Pertussis toxin directly activates endothelial cell p42/p44
MAP kinases via a novel signaling pathway
Joe G. N.
Garcia1,
Peiyi
Wang1,
Feng
Liu1,
Marc B.
Hershenson2,
Talaibek
Borbiev1, and
Alexander D.
Verin1
1 Division of Pulmonary and Critical Care Medicine,
Department of Medicine, Johns Hopkins University School of Medicine,
Baltimore, Maryland 21224; and 2 Department of Pediatrics,
University of Chicago School of Medicine, Chicago, Illinois
60637
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ABSTRACT |
Bordetella pertussis
generates a bacterial toxin utilized in signal transduction
investigation because of its ability to ADP ribosylate specific G
proteins. We previously noted that pertussis toxin (PTX) directly
activates endothelial cells, resulting in disruption of monolayer
integrity and intercellular gap formation via a signaling pathway that
involves protein kinase C (PKC). We studied the effect of PTX on the
activity of the 42- and 44-kDa extracellular signal-regulated kinases
(ERK), members of a kinase family known to be activated by PKC. PTX
caused a rapid time-dependent increase in bovine pulmonary artery
endothelial cell ERK activity that was significantly attenuated by
1) pharmacological inhibition of MEK, the upstream ERK
activating kinase, 2) an MEK dominant-negative construct,
and 3) PKC inhibition with bisindolylmaleimide. There was
little evidence for the involvement of either G
-subunits, Ras
GTPases, Raf-1, p60src, or phosphatidylinositol 3'-kinases
in PTX-mediated ERK activation. Both the purified
-oligomer binding
subunit of the PTX holotoxin and a PTX holotoxin mutant genetically
engineered to eliminate intrinsic ADP ribosyltransferase activity
completely reproduced PTX effects on ERK activation, suggesting that
PTX-induced ERK activation involves a novel PKC-dependent signaling
mechanism that is independent of either Ras or Raf-1 activities and
does not require G protein ADP ribosylation.
signal transduction; endothelium; bacterial toxin; adenosine
5'-diphosphate ribosylation; extracellular signal-regulated kinases;
-oligomer; Raf-1 activation; p21 Ras activity
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INTRODUCTION |
PERTUSSIS TOXIN
(PTX) is a product of Bordetella pertussis infection and is
a widely used tool for examination of cellular signaling pathways. The
watershed discovery was that pertussis toxin (PTX) exerted its effect
as a modulatory virulence factor by ADP ribosylation of guanine
nucleotide-binding G proteins, key components of stimulus/coupling
signal transduction. The pertussis holotoxin comprises an
ADP-ribosyltransferase fragment (S1) whose targets include
the
-subunit of Gi and Go subclasses of
heterotrimeric G proteins and a
-oligomer containing several binding
subunits (S2-S6) (24, 40,
41). ADP ribosylation of G
uncouples the G protein from its
receptor in a way that disruption of a signaling pathway by PTX is
presumptive evidence of a G protein-regulated pathway. Direct effects
of PTX on cellular function, i.e., in the absence of agonist/ligand
stimulation, have also been noted in numerous cell systems (7,
27, 36, 46-48, 52). This has generally been perceived as
evidence for tonic regulation by a PTX-sensitive G protein; however,
PTX directly elicits several second messenger cascades capable of
evoking specific biochemical and physiological responses such as
Ca2+ mobilization and cAMP and diacylglycerol synthesis
(46-48, 52). For example, PTX was noted to directly
increase lung weight gain in isolated lung preparations, consistent
with lung cell activation (8, 49), although the exact
mechanism or target of the edemagenic response was not identified. We
previously noted PTX to be a potent direct stimulus for endothelial
paracellular gap formation and increases in macromolecular permeability
across confluent endothelial cell monolayers in vitro (37, 38).
In these studies, neither increases in cytosolic Ca2+ nor
increases in myosin light chain phosphorylation were noted, unique
findings compared with other models of endothelial cell permeability
(15). However, there was strong evidence that PTX-mediated endothelial cell activation was dependent on protein kinase C (PKC)
activity, because PKC inhibition attenuated the extent of PTX-induced
endothelial cell barrier dysfunction (38). The exact PKC
permeability targets responsible for PTX-mediated endothelial cell gap
formation and permeability have not yet been defined; however,
signaling pathways frequently involved in cellular activation such as
phosphatidylinositol-specific phospholipase C or phospholipase D
do not appear to participate in PTX-induced endothelial cell activation (16-18). In fact, our prior results
suggested a novel PKC-dependent model of endothelial cell permeability
that is independent of contractile protein rearrangement driven by a
myosin motor.
The mitogen-activated protein (MAP) kinase family of serine/threonine
protein kinases includes several potential participants in PTX-induced
endothelial cell activation, since several members of this family are
known to be activated by PKC. This MAP kinase family includes three
subgroups [extracellular signal-regulated kinase (ERK), c-Jun
NH2-terminal kinase (JNK), and p38] that are structurally
related, yet exhibit distinct substrate specificity and biological
effects. ERK was the first MAP kinase to be discovered and participates
in cell proliferation, contraction, apoptosis, and a number of
other important cellular responses (19). Increased ERK
activation can follow increases in either p21 Ras GTPase activity or
PKC activity, both of which result in Raf-1 kinase-mediated autophosphorylation and, subsequently, increased activity of the dual-specificity kinase MEK, the direct upstream activator of p42/p44
ERK. The mechanism by which G proteins activate p42/p44 MAP kinases is
poorly understood but has been attributed to 
-subunit involvement
as well as
-subunit-associated coupling (35). In this
study, we have examined whether p42/p44 MAP kinase activity is involved
in endothelial cell activation produced by the important G protein
modulator PTX. Our results indicate PTX to be a robust stimulus for
activation of p42 and p44 ERK1 and ERK2 activation via a novel
signaling pathway that does not involve p21 Ras GTPases, Raf-1, or G protein 
-subunits. In contrast, PKC was critical to
the MEK-dependent, PTX-mediated ERK activation. Both an S1 mutant devoid of ADP ribosyltransferase activity and the purified
-oligomer of the PTX holotoxin directly produced ERK activation. These studies strongly suggest the involvement of ERK signaling pathways in endothelial cell activation evoked by PTX.
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METHODS |
Reagents.
Bovine pulmonary artery endothelial cell cultures were maintained in
DMEM (GIBCO, Chagrin Falls, OH) supplemented with 20% (vol/vol)
colostrum-free bovine serum (Irvine Scientific, Santa Ana, CA), 15 µg/ml endothelial cell growth supplement (Collaborative Research,
Bedford, MA), 1% antibiotic and antimycotic solution (10,000 U/ml
penicillin, 10 µg/ml streptomycin, and 25 µg/ml amphotericin B;
K. C. Biologicals, Lenexa, KS), and 0.1 mM nonessential amino acids (GIBCO). Unless specified, reagents were obtained from Sigma Chemical (St. Louis, MO). Phosphate-buffered saline (PBS) and Hanks'
balanced salt solution without phenol red were purchased from GIBCO
(Grand Island, NY). Polyacrylamide gradient 4-15% ready-to-use gels were purchased from Bio-Rad (Hercules, CA). The MEK inhibitor PD-98059 was purchased from Calbiochem (La Jolla, CA). Endotoxin-free pertussis holotoxin and
-oligomer were purchased from List
Biological Laboratories (Campbell, CA). The S1 mutant,
-adrenergic receptor kinase (
ARK) minigene, hemagglutinin
(HA)-tagged ERK2, and MEK constructs were kindly provided by Drs.
Rappuoli (Sienna, Italy), W. J. Koch (Duke University,
Durham, NC), R. Pestell (Albert Einstein College of Medicine, Bronx,
NY), and M. Rosner (University of Chicago, Chicago, IL), respectively.
Bovine pulmonary artery endothelial cell cultures.
Endothelial cells were obtained from American Type Tissue Culture
Collection (CCL 209; Rockville, MD) at 16 passages, utilized at
passages 19-24, and cultured in complete media
(15, 17). The endothelial cell cultures were maintained at
37°C in a humidified atmosphere of 5% CO2-95% air and
grew to contact-inhibited monolayers with typical cobblestone
morphology. Cells from each primary flask were detached with 0.05%
trypsin, resuspended in fresh culture medium, and passaged into 30- or
60-mm dishes for MAP kinase activity, Raf-1 kinase activity, and p21
Ras activity determination.
ERK activation assays.
Endothelial cell monolayers in 35-mm dishes (100% confluence) were
either serum-starved by incubation with DMEM for 20 h or challenged in complete media and treated with either vehicle or PTX
(Calbiochem, CA) for specified periods of time. The cells were lysed
with 150 µl of boiling lysis buffer containing 10 mM Tris · HCl, pH 7.4, 1% SDS, and 1 mM sodium orthovanadate,
heated to boiling for 5 min, and centrifuged for 5 min. The protein
concentration of the resulting supernatant was determined using BCA
(bicinchoninic acid) Protein Assay Reagent (Pierce). ERK activity of
samples was assessed by either Western blotting with specific
phospho-MAPK (MAP kinase) antibody (New England BioLab, Beverly, MA) or
an in-gel MAP kinase assay. To perform the in-gel MAP kinase assay, we
separated MAP kinases from other proteins by SDS-PAGE (33) with the use of 12.5% polyacrylamide gel containing 0.5 mg/ml myelin
basic protein (MBP) (Sigma). After electrophoresis, the gel was washed
with two changes of 100 ml of 20% isopropanol in 50 mM
Tris · HCl, pH 8.0, for 2 h to remove SDS and then
incubated with 250 ml of buffer A (50 mM Tris · HCl,
pH 8.0, and 5 mM 2-mercaptoethanol) for 1 h with continuous
agitation. To denature the proteins, we incubated the gel with 100 ml
of 6 M guanidine-HCl in buffer A for 1 h with two
exchanges. The proteins in the gel were then renatured by five changes
of buffer A containing 0.04% Tween 20 at 4°C with
continuous agitation for 16 h. To assess phosphorylation of MBP,
we preincubated the gel with 25 ml of kinase buffer [40 mM HEPES-NaOH,
pH 8.0, 2 mM dithiothreitol (DTT), 0.1 mM EGTA, 0.1 mM sodium
orthovanadate, and 10 mM MgCl2] for 30 min at 25°C and
then with 10 ml of kinase buffer containing 25 µM ATP and 50 µCi of
[
-32P]ATP for 1 h with continuous agitation. The
reaction was stopped by washing the gel extensively with 5% TCA
containing 1% sodium pyrophosphate with continuous agitation until all
free radioactivity was liberated. The gel was then dried and exposed to
X-Omat film (Kodak).
Raf-1 activity assay.
Raf-1 kinase activity was assessed by using a commercially available
assay kit (Upstate Biotechnology, Lake Placid, NY). Confluent endothelia were treated with 1 µg/ml PTX or the same volume of PBS as
vehicle control for 5 min after 18 h of serum starvation. As a
positive control, cells were treated with phorbol 12-myristate 13-acetate (PMA; 100 nM) or DMSO as vehicle control (5 min). Cells were
lysed at the end of the incubation period, and Raf-1 kinase was
immunoprecipitated with 4 µg of anti-human c-Raf kinase carboxy terminus at 4°C for 2 h. This was followed by gentle agitation with 100 µl of PBS-prewashed protein G-Sepharose slurry (containing 30% protein G-Sepharose 4 Fast Flow; Amersham Pharmacia Biotech, Piscataway, NJ) for 2 h at 4°C. Immunoprecipitated active Raf was used to phosphorylate and activate GST-MAPKK
(glutathione-S-transferase-MAPK kinase), which in turn
phosphorylates p42 GST-MAPK, resulting in phosphorylation of MBP in the
presence of [
-32P]ATP. The radiolabeled substrates
were allowed to bind to P81 phosphocellulose paper, and the
radioactivity was measured in a scintillation counter.
Cotransfection with plasmids encoding HA-ERK 2 and the
G
-binding domain of
ARK1.
Endothelial cells grown to 80% confluence in 35-mm dishes were
transiently transfected with a plasmid encoding HA-ERK2
(23) and a second plasmid encoding either the G protein

-binding domain of
ARK1 (22, 29) or a
dominant-negative MEK construct (EE-MEK-2E) (57).
The
ARK minigene plasmid (pRK-
BARK1-495-689) contains the carboxy
terminus of the
-adrenergic receptor kinase (the G
-binding
domain) and was kindly provided by Dr. Walter J. Koch (Duke
University). Cells were incubated with 1 µg of total DNA (1:1
ratio of the DNA of the 2 plasmids) and 10 µl of Lipofectamine (GIBCO) in 1 ml of OPTI-MEM for 6 h. The solution was then
replaced with 1 ml of normal growth medium and incubated for 24 h,
and the cells were subsequently serum-starved in DMEM for 20 h.
The transfected endothelial cell monolayers were then treated with either PTX (1 µg/ml) for 5 min or lysophosphatidic acid (LPA; 1 µM)
for 5 min. ERK2 kinase activity was assessed by immunoprecipitation of
HA-tagged ERK2, followed by in vitro phosphorylation assay using MBP as
substrate. Briefly, after treatment with agonists, the cells were
quickly rinsed with PBS and lysed with 150 µl of immunoprecipitation
buffer containing 10 mM Tris · HCl, pH 7.4, 1% Triton X-100,
0.5% Nonidet P-40 (NP-40), 150 mM NaCl, 20 mM NaF, 0.2 mM
sodium orthovanadate, 1 mM EDTA, 1 mM EGTA, and 1% inhibitor cocktails
(Calbiochem) for 30 min at 4°C. The cells were scraped, homogenized
by being passed through a 26-gauge syringe three times, and centrifuged
for 10 min at 4°C. The soluble cell lysate (100 µl), containing
~100 µg of total protein, was incubated with mouse anti-HA antibody
for 1.5 h and then with 15 µl of protein G-Sepharose at 4°C
for 1.5 h. The immune complexes were washed three times with
immunoprecipitation buffer and three times with kinase buffer
containing 10 mM Tris · HCl, pH 7.4, 150 mM NaCl, 10 mM
MgCl2, and 0.5 mM DTT. The immune complexes were
resuspended in 40 µl of kinase buffer with 0.5 mg/ml MBP, 25 µM
ATP, and 2.5 µCi of [
-32P]ATP and incubated at
30°C for 30 min. The reaction was stopped by adding 14 µl of
boiling 4× sample buffer. The samples were then boiled for 5 min and
centrifuged for 5 min, and 15 µl of supernatant were loaded for
SDS-PAGE (33). After electrophoresis, the gel was stained
with Coomassie blue R250, destained, dried, and exposed to X-Omat film (Kodak).
Ras activity assay.
Endothelial cell monolayers were cultured in 35-mm dishes for 7 days in
serum-containing culture medium, serum-starved for 16 h, and
radiolabeled with 220 µCi/ml [32P]orthophosphate in
DMEM for an additional 4 h to label ATP pools. Cells were
challenged with 1 µg of PTX or 100 nM PMA in 1 ml of phosphate-free
DMEM for the indicated times. Medium was then removed, and
cells were lysed in 500 µl of buffer containing 25 mM Tris, pH 7.5, 150 mM NaCl, 16 mM MgCl2, 1% NP-40, 1 mM
phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, and 10 µg/ml
p21ras primary antibody (anti-v-H-ras; Calbiochem). Plates
were incubated on ice for 30 min, and then lysates were scraped from
dishes and centrifuged for 10 min (16,000 g at 4°C). An
additional 2 µg of primary antibody were added to each supernatant
and incubated on ice for 1 h, and 50 µl of lysis buffer
preequilibrated protein G-Sepharose were added to each tube and allowed
to incubate for 1 h with gentle mixing at 4°C. Protein
G-Sepharose was spun down at 80 g for 1 min at 4°C and was
washed four times with washing buffer (lysis buffer without proteinase
inhibitors and antibody). Immunoprecipitates were resuspended in 20 µl of elution buffer containing 2 mM EDTA, 2 mM DTT, and
0.2% SDS and then boiled for 3 min. Sepharose was pelleted by
centrifugation at 16,000 g for 10 min at room temperature.
Supernatants were collected and counted for radioactivity using a
scintillation counter. Equal amounts of radioactivity for each sample
were loaded on 20 × 20-cm thin-layer chromatography (TLC) plates
(Baker-flex cellulose PEI-F; J. T. Baker, Phillipsburg, NJ) and
performed in 0.75 M KH2PO4, pH 3.4. The TLC
plates were exposed to a phosphorimager plate overnight and were
read in a Molecular Dynamics PhosphorImager 445SI. The intensities
of separated [32P]GTP and [32P]GDP were
quantitated, and the data were expressed as the ratio of
[32P]GTP to [32P]GTP and
[32P]GDP.
ADP ribosylation of endothelial cell proteins.
Bacterial toxin-catalyzed ADP ribosylation of proteins contained within
endothelial cell homogenates was measured by incorporation of
32P-labeled NAD (10-20 µCi/ml in ribosylation
cocktail; NEN) as we have previously described (16, 18,
37). PTX (final concentration 1 µg/ml) was preactivated with
20 mM DTT. ADP-ribosylated proteins were separated via SDS-PAGE gels
(33) and detected by autoradiography.
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RESULTS |
PTX induces rapid ERK activation in endothelium.
We initially assessed whether MAP kinases participate in PTX-induced
endothelial cell signal transduction and cellular activation. Figure
1 depicts the rapid increase in p42, p44
ERK activation elicited by PTX as detected by either immunoblotting
with an antibody that only recognizes ERKs phosphorylated at Thr-183
and Tyr-185, a requirement for full enzymatic activity (2,
39) (Fig. 1A) or an in-gel MAP kinase assay using MBP
as substrate (Fig. 1B). PTX-induced ERK activation was
evident in serum-starved endothelium (maximal at 5 min) as well as in
cells challenged in complete medium (maximal at 15 min), with a steady
decline to basal or below basal levels thereafter (Fig. 1C).
Near-maximal stimulation was observed with concentrations as low as 10 ng/ml (Fig. 1D).

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Fig. 1.
Effect of pertussis toxin (PTX) on mitogen-activated
(MAP) kinase activity in bovine pulmonary artery endothelial cell
monolayers. A: confluent bovine endothelium from 35-mm
dishes were serum-starved for 20 h and then challenged with 1 µg/ml PTX for the indicated periods of time. After treatment, cells
were lysed with SDS lysis buffer, scraped off dishes, heated for 5 min,
and microcentrifuged. The supernatant was used for Western
immunoblotting analysis with phospho-specific extracellular
signal-regulated kinase (p-ERK) and pan-ERK antibodies (ab). Plot
represents data averaged from 4 independent experiments quantitated by
scanning densitometry. *P < 0.05 vs. control.
Inset: representative Western blot. PTX significantly
increased ERK activity in a time-dependent manner. B:
similar to results in A, PTX induced maximal ERK activity as
detected by an in-gel assay utilizing myelin basic protein (MBP) as the
in-gel substrate. C: similar to A, confluent
bovine endothelium was challenged with 1 µg/ml PTX but without serum
starvation, and ERK activity was assessed by p-ERK blotting.
D: increasing concentrations of PTX were added to confluent
serum-starved endothelium and then analyzed by Western blotting with
phospho-specific ERK antibodies as described in A.
Concentrations of PTX as low as 10 ng/ml produce near-maximal ERK
activation.
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PTX-mediated ERK activation does not involve ADP ribosylation.
We have previously shown that PTX catalyzes the ADP ribosylation of
40-kDa G proteins in human and bovine endothelial cells that are not
substrates for ADP ribosylation by other bacterial toxins (18,
37). These 40-kDa proteins have previously been shown to
comigrate with a band that is immunoreactive with antibodies directed
against a synthetic peptide corresponding to an amino acid sequence
common to all known G
proteins and migrates to the expected position
of Gi
(18). Prior studies also indicated that maximal PTX-mediated G
ADP ribosylation occurs at 1-2 h (16), a time frame that differs markedly from that of the
maximal PTX-mediated ERK activation (5-15 min) shown in Fig. 1. To
assess the linkage between ADP ribosylation and ERK activation, we next performed experiments to carefully detect the earliest evidence of
PTX-mediated ADP ribosylation in endothelium. Lysates were retrieved
after several defined periods of PTX exposure (1 µg/ml) to allow
endogenous ADP ribosylation, which is assessed by comparison with the
magnitude of the subsequent activated PTX-induced ADP ribosylation assessed in vitro. These studies confirmed that
ADP ribosylation evoked by PTX in intact cells begins after 30 min and
continues to increase up to 120 min (Fig.
2). Thus PTX-induced ADP
ribosyltransferase activity occurs well after the point of maximal ERK
activation (5-15 min), suggesting that G
ADP ribosylation is
unlikely to account for PTX-mediated ERK activation in endothelium. Because of the important implication of these findings, two strategies were next used to further confirm the lack of involvement of G protein
ADP ribosylation in PTX-induced ERK activation in endothelium. One
series of experiments utilized the purified
-oligomer binding subunit, which is the PTX component responsible for cellular binding and facilitating toxin entry into the cell but which is devoid of ADP
ribosyltransferase activity (24). Figure
3A depicts the time-dependent
increase in ERK activation produced by the PTX
-oligomer-binding
subunit over control values, which mirrors the holotoxin effects. The
-oligomer preparation is reported by the manufacturer (List
Biological Laboratories) as potentially containing up to 0.01%
contamination by the PTX holotoxin, resulting in a holotoxin
concentration of <0.01 ng/ml. Although we found that this toxin
concentration does not affect endothelial cell ERK activity (data not
shown), we utilized a second strategy employing a genetically
engineered PTX holotoxin S1 mutant with two site-directed mutations that totally eliminates ADP ribosyltransferase activity (40). Similar to the
-oligomer, the S1 PTX
mutant significantly increased ERK activation in a time-dependent
fashion that closely mimicked the native holotoxin (Fig.
3B). Together, these results strongly indicate that PTX
effects on MAP kinase activity are completely independent of G
ADP
ribosylation and suggest that ligation of a PTX receptor on the cell
surface by the PTX binding subunit is sufficient to activate ERK.

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Fig. 2.
Time course of PTX-mediated ADP ribosylation of
endothelial cell G proteins. Endothelial cell monolayers in 100-mm
dishes were incubated with either vehicle (lanes 1 and
2) or 1 µg/ml PTX (lanes 3-7) for the
indicated periods of time, lysed, and homogenized, followed by
centrifugation at 100,000 g. Pellets were collected, and 60 µg of protein were used for in vitro PTX-catalyzed (1 µg/ml) ADP
ribosylation reaction (except for control, lane 1). The
reactions were carried out in the presence of 5 µCi of
32P-labeled NAD at 30°C for 30 min. Proteins were TCA
precipitated and electrophoresed on 15% SDS-PAGE. The resulting gel
was dried and exposed to Kodak X-OMAT film for 5 h. Lanes
1 and 2 were not pretreated with PTX, and PTX was
omitted from the in vitro reaction mixture that was loaded in
lane 1 but was present in the reaction mixture loaded in
lane 2. Lanes 3-7 represent samples derived
from monolayers pretreated with PTX, which results in diminished
incorporation of 32P during subsequent in vitro
PTX-catalyzed ADP ribosylation. Together, these results indicate that
PTX-induced ADP ribosylation begins after 30 min of PTX challenge, a
time frame that markedly differs from the time course of ERK activation
shown in Fig. 1.
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Fig. 3.
Effect of PTX, -oligomer, and the S1 PTX
mutant on endothelial cell MAP kinase activation. A:
endothelial cell monolayers were treated with 1 µg/ml PTX or 0.734 µg/ml -oligomer (equimolar amount) for 5, 10, and 30 min. The
soluble cell lysates containing comparable total protein were used for
Western blotting with phospho-specific ERK1/ERK2 antibodies.
B: endothelial cell monolayers were treated for 5, 30, or 60 min with PTX (1 µg/ml)or an S1 PTX mutant (1 µg/ml)
that is completely devoid of ADP ribosyltransferase activity. The
soluble cell lysates containing comparable total protein were used for
Western blotting with phospho-specific ERK1/ERK2 antibodies. These data
confirm that ADP ribosyltransferase activity is not required for
PTX-induced endothelial cell ERK activation.
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Independence of ERK activation from Ras and Raf-1 activity.
One well-recognized pathway for ERK induction is via G protein

-subunit activation of Ras GTPases, which increase
Raf-1-mediated phosphorylation of MEK and subsequent ERK activities
(22, 50, 55). Although, in general, these responses have
been noted in response to ligation of specific G protein-coupled
receptors, we next examined whether PTX employs G protein

-subunit interaction with p21 Ras to increase ERK activities.
Endothelial cell monolayers were cotransfected with plasmids encoding
HA-ERK2 and a
ARK minigene encoding a peptide that serves as a
dominant negative for 
-subunit activities (21, 29).
Epitope-tagged ERK was immunoprecipitated, and enzymatic activity was
assessed by in vitro MBP phosphorylation. These studies demonstrated
that 
inhibition does not significantly alter PTX-induced HA-ERK
activity, whereas LPA-stimulated ERK activation was significantly
attenuated by 
inhibition with the
ARK minigene (Fig.
4). Consistent with these results,
measurements of Ras-associated GTP levels were not increased after PTX
(Fig. 5), indicating the absence of Ras
activation, although there was substantial evidence of increased Ras
GTP content after the PKC-activating phorbol ester (PMA) (Fig. 5 and
Table 1). Together, these
studies indicate that PTX-mediated ERK activation does not follow a Ras GTPase-dependent pathway.

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Fig. 4.
Effect of G protein  -subunit inhibition on
PTX-induced ERK activation. Endothelial cells were cotransfected with
plasmids encoding hemagglutinin (HA)-tagged ERK 2 and the
G -binding domain of the -adrenergic receptor kinase
( ARK-1). After transfection, the cells were treated with either
vehicle, PTX (1 µg/ml), or lysophosphatidic acid (LPA; 1 µM). The
cell lysates were immunoprecipitated with HA antibody, and the immune
complexes were used for an in vitro kinase assay utilizing MBP as
substrate. Phosphorylation of MBP was visualized by gel electrophoresis
followed by autoradiography. Unlike LPA, PTX-induced ERK activation
does not involve G protein  subunit-mediated activation.
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Fig. 5.
Effect of PTX on Ras activity. Bovine pulmonary endothelial cells
were incubated with either PTX (1 µg/ml agonist) or phorbol
12-myristate 13-acetate (PMA; 100 nM positive control) for 5 min. The
cell lysates were subjected to immunoprecipitation with 14 µg of
p21ras antibody. Equal amounts of protein were loaded, and
the active GTP-bound Ras and the inactive GDP-bound Ras were separated
by thin-layer chromatography. Ras activity is presented as the ratio of
active Ras to total Ras. Data are means ± SE for 5 independent
experiments. Unlike PMA, PTX does not affect Ras activity.
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ERK activity, in response to growth factor and tumor promoter
stimulation, is known to depend on Src and phosphatidylinositol 3'-kinase (PI 3K) activities (4, 6, 11, 51).
However, our experiments with specific Src and PI 3K inhibitors (PP-2
and LY-294002, respectively) do not support involvement of these
kinases in PTX-induced ERK activation (data not shown). In addition,
phorbol ester- and growth factor-mediated ERK stimulation is dependent on PKC-mediated phosphorylation of Raf-1, a serine/threonine kinase situated upstream to the ERK kinase MEK (5, 12, 30, 31, 45). To explore this potential signal sequence, we pretreated endothelial cell monolayers with the PKC inhibitor bisindolylmaleimide, which significantly reduced PTX-induced ERK activation (Fig.
6). However, kinase activity in Raf-1
immunoprecipitates obtained from PTX-stimulated endothelial cell
monolayers and assessed by quantifying MBP phosphorylation was not
increased compared with PMA (Fig. 7) and
was similar to control values from vehicle-stimulated monolayers,
suggesting that, unlike PMA, PTX-induced ERK activation may proceed via
a Raf-1-independent pathway. Consistent with this notion, treatment of
endothelial cells with forskolin, which decreases ERK activity via
cAMP-mediated inactivation of Raf-1 (10, 20, 56),
decreased the basal level of ERK activity but did not significantly alter PTX-induced ERK activation (data not shown). In contrast to the
lack of Raf-1 involvement, PTX-induced MEK activation appears to be
essential to subsequent increases in ERK activity, because PD-098059, an inhibitor of the upstream ERK-activating dual kinase MEK
(13), abolished PTX-induced ERK activation (Fig.
8A). These studies were
confirmed by manipulating the activity of ERK in vivo by transiently
expressing a dominant-negative MEK construct that is unable to be
phosphorylated by substitution of the regulatory serine phosphorylation
sites with alanine, thereby inhibiting signaling through the ERK
pathway (57). Endothelial cell cotransfection of
a plasmid encoding this MEK mutant with HA-ERK2 established a causal
relationship between PTX-mediated MEK activation and subsequent ERK
activation (Fig. 8B).

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Fig. 6.
Effect of protein kinase C (PKC) inhibition on
PTX-induced ERK activation. Bovine endothelial cells were preincubated
with either vehicle (0.1% DMSO) or the specific PKC inhibitor
bisindolylmaleimide (Bis, 1 µM) for 1 h and then challenged with
PTX (1 µg/ml) for 5 min. ERK activation was assessed by Western
immunoblotting with specific p-ERK antibody. Representative blot
(n = 3) demonstrates that PKC inhibition significantly
attenuates PTX-induced ERK activation in endothelium.
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Fig. 7.
Effect of PTX on Raf-1 activity. Confluent endothelial
cell monolayers were serum-deprived for 18 h and then incubated
with either vehicle (PBS) or PTX (1 µg/ml) for 5 min. As a positive
control, endothelial cells were incubated with 100 nM PMA or 0.1% DMSO
(vehicle) for 5 min. Raf-1 was immunoprecipitated from cell lysates by
an antibody directed against the COOH terminus of human c-Raf kinase.
Raf-1 activity was measured using a Raf-1 kinase cascade assay as
described in METHODS. Data are presented as means ± SE; n = 3. *P < 0.05 vs. basal activity (dotted
line). Whereas phorbol ester-stimulated PKC activation produces a
>5-fold increase in Raf-1 activity, PTX does not increase Raf-1
activity.
|
|

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|
Fig. 8.
Effect of MEK (the MAP kinase kinase) inhibition on
PTX-induced ERK activation. A: confluent endothelial cells
(n = 3) pretreated with either vehicle (0.1% DMSO) or
PD-098059 (50 µM) for 30 min were stimulated with PTX (1 µg/ml) for
5 min. ERK activity was determined as described in METHODS.
Specific inhibition of MEK by PD-098059 significantly attenuated the
basal level of ERK phosphorylation and completely abolished PTX-induced
ERK activation. B: endothelial cells were cotransfected with
HA-ERK (vector) and a construct encoding an MEK dominant-negative
protein (DN MEK). The PTX-stimulated ERK activity in HA-ERK
immunoprecipitates is completely abolished by MEK inhibition
(n = 3).
|
|
 |
DISCUSSION |
Pertussis infection remains a worldwide health problem affecting
target tissues such as the respiratory tract, with many of these
pathological features directly attributed to the holotoxin generated
during this bacterial infection (24, 41). We have previously noted that PTX elicits substantial endothelial cell activation with increases in paracellular gap formation and loss of
semiselective vascular barrier properties (37, 38).
Although the exact signaling pathways by which PTX produces endothelial cell activation were previously elusive, we have now identified a major
role for the 42- and 44-kDa MAP kinases known as ERK. We monitored ERK
activity 1) by immunoprecipitating the ERK kinase and
measuring its activity toward an in vitro substrate, MBP (the most
direct measure of its activity); 2) by Western blotting with antibodies that specifically recognize the phosphorylated and, hence,
activated form of the kinase; and 3) by performing an in-gel assay of MBP phosphorylation. These studies unequivocally demonstrate that PTX is a rapid and potent inducer of ERK activation. We also initiated studies to more precisely define the cellular events evoked
by PTX that lead to ERK activation in endothelium. Because disruption
of signaling pathway by PTX is often used as evidence that ADP
ribosylation of specific heterotrimeric G proteins is involved, our
initial experiments assessed this pathway. As noted before, the
pertussis holotoxin consists of the A protomer, which contains ADP
ribosyltransferase activities (24), whereas the
-oligomer binds the toxin to target cells and increases the
efficiency of noncovalently bound S1 entry and
translocation to target sites (24, 53). We utilized
commercially available
-oligomer subunits as well as a recombinant
S1 mutant to clarify specific mechanisms by which PTX may
increase MAP kinase activities. Interestingly, the S1
mutant and the catalytically inactive
-oligomer of the toxin
completely reproduced the holotoxin effects on ERK activation. In
addition, comparison of the temporal sequences of PTX-mediated MAP
kinase activities and ADP ribosylation argues effectively that PTX
effects on endothelial cell ERK activation are entirely independent of
G protein ADP ribosylation. These data are consistent with limited
reports suggesting that mere binding of the
-oligomer to eukaryotic
cells can alter cellular function independently of ADP ribosylation
(27, 46-48, 53). For example, purified
-oligomer
induces mitogenic stimulation of human T cells (46, 48),
enhances glucose oxidation in adipocytes (53), promotes influx of extracellular Ca2+ (38), and
promotes leukemic cell adhesion (52). Although the exact
identity of these binding sites on endothelium remains unknown,
carbohydrate moieties have been speculated to be crucial components of
-oligomer binding sites (24). Given the recent report
that PTX appears to induce leukemic cell adhesion via integrin receptor
(CD11/CD18) binding (54), it is tempting to speculate that
integrin ligation is directly coupled to signaling pathways that can
initiate MEK activity as well as other signaling cascades in
PTX-challenged endothelium. Further studies are required to evaluate
this stimulus/coupling pathway.
We had initially hypothesized that PTX-induced ERK activation might
follow a well-recognized activation sequence of G protein 
-subunit release after receptor occupancy, followed by Ras- and
Raf-1-dependent pathways leading to the sequential activation of MEK
and ERK. We cotransfected HA-ERK2 with the
ARK minigene encoding a
peptide that functions as a dominant negative for 
-mediated activation sequences. The
ARK minigene totally abolished
receptor-mediated ERK activation elicited by LPA, an important
platelet-derived phospholipid growth factor, but did not attenuate MAP
kinase activity after PTX, suggesting that PTX does not stimulate ERK
via a pathway involving G protein subunit dissociation. Consistent with
the results obtained with the
ARK minigene, the PTX-induced ERK
response appears to be Ras independent, as convincingly demonstrated by direct measurements of Ras activity.
It was not unexpected that PTX-induced ERK activity is dependent on the
activation of the dual-specificity kinase MEK. Growth factors induce
ERK activation when activated Raf kinase phosphorylates two regulatory
serine residues (Ser-218, Ser-222) on MEK1, which facilitates
MEK-mediated phosphorylation of the regulatory threonine (Thr-183) and
tyrosine (Tyr-185) residues on ERK2. ERK phosphorylation at these sites
increases ERK catalytic activity, whereas removal of either phosphate
eliminates this activity (2). PTX-mediated ERK activation
was abolished by the synthetic MEK inhibitor PD-98059 as well as by
expression of a MEK dominant-negative construct. These results are
entirely consistent with the notion that MEK1 participates in the
signaling pathway utilized by PTX to completely produce ERK activation
in cultured bovine endothelium.
The mechanism by which MEK is activated independently of Ras and Raf-1
after PTX challenge is not clear but appears to require PKC involvement
as described in other cell types (9, 25, 32, 34, 42, 43).
Ras- and Raf-1-independent ERK activation has been reported
(44), and although not addressed in our study, our data
appear to suggest a role for B-Raf or other Raf-like molecules
including MEKK2, MEKK3, and the p21-activated kinase (PAK) in the
MEK-dependent endothelial cell ERK activation as noted in
nonoverlapping studies (3, 14). We had previously noted in
bovine endothelium that PTX produces substantial PKC activation and
translocation to the plasma membrane in a temporal sequence compatible
with a role for PKC in PTX-induced endothelial cell barrier dysfunction
(38). The exact mechanism by which PTX accomplishes PKC
activation as well as the exact PKC isoforms involved in this response
is also unclear. In addition to conventional PKC isotypes, PKC
- and
-isoforms have been suggested to participate in MAP kinase
regulation in specific cell systems (9, 34, 42). Recently,
we described ERK activation in response to the PKC-activating phorbol
esters, which proceeded in a Ras- and Raf-1-dependent fashion. While
this certainly suggests PKC isotype-specific activation after PTX and
phorbol esters, further work is needed to fully understand the complex
regulatory mechanism that involves ERK activation.
In summary, we have explored early signaling events involved in
PTX-induced endothelial cell activation and have identified a signaling
cascade involving MEK and PKC in the enhancement of ERK MAP kinase
activities. PTX-induced ERK activation was completely independent of
Ras or Raf-1 activities and does not depend on G
ADP ribosylation.
The physiological importance of PTX-mediated ERK activation in human
disease is unknown but is under study. However, ERK has been noted to
regulate the stability of the endothelial cell-cell junctions and force
development (26, 28), suggesting that PTX may utilize
ERK-modified cytoskeletal targets (1) in a manner relevant
to human lung epithelial or endothelial cell barrier dysfunction. Our
results, which demonstrate rapid ERK activation after cellular
interaction with PTX, provide a provocative and potentially important
mechanism by which PTX may promulgate the inflammatory response to this
bacterial infection, resulting in significant increases in mucosal and
vascular permeability.
 |
ACKNOWLEDGEMENTS |
We gratefully acknowledge the contributions of Drs. Joel Moss
(National Institutes of Health Pulmonary and Critical Care Branch), Rino Rappuoli (Sienna, Italy), W. J. Koch (Duke University,
Durham, NC), R. Pestell (Albert Einstein College of Medicine,
Bronx, New York), and M. Rosner (University of Chicago, Chicago,
Illinois) for generously providing reagents for this study, Lakshmi
Natarajan and Anila Ricks-Cord for superb technical assistance, and
Ellen G. Reather for expert manuscript preparation.
 |
FOOTNOTES |
This work was supported by National Heart, Lung, and Blood Institute
Grants HL-50533 and HL-58064 and by the Dr. David Marine Endowment
(J. G. N. Garcia).
Address for reprint requests and other correspondence: J. G. N. Garcia, Johns Hopkins Asthma and Allergy Center, 5501 Hopkins Bayview Circle, 4B.77, Baltimore, MD 21224-6801 (E-mail:
drgarcia{at}jhmi.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 6 October 2000; accepted in final form 15 December 2000.
 |
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