104 cells/cm2) on collagen-coated polyethylene terephtalate cell culture inserts (3-mm 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 ml of culture medium containing PMT. After 60 min of stimulation, 500 ml 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 ml of medium was collected from the lower compartment and mixed with 860 ml of reaction buffer (50 mM NaH2PO4, 5 mM guaiacol) and 100 ml 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
Preparation of myosin-enriched cell fractions
For fluorescence staining, HUVEC were plated (2 3 104 cells/cm2) on Eppendorf Cellocate glass coverslips (Eppendorf, Hamburg, Germany) coated with 100 mg/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 Ca21 and 1 mM Mg21, permeabilized for 5 min in cold acetone (220°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).
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 ml of homogenization buffer (50 mM Tris-aminomethane (pH 7.5), 0.1 mM EDTA, 28 mM b-ME, leupeptin, pepstatin, Pefabloc (Boehringer Mannheim, Mannheim, Germany), and aprotinin, 1 mg/ml each) was added. Plates were incubated at 280°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 mg/ml of leupeptin, pepstatin, aprotinin, and Pefablock) for 1 h at 4°C and subsequently centrifuged at 4500 3 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 b-ME, pH 7.0) and centrifuged for 30 min at 10,000 3 g at 4°C. The resulting pellet was resolved in 10 ml of high salt buffer. This myosinenriched 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.
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 SDSPAGE 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 mg/ml (RBD) and 0.96 mg/ml (PH) into the cytoplasm of HUVEC. The PP1 catalytic subunit (a-isoform; Calbiochem) was diluted with phosphatase buffer (100 mM K1 glutamate and 39 mM K1 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 3 10215 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.
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 [g-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 3 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 [g-32P]phosphorylase a. For this purpose myosin-enriched fractions were diluted with 30 ml of assay buffer (50 mM Tris, 0.1 mM EDTA, 28 mM b-ME, and 6.25 mM caffeine, pH 7.0) and mixed with 20 ml 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 3 g for 3 min. The radioactivity released in the supernatant was measured using a Wallac 1410 liquid scintillation counter (Gaithersburg, MD).
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015
Cell culture
5642
FIGURE 1. P. multocida toxin increases endothelial permeability. HUVEC were pretreated for 24 h without or with 5 mg/ml C3 transferase (C3) from C. botulinum or for 15 min with 6 mM of the PP1 inhibitor tautomycin (Taut) and then stimulated with PMT (1 h, 40 ng/ml). Transendothelial diffusion of HRP was determined as described in Materials and Methods. Bars represent the mean 6 SD of one experiment performed in quadruplicate. Where absent, error bars are within symbols.
PMT increases transendothelial permeability by a Rhodependent mechanism 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 ADPribosylates 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 mg/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 mM; Fig. 1). Together these data suggest that PMT increases endothelial permeability through activation of Rho and inhibition of myosinbound MLC phosphatase (PP1 M) activity. PMT-induced shape change in HUVEC is mediated by Rho and its target Rho kinase 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 mg/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. PMT inactivates PP1 and increases MLC phosphorylation 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 mg/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. Pharmacologic inhibition of PP1 M in HUVEC was shown to increase MLC phosphorylation, cell contractility, and permeability (23). We therefore tested whether inactivation of PP1 M by PMT is associated with increased MLC phosphorylation. HUVEC were stimulated with PMT, and un- (P0), mono- (P1), and diphosphorylated (P2) MLCs were separated by urea-gel electrophoresis and detected by a specific Ab against MLC in Western blot. Protein bands were quantified by densitometry, and phosphate incorporation into MLC was calculated (see Materials and Methods). We found that MLC phosphorylation increased in a time-dependent manner within 24 h of PMT stimulation (Fig. 4B). Taken together these data suggest that PMT inactivates PP1 M via Rho/Rho kinase, and this causes an increase in MLC phosphorylation.
Discussion Here we describe a signal transduction pathway by which P. multocida toxin stimulates actin reorganization and an increase in permeability of human endothelial layers. We propose that PMT activates Rho, which, via its target Rho kinase, inhibits PP1 M. Inhibition of PP1 leads to an increase in MLC phosphorylation, supposedly because MLC kinase activity is not counter balanced
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015
Results
P. multocida TOXIN SIGNALING IN ENDOTHELIAL CELLS
The Journal of Immunology
5643
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015 FIGURE 2. PMT-induced intercellular gap formation and actin reorganization are Rho dependent. HUVEC were not stimulated (A and B), were treated with C3 transferase (24 h, 5 mg/ml; C and D), were stimulated with PMT (1 h, 40 ng/ml; E and F), or were pretreated with C3 transferase (24 h, 5 mg/ml) and then stimulated with PMT (G and H). A, C, E, and G show phase contrasts, and B, D, F, and H show actin staining using rhodamine phalloidin. Representative pictures are shown. Bar represents 30 mm.
by PP1. MLC kinase activity seems to be active in unstimulated endothelial cells (23) and is most likely further stimulated by the PMT-induced increase in intracellular Ca21 release (7). Consistent with this idea, we found that the MLC kinase inhibitor KT5926 inhibits PMT-induced actin reorganization. Hence, to increase MLC phosphorylation PMT seems to inhibit PP1 activity in the presence of a basal or slightly activated Ca21-calmodulin-dependent MLC kinase activity. Interestingly, PMT increased MLC phosphorylation only moderately compared with mediators such as histamine, but for a time period of at least 24 h, while histamineinduced MLC-phosphorylation returned to basal levels in 30 min or less (25). We suggest that PMT exerts its effect on cell shape via a moderate but prolonged elevation of MLC phosphorylation. Fig.
5 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 Ca21dependent MLCK activity, an increase in MLC phosphorylation that together with actin drives contraction. Two toxins from E. coli, CNF1 and CNF2, show sequence homology to PMT. PMT and CNF2 share 27% identical and 85% conserved residues at the N terminus (26). CNF1 was reported to deamidate glutamine residue 63 of Rho, which puts Rho into the constitutively active GTP-bound state (27, 28). Furthermore, dermonecrotizing toxin from Bordetella bronchiseptica, which shares homologous sequences with CNF at the C terminus, also modifies and activates Rho by deamidation (29).
5644
P. multocida TOXIN SIGNALING IN ENDOTHELIAL CELLS
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015 FIGURE 3. PMT-induced actin rearrangements are mediated by Rho/Rho kinase and PP1. Single cells were microinjected with the RBD of Rho kinase (A and B) or the PH domain of Rho kinase (C and D) and then stimulated with PMT (1 h, 40 ng/ml). Alternatively, cells were treated with PMT and then injected with the catalytic domain of PP1 (E and F). To identify injected cells, rat IgG was coinjected with all constructs. All cells were double stained for actin using rhodamine-phalloidin and for rat IgG using FITC-labeled anti-rat IgG. A, C, and E show actin staining (rhodamine fluorescence), and in B, D, and F, injected cells can be identified by FITC fluorescence. In G, cells were pretreated with the selective MLCK inhibitor KT-5926 (24 h, 5 mM) and then stimulated with PMT. KT-5926 could completely block the PMT effect. Photographs are representative of three experiments. Bar represents 30 mm.
The Journal of Immunology
5645
FIGURE 4. Inactivation of PP1 by PMT correlates with an increase in MLC phosphorylation. HUVEC were stimulated with PMT (40 ng/ml), and at the indicated time points PP1 M activity (A) and MLC phosphorylation (B) were determined as described in Materials and Methods. p, PP1 M activity in C3-pretreated cells (24 h, 5 mg/ml). Phosphatase values in A represent the mean 6 SEM of three experiments. Phosphorylation values in B represent the mean of one experiment performed in triplicate. The inset shows the original gel from which the phosphorylation values are derived through densitometric analysis as described in Materials and Methods.
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.
Acknowledgments 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 Gqa, which couples to and activates PLCb1 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 Gqa (6). Furthermore, in that study microinjection of antiRhoA 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 Gqa/PLCb 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
We thank Barbara Bo¨hlig for expert technical assistance, Manfred Schliwa (Institut fu¨r Zellbiologie, LMU Mu¨nchen) for help with microinjection, and Markus Bauer for densitometry.
References 1. Griego, R. D., T. Rosen, I. F. Orengo, and J. E. Wolf. 1995. Dog, cat and human bites: a review. J. Am. Acad. Dermatol. 33:1019. 2. Elling, F., K. B. Pedersen, P. Hogh, and N. T. Foged 1988. Characterisation of the dermal lesions induced by a purified protein from toxigenic Pasteurella multocida. Acta Pathol. Microbiol. Infectiol. Scand. 96:50. 3. Wysolmerski, R. B., and D. Lagunoff. 1990. Involvement of myosin light chain kinase in endothelial cell retraction. Proc. Natl. Acad. Sci. USA 87:16. 4. Shasby, M. D., T. Stevens, D. Ries, A. B. Moy, J. M. Kamath, A. M. Kamath, and S. S. Shasby. 1997. Thrombin inhibits myosin light chain dephosphorylation in endothelial cells. Am. J. Physiol. 272:L311. 5. Rozengurt, E., T. Higgins, N. Chanter, A. J. Lax, and J. M. Staddon. 1990. Pasteurella multocida toxin: potent mitogen for cultured fibroblasts. Proc. Natl. Acad. Sci. USA 87:123. 6. Wilson, B. A., X. Zhu, M. Ho, and L. Lu. 1997. Pasteurella multocida toxin activates the inositol trisphosphate signalling pathway in Xenopus oocytes via Gqa-coupled phospholipase C-b1. J. Biol. Chem. 272:1268. 7. Staddon, J. M., C. J. Barker, A. J. Murphy, N. Chanter, A. J. Lax, R. H. Michell, and E. Rozengurt. 1991. Pasteurella multocida toxin, a potent mitogen, increases inositol 1,4,5-trisphosphate and mobilises Ca21 in Swiss 3T3 cells. J. Biol. Chem. 268:480. 8. Staddon, J. M., N. Chanter, A. J. Lax, T. E. Higgins, and E. Rozengurt. 1990. Pasteurella multocida toxin, a potent mitogen, stimulates protein kinase C-dependent and independent protein phosphorylation in Swiss 3T3 cells. J. Biol. Chem. 265:11841. 9. Lacerda, H. M., A. J. Lax, and E. Rozengurt. 1996. Pasteurella multocida toxin, a potent intracellularly acting mitogen, induces p125 FAK and paxillin tyrosine
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015
FIGURE 5. PMT signaling network controlling vascular permeability. Rho-GTP, GTP-bound Rho; Rho-K, Rho kinase; Ca11-Cam, calciumcalmodulin; MLC-P, phosphorylated MLC; MLC-Pase, PP1; MLC-Pase-P, phosphorylated PP1; EC, endothelial cell.
5646
10. 11. 12.
13.
14.
15.
16.
17.
19.
20.
21. 22.
23. Verin, A. D., C. E. Patterson, M. A. Day, and J. G. N. Garcia. 1995. Regulation of endothelial cell gap formation and barrier function by myosin-associated phosphatase activities. Am. J. Physiol. 269:L99. 24. Chisholm, A. A. K., and P. Cohen. 1988. Identification of a third form of protein phosphatase 1 in rabbit skeletal muscle that is associated with myosin. Biochim. Biophys. Acta 968:392. 25. Moy, A. B., S. S. Shasby, B. D. Scott, and D. M. Shasby. 1993. The effect of histamine and cyclic adenosine monophosphate on myosin light chain phosphorylation in human umbilical vein endothelial cells. J. Clin. Invest. 92:1198. 26. Oswald, E., M. Sugai, A. Labigne, H. C. Wu, C. Fiorentini, P. Boquet, and A. O’Brien. 1994. Cytotoxic necrotising factor type 2 produced by virulent Escherichia coli modifies the small GTP-binding protein Rho involved in assembly of actin stress fibers. Proc. Natl. Acad. Sci. USA 91:3814. 27. Schmidt, G., P. Sehr, M. Wilm, J. Selzer, M. Mann, and K. Aktories. 1997. Gln 63 of Rho is deamidated by Escherichia coli cytotoxic necrotising factor-1. Nature 387:72. 28. Flatau, G., E. Lemichez, M. Gauthier, P. Chardin, S. Paris, C. Fiorentini, and P. Boquet. 1997. Toxin-induced activation of the G protein p21 Rho by deamidation of glutamine. Nature 387:729. 29. Horiguchi, Y., N. Inoue, M. Masuda, T. Kashimoto, J. Katahira, N. Sugimoto, and M. Matsuda. 1997. Bordetella bronchiseptica dermonecrotizing toxin induces reorganization of actin stress fibers through deamidation of Gln-63 of the GTP-binding protein Rho. Proc. Natl. Acad. Sci. USA 94:11623. 30. Takai, Y., T. Sasaki, K. Tanaka, and T. Nakanishi. 1995. Rho as a regulator of the cytoskeleton. Trends Biochem. Sci. 20:227. 31. Zhang, C., M. Schmidt, C. von Eichel-Streiber, and K. H. Jakobs. 1996. Inhibition by toxin B of inositol phosphate formation induced by G-protein-coupled and tyrosine kinase receptors in N1E-115 neuroblastoma cells: involvement of Rho proteins. Mol. Pharmacol. 50:227. 32. Nobes, C. D., P. Hawkins, L. Stephens, and A. Hall. 1995. Activation of the small GTP-binding proteins Rho and Rac by growth factor receptors. J. Cell Sci. 108: 225. 33. Gohla, A., R. Harhammer, and G. Schultz. 1998. The G-protein G13 but not G12 mediates signaling from lysophosphatidic acid receptor via epidermal growth factor receptor to Rho. J. Biol. Chem. 273:4653. 34. Rabiet, M. J., J. L. Plantier, Y. Rival, Y. Genoux, M. G. Lampugnani, and E. Dejana. 1996. Thrombin-induced increase in endothelial permeability is associated with changes in cell-to-cell junction organisation. Arterioscler. Thromb. Vasc. Biol. 16:488. 35. Ratcliffe, M. J., L. L. Rubin, and J. M. Staddon. 1997. Dephosphorylation of the cadherin-associated p100/p120 proteins in response to activation of protein kinase C in epithelial cells. J. Biol. Chem. 272:31894.
Downloaded from http://www.jimmunol.org/ by guest on July 17, 2015
18.
phosphorylation, actin stress fiber formation, and focal contact assembly in Swiss 3T3 cells. J. Biol. Chem. 271:439. Tapon, N., and A. Hall. 1997. Rho, Rac and CDC42 GTPases regulate the organisation of the actin cytoskeleton. Curr. Opin. Cell Biol. 9:86. Machesky, L. M., and A. Hall. 1996. Rho: a connection between membrane receptor signalling and the cytoskeleton. Trends Cell Biol. 6:304. Watanabe, G., Y. Saito, P. Madaule, T. Ishizaki, K. Fujisawa, N. Morii, H. Mukai, Y. Ono, A. Kakizuka, and S. Narumiya. 1996. Protein kinase N (PKN) and PKN-related protein rhophilin as targets of small GTPase Rho. Science 271: 645. Matsui, T., M. Amano, T. Yamamoto, K. Chihara, M. Nakafuku, M. Ito, T. Nakano, K. Okawa, A. Iwamatsu, and K. Kaibuchi. 1996. Rho-associated kinase, a novel serine/threonine kinase, as a putative target for the small GTP binding protein Rho. EMBO J. 15:2208. Leung, T., X. Chen, E. Manser, and L. Lim. 1996. The p160 RhoA-binding kinase ROKa is a member of a kinase family and is involved in the organisation of the cytoskeleton. Mol. Cell. Biol. 16:5313. Ishizaki, T., M. Maekawa, K. Fujisawa, K. Okawa, A. Iwamatsu, A. Fujita, N. Watanabe, Y. Saito, A. Kakizuka, N. Morii, et al. 1996. The small GTPbinding protein Rho binds to and activates a 160 kDa Ser/Thr protein kinase homologous to myotonic dystrophy kinase. EMBO J. 15:1885. Kimura, K., M. Ito, M. Amano, K. Chihara, Y. Fukata, M. Nakafuku, B. Yamamori, Y. Feng, T. Nakano, K. Okawa, et al. 1996. Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273:245. Amano, M., K. Chihara, K. Kimura, Y. Fukata, N. Nakamura, Y. Matsuura, and K. Kaibuchi. 1997. Formation of actin stress fibers and focal adhesions enhanced by Rho-kinase. Science 275:1308. Aepfelbacher, M., M. Essler, E. Huber, M. Sugai, and P. C. Weber. 1997. Bacterial toxins block endothelial wound repair: evidence that Rho GTPases control cytoskeletal rearrangements in migrating endothelial cells. Arterioscler. Thromb. Vasc. Biol. 17:1623. Lampugnani, M. G., M. Resnati, E. Dejana, and P. C. Marchisio. 1991. The role of integrins in the maintenance of endothelial monolayer integrity. J. Cell Biol. 112:479. Ridley, A. J., and A. Hall. 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. Daniel, J. L., and J. R. Sellers. 1992. purification and characterisation of platelet myosin. Methods Enzymol. 215:83. Patterson, C. E., H. W. Davis, K. L. Schaphorst, and J. G. N. Garcia. 1994. Mechanisms of cholera toxin prevention of thrombin- and PMA-induced endothelial cell barrier dysfunction. Microvasc. Res. 48:212.
P. multocida TOXIN SIGNALING IN ENDOTHELIAL CELLS
