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of June 15, 2017.
Platelet-Activating Factor Receptor and
Innate Immunity: Uptake of Gram-Positive
Bacterial Cell Wall into Host Cells and
Cell-Specific Pathophysiology
Sophie Fillon, Konstantinos Soulis, Surender Rajasekaran,
Heather Benedict-Hamilton, Jana N. Radin, Carlos J.
Orihuela, Karim C. El Kasmi, Gopal Murti, Deepak Kaushal,
M. Waleed Gaber, Joerg R. Weber, Peter J. Murray and
Elaine I. Tuomanen
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The Journal of Immunology is published twice each month by
The American Association of Immunologists, Inc.,
1451 Rockville Pike, Suite 650, Rockville, MD 20852
Copyright © 2006 by The American Association of
Immunologists All rights reserved.
Print ISSN: 0022-1767 Online ISSN: 1550-6606.
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J Immunol 2006; 177:6182-6191; ;
doi: 10.4049/jimmunol.177.9.6182
http://www.jimmunol.org/content/177/9/6182
The Journal of Immunology
Platelet-Activating Factor Receptor and Innate Immunity:
Uptake of Gram-Positive Bacterial Cell Wall into Host Cells
and Cell-Specific Pathophysiology
Sophie Fillon,1* Konstantinos Soulis,1* Surender Rajasekaran,1† Heather Benedict-Hamilton,†
Jana N. Radin,* Carlos J. Orihuela,* Karim C. El Kasmi,* Gopal Murti,‡ Deepak Kaushal,§
M. Waleed Gaber,¶ Joerg R. Weber,储 Peter J. Murray,* and Elaine I. Tuomanen2*
P
roinflammatory bacterial components are at least partially
responsible for causing the clinical features of sepsis, a
syndrome that causes ⬎100,000 deaths each year in the
United States (1). In the case of Gram-positive infection, a key
bacterial element recognized by the innate immune system is the
cell wall, a complex network of peptidoglycan covalently linked to
teichoic acids, proteins, and lipoproteins (2). Host peptidoglycan
recognition proteins bind pathogen-associated molecular patterns
(PAMPs)3 and degrade the cell wall (3, 4), releasing a wide variety
of smaller proinflammatory fragments. Extracellular Gram-positive
cell wall components, lipoproteins, and lipopeptides interact with
membrane-bound TLRs, particularly TLR2 (5–7), and induce a proinflammatory response. Nod proteins are proposed to function as intracellular surveillance proteins for the recognition of subcomponents of
peptidoglycan (8, 9). Nod2-deficient macrophages respond to TLR
agonists but are refractory to muramyl dipeptide stimulation (10 –14).
How peptidoglycan might reach the cytoplasm is unclear.
Streptococcus pneumoniae, a leading cause of pneumonia, sepsis, and meningitis (15), has served as an important model organ-
*Department of Infectious Diseases, †Division of Critical Care Medicine, ‡Department of Molecular Biotechnology, and §Hartwell Center for Bioinformatics and Biotechnology, St. Jude Children’s Research Hospital, Memphis TN 38105; ¶Department
of Biomedical Engineering, University of Tennessee, Memphis, TN 38163; and 储Department of Neurology, Charite-Universitaetsmedizin, Berlin, Germany
Received for publication April 26, 2006. Accepted for publication July 25, 2006.
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
S.F., K.S., and S.R. contributed equally to this work.
2
Address correspondence and reprint requests to Dr. Elaine I. Tuomanen, Department
of Infectious Diseases, St. Jude Children’s Research Hospital, 332 North Lauderdale
Street, Memphis, TN 38105. E-mail address: [email protected]
3
Abbreviations used in this paper: PAMP, pathogen-associated molecular patterns;
CDP, cytidine diphosphate; DAPI, 4⬘,6-diamidino-3-phenylindole; GPCR, G proteincoupled receptor; LVdevP, left ventricular developed pressure; PAF, platelet-activating factor; PAFr, PAF receptor; PLC, phospholipase C; RT, room temperature; TMB,
tetramethylbenzidine.
Copyright © 2006 by The American Association of Immunologists, Inc.
ism for determining the innate immune response to a Gram-positive bacterial cell wall because not only are the detailed chemical
structures of its cell wall components well characterized (reviewed
by Ref. 2), but these structures have also been linked to specific
pathologies in disease (16 –20). Cell wall fragments are released in
vivo during pneumococcal growth and especially during antibiotic-induced bacterial death and are likely to contribute to inflammation and damage during infection.
For the progression of invasive infection, pneumococci traverse
human cells by binding to the platelet-activating factor (PAF) receptor (PAFr) (21, 22), a widespread, G protein-coupled receptor
(GPCR) that naturally recognizes the phosphorylcholine determinant on the eukaryotic proinflammatory chemokine PAF. Like
PAF, the pneumococcal cell wall displays phosphorylcholine that
binds to the PAFr, initiating bacterial uptake. Recently, uptake via
PAFr has also been shown for several other respiratory pathogens
that display phosphorylcholine on their surfaces (23–25). Ligation
of PAFr induces pleiotropic effector responses that are highly dependent on the cell context, in part due to coupling to a variety of
pathways (26, 27). Two signal transduction cascades that have
been studied extensively are the G protein-coupled phosphorylation of protein kinases, which leads to chemotaxis, and the G protein-independent PAFr interaction with the scaffold protein ␤-arrestin-1, which results in receptor recycling by endocytosis (27–
33). The ␤-arrestin-dependent pathway is responsible for
pneumococcal uptake via PAFr (21, 34), but no direct PAF-like G
protein-stimulating activity has been described for the bacteria
(35). The absence of PAFr modifies the progression of pneumococcal disease in several experimental models (34, 36, 37) while
responses to endotoxin challenge are unchanged (38). We sought
to determine the trafficking and inflammatory consequences of the
interaction of the phosphorylcholine-containing cell wall with the
PAFr at the level of cellular physiology as well as in animal
models.
0022-1767/06/$02.00
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The current model of innate immune recognition of Gram-positive bacteria suggests that the bacterial cell wall interacts with host
recognition proteins such as TLRs and Nod proteins. We describe an additional recognition system mediated by the plateletactivating factor receptor (PAFr) and directed to the pathogen-associated molecular pattern phosphorylcholine that results in the
uptake of bacterial components into host cells. Intravascular choline-containing cell walls bound to endothelial cells and caused
rapid lethality in wild-type, Tlr2ⴚ/ⴚ, and Nod2ⴚ/ⴚ mice but not in Pafrⴚ/ⴚ mice. The cell wall exited the vasculature into the heart
and brain, accumulating within endothelial cells, cardiomyocytes, and neurons in a PAFr-dependent way. Physiological consequences of the cell wall/PAFr interaction were cell specific, being noninflammatory in endothelial cells and neurons but causing
a rapid loss of cardiomyocyte contractility that contributed to death. Thus, PAFr shepherds phosphorylcholine-containing bacterial
components such as the cell wall into host cells from where the response ranges from quiescence to severe pathophysiology. The
Journal of Immunology, 2006, 177: 6182– 6191.
The Journal of Immunology
Materials and Methods
Preparation of bacterial components
Intravital fluorescence microscopy
Forty wild-type vs 17 Pafr⫺/⫺ littermates (34), 8 Tlr2⫺/⫺mice (The Jackson Laboratory), and 4 Nod2⫺/⫺ mice (10) vs 6 wild-type C57BL/6 littermates 8 –10 wk old were maintained in biosafety level 1 and level 2 facilities at the St. Jude Children’s Research Hospital Animal Facility
(Memphis, TN). All experimental procedures were done with mice anesthetized with either inhaled isoflurane (Baxter Healthcare) at 2.5% or MKX
(1 ml of ketamine (Fort Dodge Laboratories) at 100 mg/ml, 5 ml of xylazine (Miles Laboratories) at 100 mg/ml, and 21 ml of PBS). MKX was
administered by i.p. injection at a dose of 50 ␮l per 10 g of body weight.
All experiments were done in compliance with National Institutes of
Health and institutional guidelines.
For cranial window studies, animals were placed in a stereotaxic frame
(Kopf Instruments) and their body temperatures were maintained at ⬃37°C
by a silicon heating mat. As described previously (46), the scalp and underlying tissue over the parietal cortex were removed on one side of the
midsagittal suture using a low-speed dental drill with irrigation, creating a
rectangular cranial window extending from the bregma to the lambdoid
sutures. The dura mater was punctured and excised. Using cyanoacrylate
glue, a 0.5 ⫻ 0.5-cm glass plate was fixed to the bone surrounding the
cranial window. After surgery, mice were allowed to recover for 7–14 days
before data collection was initiated. Intravital microscopy was used to visualize the attachment of a FITC-labeled cell wall to the microvessels of
the brain (46). Mice with cranial windows were anesthetized, immobilized
on a stereotaxic frame, and placed under an industrial scale microscope
(model MM-11; Nikon) with a camera assembly and bright field and fluorescent light sources. Baseline pictures of the cerebral microvasculature
were taken. Mice were injected retro-orbitally or by tail vein with a bolus
of FITC-labeled cell wall equivalent to 5 ⫻ 107 CFU of pneumococci, and
video images and still pictures of the injection, using an excitation wavelength of 490 nm and emission wavelength of 520 nm, were captured using
MetaMorph software (Universal Imaging). Images were captured at 1, 3,
and 5 h after injection. Cell wall fragments attached to the microvasculature were subsequently quantitated (46). For experiments with cytidine
diphosphate (CDP)-choline, mice were given CDP-choline (500 mg/kg;
Biomol) 16 and 1 h before i.v. treatment with 5 ⫻ 107 cell wall equivalents.
For some mice, CDP-choline was given 10 min after cell wall attachment
to the brain vasculature was visualized (n ⫽ 3). For experiments with PAFr
antagonist (CV-6209, 1 mg/kg; Biomol), mice (n ⫽ 6) were treated i.v. at
16 and 1 h before cell wall challenge or received a single dose 10 min after
challenge with 5 ⫻ 107 cell wall equivalents (n ⫽ 3). Six hours after
challenge, animals were sacrificed for histopathology.
To determine more precisely the distribution of cell wall, mice were
sacrificed and the brain and heart were excised and submerged in 10%
buffered formalin (Globe Scientific). The organs were then embedded in
paraffin, cut into 5-␮m-thick sections, and viewed at ⫻100 magnification
using fluorescence or confocal multiphoton microscopy or stained with
H&E for light microscopy.
To follow cell wall distribution from the cerebrospinal fluid, cell wall
(2 ⫻ 107 bacterial equivalents) was injected into the lumbar subarachnoid
space of 4-wk-old C57BL/6 mice as described (n ⫽ 3) (47). Three and 6 h
later, animals were sacrificed, brains were fixed, and 5-␮m sections were
stained using anti-choline TEPC-15 Ab (Sigma-Aldrich) and FITC-labeled
secondary anti-IgA Ab (Sigma-Aldrich) as described previously (4). Distribution of cell wall was determined by fluorescence microscopy.
To view the intracellular distribution of cell wall, deparaffinized brain
sections of animals challenged with FITC-labeled cell wall were blocked
with 1⫻ TBS, 0.5% BSA, 0.5% Triton X-100, and 4% goat serum for 30
min. For brain, neurons were detected using the neuronal marker NeuN
(1/500; Chemicon) in blocking solution overnight at 4°C in a humid chamber. Brain sections were washed twice with TBS for 5 min and incubated
with the secondary Ab, Alexa Fluor 594 goat anti-mouse IgG (1/50; Molecular Probes) for 1 h at RT, rinsed twice in TBS, and stained with 4⬘,6diamidino-3-phenylindole (DAPI; a nuclear stain) and viewed at ⫻100 in
a Zeiss 510 NLO confocal-multiphoton microscope. The FITC label was
viewed in the confocal mode using an argon laser, and the DAPI label was
observed in the multiphoton mode with the infrared laser set at the appropriate wavelength and filter combination for viewing DAPI. The images
were collected as Z series, and maximal projections were derived using the
Zeiss software. Heart sections were deparaffinized and coverslipped with
Vectashield Hard Set mounting medium with DAPI. No staining was done
other than with DAPI nuclear stain.
Langendorff isolated heart
All experiments were performed in compliance with National Institutes of
Health and institutional guidelines according to the method of Gao et al.
(48). Three- to 4-wk-old Sprague Dawley rats (four per group; Harlan
Breeders) were anesthetized i.m. with a combination of ketamine (35 mg/
kg) and xylazine (5 mg/kg). Once the animal was completely unresponsive,
a midsternal incision was made and the heart was excised and placed immediately in cold Krebs-Henseleit solution consisting of 118 mM NaCl,
4.75 mM KCl, 1.19 mM KH2PO4, 1.19 mM MgSO4䡠7H2O, 1.8 mM
CaCl2䡠2H2O, 25 mM NaHCO3, and 11 mM glucose. Within 30 s the hearts
were mounted onto a Langendorff perfusion apparatus (Radnotti Glass
Technology), perfused with Krebs-Henseleit solution at 37°C, and oxygenated with 95% oxygen and 5% carbon dioxide with the pH adjusted to
7.35–7.45 at a constant pressure of 60 mm Hg with flow maintained at 8
cc/min. After the hearts were mounted, an incision was made into the left
atrium and a microtip SPR-524 1.25 F Millar catheter (AD Instruments)
was passed into the left ventricle. The Millar catheter was calibrated before
every experiment. The heart was then suspended in a warm bath and the
end diastolic pressure was adjusted to between 5 and 8 mm Hg. After a
20-min equilibration period the peak systolic pressure and end diastolic
pressure were measured at 5-min intervals. Left ventricular developed pressure (LVdevP) was derived by subtracting left ventricular end diastolic
pressure from left ventricular peak systolic pressure. The hearts were exposed to various interventions after equilibration. Choline (n ⫽ 4) or ethanolamine cell wall (n ⫽ 4) was administered through a side arm just above
the heart. Controls (n ⫽ 4) received saline alone. The rate of infusion was
kept at 1.25% of coronary flow using a programmable WPI 200 infusion
pump (World Precision Instruments) and adjusted to keep the final pneumococcal cell wall concentration of 106 CFU equivalents per milliliter
constant. For some experiments, rats (n ⫽ 4) were treated continuously
with the PAFr antagonist BN52021 (10 ␮M; BIOMOL) beginning 30 min
before cell wall infusion. The amplitude contractility and LVdevP data
were assessed using one-way ANOVA and the Tukey honestly significant
difference test.
To follow the cell wall into cardiac muscle in vivo, 3- to 4-wk-old
Sprague Dawley rats (Harlan; n ⫽ 3) were injected i.v. with 2 ⫻ 108 FITC
cell wall and sacrificed 3 h later. Hearts were excised and placed in 10%
formalin (Globe Scientific) For PAF antagonist studies, rats received 0.1
mg of CV6209 (Biomol) i.v. at 24 and 1 h before administering the cell
wall. For all animals, heart tissue was deparaffinized using EZ-DeWax
(BioGenex) and coverslipped using Vectashield mounting medium with
DAPI (Vector Laboratories). Slides were then viewed at ⫻40 magnification on a Zeiss 510 NLO confocal multiphoton microscope. The FITC label
was viewed in the confocal mode using an argon laser, and the DAPI label
was observed in the multiphoton mode with the infrared laser set at the
appropriate wavelength and filter combination for viewing DAPI. The images were collected as Z series, and maximal projections were derived
using the Zeiss software. The amount of cell wall present was determined
by manually counting the particles present in each image.
Eukaryotic cell culture and transfection
The rat brain capillary endothelial cell line rBCEC6 (49) or A549 lung
epithelial cell line (American Type Culture Collection) was grown to confluence (1.5 ⫻ 108 cells in 150-mm tissue culture plates) in humidified 5%
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S. pneumoniae serotype 4, strain TIGR4 (39), or unencapsulated strain R6
were grown on tryptic soy agar (Difco) supplemented with 3% defibrinated
sheep blood or in defined semisynthetic casein liquid medium supplemented with 0.5% yeast extract. Radiolabeling of S. pneumoniae strain R6
or its cell wall was performed by growth in (C⫹Y) medium with reduced
choline (1 ␮g/ml) and supplemented with 2 ␮Ci/ml [methyl-3H]choline
(Amersham Bioscience) (16, 40). Highly purified cell wall was prepared as
described previously (16). Briefly, bacteria were boiled in SDS, mechanically broken by shaking with acid-washed glass beads, and treated sequentially with DNase, RNase, trypsin, LiCl, EDTA, and acetone. Cell
wall was confirmed to be free of protein by analysis for non-cell wall
amino acids using mass spectrometry. The absence of contaminating endotoxin was confirmed by Limulus test (Associates of Cape Cod). As determined by electron microscopy, the majority of cell wall particles were
⬍30 nm in size, a value within the size range capable of passing nuclear
pores (41, 42). Ethanolamine-containing cell wall was prepared by replacing choline with 20 ␮g/ml ethanolamine (Sigma-Aldrich) in chemically
defined growth medium (43). Peptidoglycan free of teichoic acid was prepared from insoluble cell wall by treatment with 48% hydrofluoric acid
(44). Cell wall was directly labeled with 1 mg/ml FITC (Sigma-Aldrich)
solution in carbonate buffer (pH 9.2) for 1 h at room temperature (RT) in
the dark and washed twice with PBS containing Ca2⫹ and Mg2⫹ (45).
6183
6184
Imaging of intracellular cell wall
All cell lines were seeded on glass chamber slides (Nalge Nunc International), and at 30% confluence the cells were exposed to a 0.5–1 ⫻ 108
CFU/ml equivalent of FITC-labeled cell wall for 2 h. Cells were washed
with PBS, fixed with 4% paraformaldehyde (Electron Microscopy Sciences) in PBS at RT for 20 min, and then rinsed and permeabilized with 1%
Triton X-100 (Sigma-Aldrich) at RT. For some experiments, internalized
FITC-labeled cell wall was detected after staining the eukaryotic cells for
10 min with 50 ␮g/ml propidium iodide (Sigma-Aldrich).
For the colocalization of cell wall with PAFr or the nuclear envelope
marker lamin, samples were blocked with 1% BSA (Sigma-Aldrich) in
PBS at RT followed by goat Ab to mouse PAFr (1/100 or 1/250; Santa
Cruz Biotechnology) or human lamin (1/100; Santa Cruz Biotechnology)
overnight at 4°C. Cells were washed with blocking buffer and incubated
with the secondary Ab Alexa Fluor 594 rabbit anti-goat IgG (1/100 or
1/200; Molecular Probes) at RT. Cells were washed twice with PBS and
mounted in a medium containing the DNA-specific dye TO-PRO-3 (Molecular Probes) to stain the nucleus blue. Primary endothelial cells were
stained with monoclonal anti-CD31 (1/50; Sigma-Aldrich) and the secondary Ab, Alexa Fluor 594 goat anti-mouse IgG (1/200; Molecular Probes).
The samples were examined in a Leica TCS NT/SP confocal laser scanning
microscope equipped with argon (488 nm), krypton (568 nm), and heliumneon (633 nm) lasers. The three lasers permitted the imaging of FITC
(green; emission at 518 nm), Texas Red (red; emission at 570 nm), and
TO-PRO-3 (far red; emission at 661 nm, pseudocolored blue) fluorochromes respectively. The samples were examined with ⫻100 Plan Apochromat, numerical aperture 1.4, oil immersion objective. Scanning was
performed in X, Y, and Z planes at a laser power of 100% on all lasers.
When collecting the Z series, the step size was set at 0.5 ␮m. The three
channel images and an overlay image were recorded using the Leica PowerScan software and rescaled and ␥ corrected with the Adobe Photoshop.
Three-dimensional reconstruction was done using the Leica Power Scan
software. Orthogonal sections of the image stack were obtained using the
sectioning feature of the Leica LCS software. The sectional views in the
XZ and YZ planes enabled unambiguous localization of a given object in
a specific cellular compartment.
Effect of cell wall on myocyte contractility in vitro
HL-1 atrial myocytes (a gift from Dr. W. Claycomb, Louisiana State University, New Orleans, LA) were maintained in Claycomb’s medium (JRH
Biosciences) supplemented with 10% FBS (JRH Biosciences), 2 mM Lglutamine (Invitrogen Life Technologies), and 0.1 mM norepinephrine
(Sigma-Aldrich). Contractility studies were performed with a video edge
motion detector (Crescent Electronics) interfaced to a standard closed circuit television video camera (Javelin Electronics) attached to a Nikon Diaphot microscope (⫻40 magnification; Nikon Ph3 DL objective). Output
was interfaced to a Windograf 980 chart recorder (Gould) and images were
recorded using a video signal recorder (Sony). HL-1 cells are spontaneously contractile when confluent and did not require pacing. Edge detection
of beating was recorded at 60 Hz on myocyte monolayers at RT in Claycomb’s medium supplemented with 10% FBS (JRH Biosciences), 2 mM
L-glutamine (Invitrogen Life Technologies), and 0.1 mM norepinephrine
(Sigma-Aldrich). Cells were allowed to contract for 5 min to obtain a
baseline amplitude and then challenged with choline (n ⫽ 4), ethanolamine
cell wall (n ⫽ 4) (5 ⫻ 107 CFU equivalents per milliliter in 1 ml of
supplemented Claycomb’s medium), or medium alone (n ⫽ 3). Contractility was recorded for 1 h. Cells were exposed to 10 ␮M BN52021 PAFr
antagonist or PLC inhibitor U73122 (10 ␮M; BIOMOL) for 1 h and then
to cell wall, and contractility was recorded for 1 h (n ⫽ 4).
Preparation of cellular extracts
Nuclear and cytosolic fractions of eukaryotic cells were prepared as described previously (53). Briefly, whole cells were lysed by mechanical
disruption through a 25-gauge needle in hypotonic buffer and centrifuged
to pellet the crude nuclei and membrane fraction; the supernatant was harvested as the cytosolic fraction. The pellet was resuspended in high salt
buffer, disrupted mechanically again, and centrifuged at 25,000 ⫻ g to
pellet nuclear debris; the supernatant was harvested as the nuclear extract.
Extracts were measured for radioactivity in their entirety. Protein concentrations were determined using the Bio-Rad protein concentration reagent
as described by the manufacturer. Absence of cross-contamination of cytoplasmic and nuclear fractions was confirmed (data not shown). For some
experiments, whole or fractionated endothelial cell extracts were subjected
to SDS PAGE and analyzed by Western blotting to detect phosphorylated
ERK-1/ERK-2 kinases using the PhosphoPlus p44/42 MAPK (Thr202/
Tyr204) (54).
Cytokine ELISA
Primary neurons, rBCEC6 cells, A549 epithelial cells, or HL-1 cardiomyocytes were seeded in duplicate in 24-well plates at 250,000 cells/well in
1 ml of medium. The next day the medium was changed and 100 ␮l of
supernatant was recovered at 0, 2, 4, 6, and 24 h after stimulation with 5 ⫻
106 or 1 ⫻ 107/ml bacterial equivalents of cell wall or 100 ng/ml LPS from
Escherichia coli serotype O111:B4 (Sigma-Aldrich) or 10 ␮g/ml
Pam3Cys. Supernatant was stored at ⫺20°C until assayed. To determine
cytokines in supernatants, microtiter plates (Maxisorb; Nalge Nunc International) were coated with the respective anti-mouse capture Abs (50 ␮l/
well) against IL-1␤ (4 ␮g/ml) and TNF-␣ (3 ␮g/ml; e-Bioscience) in coating buffer (0.1 M Na2CO3H2O and 0.1 M NaHCO3) overnight at 4°C. The
next day, plates were washed with ELISA washing buffer (1% Tween 20,
1 mM Tris-base, and 154 mM NaCl), blocked with 300 ␮l/well ELISA
buffer (10% FCS in PBS), and incubated for 2 h at RT. After washing, 50
␮l per well of a 1/2 dilution in ELISA-buffer of cell supernatant and duplicate serial 2-fold dilutions of the respective standards (50 ng/ml starting
concentration) were added and incubated overnight at 4°C. Plates were
then washed three times, and biotin-conjugated detection Abs for IL-1 (300
ng/ml; BD Pharmingen) and TNF-␣ (1.6 ␮g/ml, eBioscience) were diluted
in ELISA buffer (50 ␮l/well) and added for 45 min at RT. After washing
the plates three times, streptavidin peroxidase (50 ␮l/well; Sigma-Aldrich)
diluted 1/1000 in ELISA buffer was added for 30 min at RT. Plates were
soaked in washing buffer for 1 min and washed five times, the tetramethylbenzidine peroxidase substrate system (Kirkegaard & Perry Laboratories) was applied at 50 ␮l/well, reactions were stopped with 10% phosphoric acid (50 ␮l/well), and absorbance was measured at 405 nm in an
ELISA reader (Molecular Dynamics). Cytokine concentrations were calculated according to the respective standard curves.
Microarray analysis
The response of rBCEC6 cells and primary rat neurons to cell wall was
interrogated using rat whole genome-spanning 60-mer oligonucleotide microarrays representing ⬎20,000 characterized rat transcripts (Agilent Technologies). Triplicate RNA samples were isolated from three independent
biological experiments exposing rBCEC6 cells and neurons to TNF for 2 h
and then treatment with or without 1 ⫻ 107 CFU/ml choline cell wall for
135 min or 5 ⫻ 106 CFU/ml Lyt4-4 heat-killed bacteria (1 h at 70°C). RNA
was isolated from the monolayer using Qiagen RNeasy Midi kit with oncolumn DNase digestion following the manufacturer’s protocol. RNA samples were labeled in triplicate with Cy-3 (untreated) and Cy-5 (treated)
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CO2 at 37°C. Cells were activated with 10 ng/ml TNF-␣ (Research Diagnostics) for 2 h and then challenged with 1 ⫻ 107 bacteria per milliliter or
cell wall at 5 ⫻ 105 bacterial equivalents per milliliter. For some studies,
radiolabeled cell wall was used and nuclear and cytoplasmic fractions were
analyzed for cell wall uptake (as described below). Both cell lines were
demonstrated to express PAFr and ␤-arrestin (data not shown). Pertussis
toxin (1 ␮g/ml; Sigma-Aldrich) was added for 16 –20 h to define the role
of the Gi protein as described by Zhang et al. (50). To determine cell
survival, cultures were exposed to 5 ⫻ 107 cell equivalents of cell wall for
12, 24, and 48 h and then washed in annexin buffer, stained with FITCannexin V (Roche) and propidium iodide according to the manufacturer’s
instructions, and subjected to FACS analysis with a FACSCalibur flow
cytometer and CellQuest software (BD Biosciences).
Primary neurons from embryonic day 15 embryos of wild-type or
pafr⫺/⫺ mice were isolated as described (51). Primary cerebral microvascular endothelial cells were isolated from 6- to 12-wk-old pafr⫺/⫺ mice or
wild-type littermates according to published methods (52) with the following modifications. Briefly, the cortices were homogenized in PBS, pelleted,
and resuspended in collagenase/dispase (1 mg/ml; Roche) for 30 min at
37°C with stirring. The digest was centrifuged, washed twice with PBS,
resuspended in 30 ml of 15% dextran, and centrifuged at 4500 ⫻ g for 30
min at 4°C. The top layer of myelin was discarded and the pellet was again
resuspended in collagenase/dispase and digested for 30 min at 37°C with
stirring. The digest was pelleted, washed with PBS, resuspended in EGM
endothelial cell growth medium (EGM; Cambrex), and cells were plated on
collagen-coated plates. FITC and radiolabeled cell wall challenges were
performed as for RBCEC6 cells.
COS-1 cells (American Type Culture Collection) were grown to monolayers of 3 ⫻ 106 cells and transfected using the FuGENE kit (Roche) with
0.4 ␮g of expression vector DNA for PAFr-GFP, mutant PAFr D289AGFP, and 0.8 ␮g of DNA for ␤-arrestin-1 and mutant ␤-arrestin V53D (31,
34). Successful transfection with PAFr-GFP plasmid was monitored by
fluorescence microscopy.
PAFr-MEDIATED EFFECTS OF CELL WALL
The Journal of Immunology
6185
monoreactive dyes (Amersham Biosciences) using reverse transcriptionbased indirect labeling (具http://www.hartwellcenter.org/bio_services/fungen/cDNA.php#PROTOCOLS典). Labeled products were hybridized overnight, washed, and scanned using a GenePix 4000B scanner (Axon
Instruments). The text data files were prefiltered to remove unreliable data
and normalized using Spotfire DecisionSite for Functional Genomics version 7.2 (Spotfire) to remove dye bias. Fold change of expression levels
and significance of differential gene expression were calculated using Spotfire. Genes were considered to be differentially expressed based on both the
fold change and the p value of significance as calculated by the pairwise t
test. Raw data are available at the Gene Expression Omnibus repository
(GSE5545). The Ingenuity Pathways (Ingenuity Systems) analysis application was used to visualize and explore the results of microarray analysis.
Results
Cell wall accumulates on the vascular endothelium and kills
mice
Cell wall exits the vasculature and enters nonphagocytic
eukaryotic cells
Examination of in vivo cranial window images revealed that cell
wall was not only attached to the endothelium of the vessel wall
but was also visible outside the vasculature within the brain parenchyma (Fig. 1A). Translocation across cell barriers also occurred when cell wall was instilled directly into the cerebrospinal
fluid from where it accumulated within endothelial cells of the
choroid plexus (Fig. 2A) and within the brain parenchyma contiguous to the ventricles (Fig. 2B). Localization of extravascular cell
wall in brain sections from animals studied by cranial window
showed that cell wall was present within neurons (Fig. 2C) and
cardiomyocytes (Fig. 2D). Thus, cell wall was distributed beyond
FIGURE 1. Imaging of intravascular cell wall trafficking via cranial
windows in wild-type and receptor-deficient mice. A and B, FITC-labeled
cell wall (CW) equivalent to 5 ⫻ 107 CFU was injected i.v. into 8- to
10-wk-old C57BL/6 (wild-type) or Pafr⫺/⫺, Tlr2⫺/⫺, and Nod2⫺/⫺ mice or
their littermate controls. At 1 h after injection, images of the microvasculature were acquired through a cranial window with a fluorescence microscope (excitation at 490 nm and emission at 520 nm) and a magnification
of ⫻40. Fragments outside the vasculature were in areas devoid of blood
flow (full movie is available at 具http://www.stjuderesearch.org/data/sepsis典;
user name is sepsis, and password is _sepsis_). An image taken 1 h postPBS injection is shown as a baseline control. Quantification identified an
average of 208 ⫾ 84 cell wall particles bound per field of vision in wildtype mice (n ⫽ 8) compared with 26 ⫾ 12 particles for Pafr⫺/⫺ mice (n ⫽
5; p ⫽ 0.0009). C, For some experiments, peptidoglycan (PG) or ethanolamine (EA) cell wall was substituted for choline cell wall as indicated.
Survival at 5 h is shown as a percentage (number of animals).
the vascular space and entered nonphagocytic parenchymal cells of
the brain and heart.
Analysis of in vivo tissue sections indicated that cell wall within
neurons and cardiomyocytes localized within the cytoplasm and
the nucleus (Fig. 2, E and F). In contrast to control mice, virtually
no cell wall was found in the cytoplasm or nucleus of neurons of
Pafr⫺/⫺ animals (Fig. 2G) but was present in neurons of Tlr2⫺/⫺
and Nod2⫺/⫺ mice (Fig. 2G). These data suggested that cell wall
could enter the cytoplasm and even the nucleus of nonphagocytic
endothelial cells, neurons, and cardiomyocytes in a PAFr-dependent manner. We next initiated a series of experiments to determine the consequences to the host cell of indwelling Gram-positive cell walls.
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Pneumococci use PAFr to cross from one body compartment to
another, i.e., from lung to blood for the progression of pneumonia
to bacteremia or from blood to brain for the development of meningitis. Indeed, PAFr-deficient mice show improved survival in a
pneumococcal sepsis model (50 vs 0% survival of Pafr⫺/⫺ vs wild
type at day 8). To measure the contribution of the PAFr to host cell
binding and the uptake of cell wall, we chose a whole animal
imaging model to track FITC-labeled cell wall. In this model, the
blood-borne cell wall represents material that would be released
from lysing pneumococci as a result of antibiotic therapy. Proteinfree, teichoic acid-bearing cell wall was inoculated into the blood
stream of wild-type mice. We observed that wild-type mice rapidly
became moribund, with most mice (17 of 19) dying within 2 h of
injection. The distribution of FITC-labeled cell wall in the vascular
compartment was determined by continuously imaging cerebral
vessels through a cranial window. In the first 3 min after injection,
rapid streaming of FITC-labeled cell wall was seen along with
blood inside the vessels. By 1 h, sustained binding of cell wall to
the microvasculature had occurred (Fig. 1A). In contrast to wildtype mice, Pafr⫺/⫺ mice had virtually no cell wall attached to
microvessels up to 5 h after FITC-labeled cell wall injection, and
most of the mice survived (Fig. 1A). Mice deficient in two other
cell wall recognition proteins, TLR2 and Nod2, showed abundant
cell wall margination and low survival rates similar to those of
control mice (Fig. 1B). These data suggested that in this model
PAFr, but not TLR2 or Nod2, played a predominant role in the
trafficking of cell wall and the associated lethality.
We next conducted experiments to document the specific requirement for phosphorylcholine for the binding of bacterial elements to PAFr. Cell wall bearing phosphorylethanolamine in place
of phosphorylcholine failed to bind to cerebral vessels, and all of
the animals survived i.v. challenge (Fig. 1C). Similarly, peptidoglycan free of choline-bearing teichoic acid failed to bind cerebral vessels or induce mortality (Fig. 1C).
6186
PAFr-MEDIATED EFFECTS OF CELL WALL
Uptake of cell wall into host cells in vitro
Effect of cell wall/PAFr interaction on pathophysiology in vitro
and in vivo
More malleable in vitro culture systems of rBCEC6 brain endothelial cells, primary neurons, and A549 lung epithelial cells were
used to analyze the kinetics of cell wall uptake and distribution.
Living or heat-killed bacteria bearing radiolabeled cell wall or purified radiolabeled cell wall were added to cell monolayers, and the
3
H signal was measured in the cytosolic and nuclear fractions at
45-min intervals. Cytoplasmic and nuclear localization of the cell
wall label was detected for living (Fig. 3A) and heat-killed bacteria
(Fig. 3B). Purified cell wall showed the greatest intracellular accumulation, peaking at 135 min (Fig. 3C). Unlabeled wall strongly
competed with the uptake of labeled wall in a dose-dependent way,
supporting specificity of the uptake process (data not shown). Approximately 1% of the total radiolabel added was internalized into
Extracellular cell wall has been described as inducing a variety of
cellular responses, including induction of apoptosis (59 – 61).
However, apoptosis was not induced by the uptake of cell wall into
neurons, A549 epithelial cells, cardiomyocytes, and endothelial
cells, because annexin V staining of both control untreated cells
and cell wall-challenged cells remained below 10% of each cell
population over 48 h ( p not significantly different; data not
shown). In phagocytic cells, extracellular cell wall interactions
with TLR2 activate NF-␬B for cytokine production (62). Similarly, Nod proteins are proposed to form an intracellular equivalent
of the TLR system, recognizing peptidoglycan fragments and leading to the activation of NF-␬B (44, 63, 64). According to this
hypothesis, we should expect to observe the induction of a range
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FIGURE 2. Distribution of cell wall in organs. A, Upon injection into
the cerebrospinal fluid of a mouse, cell wall (2 ⫻ 107 bacterial equivalents)
localized within the cells of the choroid plexus as detected at 3 h after
injection by fluorescent-labeled anti-choline TEPC 15 Ab (original magnification ⫻20; inset shows negative control stained after PBS injection).
B, FITC-labeled cell wall injected as in A was taken up into the ependymal
epithelium and the underlying brain parenchyma. Central black area is the
cerebrospinal fluid ventricle. C and D, Injection of mice i.v. with 5 ⫻ 107
cell wall equivalents. FITC-labeled cell wall localized within neurons
(shown in C at 5 h after injection; red is the NeuN neuronal marker) and
cardiomyocytes (shown in D at 1 h after injection). Original magnification
is ⫻100. E and F, Cell wall (green) localized to the nucleus (blue) by
confocal multiphoton imaging of neuron (E) and cardiomyocyte (F); images taken at 5 h after injection. G, Change in uptake of wall into nuclei of
neurons in receptor deficient animals; image was taken at 1 h after injection
(movies showing localization of cell wall in the nucleus can be found at
具http://www.stjuderesearch.org/data/cellwall/典; user name is cellwall, and
password is _cellwall_). For all panels, distribution of cell wall did not
differ between 1, 3, or 5 h after injection, and images shown are
representative.
the cells, with 0.7% in the nucleus and 0.4% in the cytoplasm.
Neurons accumulated 10-fold less total radiolabel than endothelial
cells, whereas virtually no radiolabel was detected in A549 epithelial cells (data not shown). These data indicated that the uptake
of cell wall varied by cell type and involved both cytoplasmic and
nuclear compartments. The PAFr was essential for uptake, because
Pafr⫺/⫺neurons showed a 60 ⫾ 2% decrease in the accumulation
of radiolabeled cell wall in the cytoplasm as compared with wildtype neurons and a 70 ⫾ 8% decrease for the nucleus (n ⫽ 4).
Consistent with in vivo images and in vitro subcellular fractionation, confocal microscopy of rBCEC6 endothelial cells exposed to
cell wall in vitro demonstrated cell wall in the cytoplasm and also
within the nuclear envelope (Fig. 3D). In ⬎50 rBCEC6 cells examined by orthogonal sectioning (Fig. 3E), 1–5 fluorescent foci
localized to the nucleus with occasional cytoplasmic foci found
adjacent to the nucleus (blue), and these cyan foci were at or near
the nuclear periphery. Similar localization in the nucleus and cytoplasm was observed using primary neurons (data not shown).
Functional PAFr can be detected on the eukaryotic cell surface
and in perinuclear and intranuclear regions (55–57). All internalized cell wall fragments colocalized with PAFr in rBCEC6 cells
(Fig. 4, A–C), cardiomyocytes (Fig. 4D), and primary neurons
(data not shown). To further substantiate the role of the PAFr in
cell wall translocation, endothelial cells isolated from wild-type
and Pafr⫺/⫺ mice were challenged with cell wall in vitro. FITClabeled cell wall was readily detected in wild-type endothelial cells
(Fig. 4E) but only rarely in Pafr⫺/⫺ cells (Fig. 4F). ␤-Arrestin-1 is
required for nuclear localization of GPCR (29, 57) and, consistent
with nuclear colocalization of PAFr and cell wall, functional ␤-arrestin-1 was essential for cell wall uptake in a COS cell transfection model (Fig. 4G). PAFr can activate multiple signaling cascades with and without G protein signaling but, characteristically,
␤-arrestin-mediated signaling through PAFr leads to the activation
of ERK1/2 and p38 MAP kinases without G protein participation
(27, 29, 30, 32, 33, 57, 58). Consistent with this finding, rBCEC6
cells, primary neurons, and HL-1 cardiomyocytes challenged with
cell wall in the presence of the G protein inhibitor pertussis toxin
did not show a decrease in the amount of uptake of cell wall (data
not shown) or a change in phosphorylation of ERK1/2 (Fig. 5A). In
contrast, although virtually no uptake of cell wall was detected,
lung epithelial cells showed strongly increased ERK phosphorylation at 60 and 120 min after cell wall challenge, virtually all of
which was pertussis toxin sensitive. Thus, indwelling cell wall
caused no G protein-mediated ERK phosphorylation in three cell
types, whereas the cell showing no uptake demonstrated strong
phosphorylation. These results point to the PAFr-␤-arrestin-1 pathway as being responsible for cell wall uptake into the cytoplasm
and nucleus of host cells.
The Journal of Immunology
6187
of NF-␬B target genes in cells that have an indwelling cell wall.
We would also anticipate that animals with high levels of cell wall
taken up by the PAFr-dependent process should develop an inflammatory response. To test these hypotheses, we treated rat endothelial cells and neurons with cell wall over time and performed
microarray analyses using rat whole genome-spanning 60-mer oligonucleotide arrays. Surprisingly, even though the cultured cells
contained cell wall throughout the cytoplasm and nucleus, no
widespread changes in gene expression were observed and, specifically, no significant changes were detected in classic NF-␬B
target genes including Tnf-␣, Il-1␤, and I-␬B (raw data are available at the Gene Expression Omnibus site as cited in Materials and
Methods). These data suggested that an indwelling cell wall was
not a strong stimulant of a typical inflammatory response and that
it appeared to have a “neutral” effect on gene expression. These
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FIGURE 3. Quantification of cell
wall uptake into cytoplasm and nucleus. S. pneumoniae bacteria or purified cell wall labeled with [methyl3
H]choline (0.00125 cpm per
bacterium or 0.168 cpm per cell wall
equivalent) were incubated with
monolayers of 1.5 ⫻ 108 rBCEC6
cells for the times indicated. Cytosolic fractions (E) and nuclear extracts
(f) were prepared and radioactivity
was counted and expressed as counts
per minute per 600-cm2 monolayer.
A, Living bacteria R6 (1 ⫻ 107 CFU/
ml). B, Heat-killed bacteria (1 ⫻ 107
CFU/ml). C, Purified cell wall (5 ⫻
105
bacterial
equivalents/ml).
Mean ⫾ SD of 3–5 experiments
each. D, Over 135 min, FITC-labeled
cell wall was distributed sparsely in
the cytoplasm of cultured rBCEC6
cells and accumulated in the nucleus.
Two nuclei are shown with membrane marked by anti-lamin Ab (red).
FITC solution alone or FITC-zymosan failed to show intracellular fluorescence (data not shown). E, To further confirm the intranuclear location
of the fluorescent cell wall foci, the
nucleus was stained with TO-PRO-3
(pseudo-colored blue) and orthogonal
sectioning on the foci was performed
within a 2-␮m area in XZ and YZ
planes. FITC-labeled cell wall component is located in the nucleus (in
crosshairs) as evidenced by the cyan
color (green foci colocalized with
blue nuclear dye). The horizontal
(XZ) section of the image stack of the
nucleus is shown at the bottom and
the vertical section (YZ) on the right
side of the image.
data were further confirmed by measuring IL-1␤ or TNF-␣ in the
supernatants of primary neurons, rBCEC6 cells, A549 cells, or
HL-1 cardiomyocytes at 0, 2, 4, 6, or 24 h after cell wall stimulation. In these experiments, no secretion of these cytokines was
observed (data not shown).
Whereas neurons and endothelial cells remained healthy and
quiescent after cell wall uptake, cardiomyocytes observed to contain indwelling cell wall in vivo showed a dramatic response. HL-1
atrial myocytes challenged with cell wall remained viable but
exhibited a rapid decrease in contractility as measured by edge
detection. The amplitude of contraction of cell wall-challenged
cells was decreased as compared with control cells and with cells
exposed to ethanolamine cell wall at all of the assessed time points
(Fig. 5B). In two of the five experiments the effect was so strong
that no amplitude of contraction could be detected at 60 min, a
6188
phenomenon not observed in any control experiment. Contractility
was preserved in cardiomyocytes pretreated with the PAFr antagonist (Fig. 5B) or phospholipase C (PLC) inhibitor (beat amplitude
of cells challenged with cell wall plus PLC inhibitor remained at
ⱖ98% of PLC inhibitor alone for 60 min; p ⬍ 0.05 by Tukey
comparison of means). These in vitro results indicated that the cell
wall/PAFr interaction in heart was associated with significant
harmful effects on cellular physiology as compared with neurons
or endothelial cells. The significance of these derangements was
further investigated in vivo.
Initiation of antibiotic therapy in the context of sepsis rapidly
releases cell wall fragments in the bloodstream, an effect that is
well known to correlate with transient impairment of cardiac function (65– 67). Such detrimental effects of cell wall uptake on cardiac physiology were demonstrable in the mouse model both in
terms of survival and cardiac dysfunction. Treatment of wild-type
mice with PAFr antagonist CV-6209 for 16 h before cell wall
challenge prevented cell wall attachment to vessels and death (Fig.
6A). Treatment of mice with the antagonist immediately after cell
FIGURE 5. A, Time course of phosphorylation of ERK-1/2 MAPKs in
rBCEC6 endothelial cells, neurons, A549 epithelial cells, and HL-1 cardiomyocytes exposed to 1 ⫻ 107 bacterial equivalents per milliliter of cell
wall (representative of 3– 4 experiments for each cell type). B, The mean
(⫾ SEM) percentage change from baseline in amplitude contractions was
calculated at 15 min intervals for HL-1 mouse atrial myocyte cell cultures
exposed to cell wall (F), cell wall plus the BN52021 PAFr antagonist (‚),
ethanolamine cell wall (), or no treatment (E). ⴱ, Significant difference
from control at p ⬍ 0.05. Experiment was performed four times.
wall administration failed to displace cell wall bound to microvessels (data not shown) or prevent lethality (three of three mice died
within 1 h). The protective benefit of interference with PAFr extended to competitive inhibition of cell wall binding by CDP-choline. Pretreatment with CDP-choline greatly diminished cell wall
binding to the endothelium and increased survival from 10 to 50%
after cell wall challenge (Fig. 6B). Rats treated with PAFr antagonist accumulated significantly less cell wall in cardiomyocytes
(2 ⫾ 2.3 particles per ⫻40 field vs untreated control 6.3 ⫾ 0.4
particles per ⫻40 field). Similarly, rats infused with an ethanolamine cell wall that lacks the choline determinant for binding
PAFr demonstrated significantly less uptake of cell wall into the
heart (0.9 ⫾ 0.2 vs control 6.3 ⫾ 0.4 particles per ⫻40 field).
The Langendorff isolated heart model was used to link the
pathophysiology seen in rats more directly to the effects of cell
wall on cardiac function. The continuous infusion of cell wall was
associated with a decrease in LVdevP in all experiments (Fig. 6B).
The effect was significant at 20 min and remained so throughout
the remainder of the experiment. There was no difference in the
LVdevP in isolated heart preparations receiving ethanolamine cell
wall or the controls (Fig. 6B). In the isolated heart preparations
treated with PAFr antagonist, LVdevP remained at control levels
until 55 min with a modest decrease thereafter (Fig. 6B). The efficacy of the PAFr antagonist and the requirement for choline on
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FIGURE 4. Interaction of cell wall with PAFr. A–C, Colocalization of
FITC-labeled choline cell wall (green in A) and Alexa Fluor 594-labeled
PAF receptor (red in B) in the nucleus (blue in C) as determined by the
merger of all three colors in the overlay image (green plus red plus blue
equals white in C). Controls in which cells were treated with the secondary
Ab alone failed to label the nuclear foci. Orthogonal sections show that the
cell wall fragment and the PAFr are within the nuclear envelope (crosshairs) (representative of ⬎50 cells in two experiments). D, FITC-labeled
cell wall (green) was visualized in cardiomyocytes colocalized with PAFr
(red) as indicated by merged colors (yellow). E, FITC-labeled cell wall was
visualized in the cytoplasm and nucleus (blue) of wild-type primary murine
brain-derived microvascular endothelial cells labeled with CD31 marker
(red). F, Only the rare FITC-labeled cell wall was observed in brain-derived microvascular endothelial cells from Pafr⫺/⫺ mice. G, COS-1 cells
(3– 4 ⫻ 106) were transfected with PAFr (P) and ␤-arrestin-1 (A) or with
combinations of wild-type and nonfunctional mutant receptors as indicated
(mP, mutant PAFr; mA, mutant arrestin). Nontransfected cells (NT) were
used as a negative control (100% ⫽ 177 ⫾ 85 cpm per 600-cm2 monolayer). Cells were challenged with radiolabeled cell wall (5 ⫻ 105 bacterial
equivalents/ml) and uptake was quantitated at 135 min (mean ⫾ SD of
three experiments; ⴱ, p ⱕ 0.019 from t test).
PAFr-MEDIATED EFFECTS OF CELL WALL
The Journal of Immunology
6189
We found that PAFr is a physiologically important innate immune
receptor that recognizes pneumococcal cell wall by virtue of the
PAMP associated with phosphorylcholine-containing bacterial
components. Phosphorylcholine has been detected on a variety of
respiratory pathogens on cell surface components as diverse as
LPS, proteins, or cell wall and is known to direct these bacteria to
the PAFr (21, 23–25). The amount of phosphorylcholine on a bacterial surface is regulated in a phase-variable fashion, indicating
that bacteria devote energy to modulating this interaction in the
context of the infected host (68). A variety of bacterial elements
bearing the PAMP phosphorylcholine become accessible to the
PAFr when released from a pathogen, for instance by cell surface
turnover or antibiotic-induced lysis. We determined that PAFr supports binding of circulating cell wall to endothelia, entry of cell
wall into organs, and, within organs, specific uptake into nonphagocytic cells such as neurons and cardiomyocytes. This egress of
cell wall out of the vascular space indicates that phosphorylcholine-containing material distributes itself widely and comes to reside within organs, an event that correlates with the death of the
animal. Survival is promoted by the absence of PAFr in mice, the
removal of phosphorylcholine from cell wall, or the treatment of
mice with CDP-choline or PAFr antagonists that block cell wall
margination on the vascular endothelium.
A striking feature of cell wall uptake was the progression of
distribution into the nucleus. PAFr is one of several GPCRs that
demonstrates both a cell surface distribution and a perinuclear distribution independent of the binding of ligand (55, 56, 69). The
FIGURE 7. Schematic comparison of the proposed PAFr-cell wall innate recognition system to known cell wall receptors and the host responses
they trigger. Cho, phosphorylcholine; M, N-acetylmuramic acid; G,
N-acetylglucosamine.
FIGURE 6. Amelioration of cell wall-induced pathophysiology by inhibition of PAFr. A, Distribution of i.v. injected FITC-labeled cell wall
(5 ⫻ 107 bacterial equivalents) was viewed at 1 h after injection through a
cranial window in wild-type mice treated with saline and PAFr antagonist
CV6209 at 16 and 1 h before cell wall challenge or CDP-choline at 16 and
1 h before cell wall challenge. Survival is indicated as a percentage of the
total number of mice per treatment group. B, The mean (⫾ SEM) percentage of baseline LVdevP for isolated rat heart preparations exposed to cell
wall (E), ethanolamine cell wall (), pretreatment of rats with BN52021
PAFr antagonist before cell wall challenge (ƒ), or no treatment (F). ⴱ,
Significant difference between challenge with cell wall vs untreated control
(p ⬍ 0.05); ⴱⴱ, significant difference between hearts challenged with cell
wall ⫾ PAFr antagonist (p ⬍ 0.05); ⫹, significant difference between PAFr
antagonist treated and untreated control hearts. Experiment was performed
four times.
the cell wall for bioactivity further indicated that cell wall/PAFr
interactions depressed contractility of the heart.
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Discussion
scaffold protein ␤-arrestin associates with GPCRs to enable their
nuclear distribution (57). Consistent with these findings, we demonstrated that the ␤-arrestin/PAFr association is required for the
localization of cell wall within the nucleus of eukaryotic cells.
Access of cell wall to nuclear material presents the interesting
possibility of novel mechanisms for modulation of gene
expression.
The binding of PAFr by macromolecular cell wall adds a new
dimension to the process of cell wall recognition by nonimmune
cells such as endothelial cells and neurons (Fig. 7). Bacterial peptidoglycan degradation products are proposed to interact with
Nod1 and Nod2 in the cytoplasm of cells leading to cellular activation via NF-␬B (70, 71). In models of Helicobacter infection,
peptidoglycan components have been demonstrated to be injected
into host cells by a bacterial type IV secretion system (70, 71). Our
results show that Gram-positive bacterial cell wall enters not only
the cytoplasm but also the nucleus of nonphagocytic endothelial
cells, cardiomyocytes, and neurons. However, this uptake pathway
differs significantly from that for the peptidoglycan fragments from
Gram-negative bacteria in several ways. First, rather than their
own injection systems, the bacteria use host cell machinery, i.e.,
PAFr and ␤ arrestin. Second, large, teichoicated cell wall components 10 –30 nm in size enter cells. Third, entry extends beyond the
cytoplasm to the nucleus. Fourth, uptake is cell specific, being
prominent in endothelial cells, cardiomyocytes, and neurons, but
not in epithelial cells. Because the Gram-positive cell wall used
here is a fragmented teichoic acid-bearing peptidoglycan, including the muramyl dipeptide subcomponent recognized by Nod2, we
expected to observe a robust inflammatory response in all cell
types carrying an indwelling cell wall. Such data would be in keeping with the concept that Nod proteins recognize cell wall within
the host cell and activate an inflammatory response (63, 64). In
contrast, we observed that most types of host cells were refractory
to significant changes in gene expression, including an absence of
induction of a classic-type NF-␬B response. The lack of induction
of an inflammatory response to indwelling cell wall is consistent
with inhibition of NF-␬B activation in response to some GPCRs by
6190
Acknowledgments
We thank G. Gao, J. Sublett, S. H. Smith-Sielicki, D. Johnson, and P. Austin
for excellent technical assistance and Dr. C. Obert, T. Chin, and
S. Bhuvanendran for methodological advice and equipment. Dr. Robert F.
Tamburro provided excellent statistical advice for the cardiac studies.
Disclosures
The authors have no financial conflict of interest.
References
1. Sprung, C., P. N. Peduzzi, C. Shatney, R. Schein, M. Wilson, J. Sheagren, and
L. Hinshaw. 1990. Impact of encephalopathy on mortality in the sepsis syndrome.
Crit. Care Med. 18: 801– 806.
2. Weber, J., P. Moreillon, and E. Tuomanen. 2003. Innate sensors for Gram positive bacteria. Curr. Opin. Immunol. 15: 408 – 415.
3. Dziarski, R. 2004. Peptidoglycan recognition proteins. Mol. Immunol. 40:
877– 886.
4. Weber, J., D. Freyer, D. Pfeil, C. Kuster, N. Schroeder, E. Tuomanen, and
R. Schumann. 2003. Recognition of pneumococcal peptidoglycan, an expanded
role for lipopolysaccharide binding protein. Immunity 19: 269 –279.
5. Barton, G., and R. Medzhitov. 2002. Toll-like receptors and their ligands. Curr.
Top. Microbiol. Immunol. 270: 81–92.
6. Yoshimura, A., E. Lien, R. Ingalls, E. Tuomanen, R. Dziarski, and D. Golenbock.
1999. Recognition of Gram positive bacterial cell wall components by the innate
immune system occurs via toll-like receptor 2. J. Immunol. 63: 1–5.
7. Aliprantis, A., R. Yang, M. Mark, S. Suggett, B. Devaux, J. Radolf, G. Klimpel,
P. Godowski, and A. Zychlinsky. 1999. Cell activation and apoptosis by bacterial
lipoproteins through toll-like receptor 2. Science 285: 736 –739.
8. Girardin, S., P. Sansonetti, and D. Philpott. 2002. Intracellular vs extracellular
recognition of pathogens — common concepts in mammals and flies. Trends
Microbiol. 10: 193–199.
9. Inohara, N., and G. Nunez. 2003. NODs: intracellular proteins involved in inflammation and apoptosis. Nat. Rev. Immunol. 3: 371–382.
10. Pauleau, A., and P. Murray. 2003. Role of Nod2 in the response of macrophages
to toll like receptor agonists. Mol. Cell. Biol. 23: 7531–7539.
11. Kobayashi, K., M. Chamaillard, Y. Ogura, O. Henegariu, N. Inohara, G. Nunez,
and R. Flavell. 2005. Nod2-dependent regulation of innate and adaptive immunity in the intestinal tract. Science 307: 731–734.
12. Maeda, S., L. Hsu, H. Liu, L. Bankston, M. Iimura, M. Kagnoff, L. Eckmann, and
M. Karin. 2005. Nod2 mutation in Crohn’s disease potentiates NF-␬B activity
and IL-1␤ processing. Science 307: 734 –738.
13. Magalhaes, J., D. Philpott, M. Nahori, M. Jehanno, J. Fritz, L. Bourhis, J. Viala,
J. Hugot, M. Giovannini, J. Bertin, et al. 2005. Murine Nod1 but not its human
orthologue mediates innate immune detection of tracheal cytotoxin. EMBO Rep.
6: 1201–1207.
14. Mariathasan, S., D. Weiss, K. Newton, J. McBride, K. O’Rourke,
M. Roose-Girma, W. Lee, Y. Weinrauch, D. Monack, and V. Dixit. 2006.
Cryopyrin activates the inflammasome in response to toxins and ATP. Nature
440: 228 –232.
15. Schuchat, A., K. Robinson, J. Wenger, L. Harrison, M. Farley, A. Reingold,
L. Lefkowitz, and B. Perkins. 1997. Bacterial meningitis in the United States.
N. Eng. J. Med. 337: 970 –976.
16. Tuomanen, E., H. Liu, B. Hengstler, O. Zak, and A. Tomasz. 1985. The induction
of meningeal inflammation by components of the pneumococcal cell wall. J. Infect. Dis. 151: 859 – 868.
17. Tuomanen, E., R. Rich, and O. Zak. 1987. Induction of pulmonary inflammation
by components of the pneumococcal cell surface. Amer. Rev. Respir. Dis. 135:
869 – 874.
18. Burroughs, M., E. Rozdzinski, S. Geelen, and E. Tuomanen. 1993. A structureactivity relationship for induction of meningeal inflammation by muramyl peptides. J. Clin. Invest. 92: 297–302.
19. Majcherczyk, P., H. Langen, D. Heumann, M. Fountoulakis, M. Glauser, and
P. Moreillon. 1999. Digestion of Streptococcus pneumoniae cell wall with its
major peptidoglycan hydrolase releases branched stem peptides carrying proinflammatory activity. J. Biol. Chem. 274: 12537–12543.
20. Majcherczyk, P., E. Rubli, D. Heumann, M. Glauser, and P. Moreillon. 2003.
Teichoic acids are not required for S. pneumoniae and S. aureus cell walls to
trigger the release of TNF. Infect. Immun. 71: 3707–3713.
21. Cundell, D., N. Gerard, C. Gerard, I. Idanpaan-Heikkila, and E. Tuomanen. 1995.
Streptococcus pneumoniae anchors to activated eukaryotic cells by the receptor
for platelet activating factor. Nature 377: 435– 438.
22. Ring, A., J. Weiser, and E. Tuomanen. 1998. Pneumococcal penetration of the
blood brain barrier: molecular analysis of a novel re-entry path. J. Clin. Invest.
102: 347–360.
23. Weiser, J., M. Shchepetov, and S. Chong. 1997. Decoration of lipopolysaccharide
with phosphorylcholine: a phase-variable characteristic of Haemophilus influenzae. Infect. Immun. 65: 943–950.
24. Weiser, J. B., J. B. Goldberg, N. Pan, L. Wilson, and M. Virji. 1998. The phosphorylcholine epitope undergoes phase variation on a 43-kilodalton protein in
Pseudomonas aeruginosa and on pili of Neisseria meningitidis and Neisseria
gonorrhoeae. Infect. Immun. 66: 4263– 4267.
25. Swords, W., M. Ketterer, J. Shao, C. Campbell, J. Weiser, and M. Apicella. 2001.
Binding of the nontypeable Haemophilus influenzae lipooligosaccharide to the
PAF receptor initiates host cell signalling. Cell Microbiol. 3: 525–536.
26. Zimmerman, G., T. McIntyre, S. Prescott, and D. Stafforini. 2002. The plateletactivating factor signaling system and its regulators in syndromes of inflammation and thrombosis. Crit. Care Med. 30: S294 –S301.
27. Honda, Z., S. Ishii, and T. Shimizu. 2002. Platelet activating factor receptor.
J. Biochem. 131: 773–779.
28. Ihida, K., D. Predescu, R. Czekay, and G. Palade. 1999. Platelet activating factor
receptor is found in a large endosomal compartment in human umbilical vein
endothelial cells. J. Cell Sci. 112: 285–295.
29. Tohgo, A., E. Choy, D. Gesty-Palmer, K. Pierce, S. Laports, R. Oakley,
M. Caron, R. Lefkowitz, and L. Luttrell. 2003. The stability of the G-protein
coupled receptor-␤-arrestin interaction determines the mechanism and functional
consequence of ERK activation. J. Biol. Chem. 278: 6258 – 6267.
30. DeFea, K., J. Zalexsky, M. Thoma, O. Dery, R. Mullins, and N. Bunnett. 2000.
␤-Arrestin-dependent endocytosis of proteinase-activated receptor 2 is required
for intracellular targeting of activated ERK1/2. J. Cell Biol. 148: 1267–1281.
31. Chen, A., D. Dupre, C. Le Gouill, M. Rola-Pleszczynski, and J. Stankova. 2002.
Agonist-induced internalization of platelet-activating factor receptor is dependent
on arrestins but independent of G-protein activation. J. Biol. Chem. 227:
7356 –7362.
32. Kato, M., H. Kimura, Y. Motegi, A. Tachibana, H. Minakami, A. Morkawa, and
H. Kita. 2002. Platelet-activating factor activates two distinct effector pathways in
human eosinophils. J. Immunol. 169: 5252–5259.
33. Deo, D., N. Bazan, and J. Hunt. 2004. Activation of platelet-activating factor
receptor-coupled G␣q leads to stimulation of Src and focal adhesion kinase via
two separate pathways in human umbilical vein endothelial cells. J. Biol. Chem.
279: 3497–3508.
34. Radin, J., C. Orihuela, G. Murti, C. Guglielmo, P. Murray, and E. Tuomanen.
2005. ␤-arrestin 1 determines the traffic pattern of PAFr-mediated endocytosis of
Streptococcus pneumoniae. Infect. Immun. 73: 7827–7835.
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␤-arrestin (72). These findings argue that although cell wall uptake
robustly activates macrophages and dendritic cells, it is neutral in
most nonimmune cell types. Recent findings from microarray and
proteomic experiments using invasive Shigella in epithelial cells
(73) and muramyl dipeptide-stimulated 293T cells overexpressing
Nod2 (74) support the notion that Nod2 does not necessarily activate groups of well-recognized NF-␬B target genes in nonphagocytic cells.
The apparent innocuous nature of cell wall fragments residing in
endothelial cells and neurons left open the question of why systemic cell wall administration leads to the death of mice. In this
regard, we established that cell wall causes PAFr-dependent cardiotoxicity and that this effect is cell specific. It is well known that
different cell types exhibit unique physiological responses to ligation of PAFr based on the activation of distinct signaling pathways
(32, 33, 58). Consistent with this fact, cell wall uptake invoked G
protein-independent signaling in neurons, cardiomyocytes, and endothelial cells, whereas lung epithelial cells demonstrated G protein-dependent signaling and lack of cell wall uptake. The heart is
known to regulate expression of the Pafr transcript in a manner
distinct from that other cell types and, in agreement with our findings concerning the cell wall, PAF induces G protein-independent
signaling in cardiomyocytes (75) that results in the activation of
PLC␥ (33). The ability of a PLC inhibitor to preserve cardiomyocyte contractility suggests this approach could serve as a targeted
therapy to prevent cardiac dysfunction in the context of treatment
of sepsis involving circulating phosphorylcholine-containing bacterial surface components.
PAFr appears to represent an innate PAMP recognition system
for phosphorylcholine that is the first one known to lead to uptake
of Gram-positive cell wall into host cells. It has been generally
believed that cell wall recognition is proinflammatory, but the lack
of response of endothelial cells and neurons to cell wall/PAFr interactions introduces the concept that, in some circumstances, recognition of cell walls is quiescent. This may enhance immune evasion during infection and promote latency or chronicity. Despite
the tolerance of cell wall in some organs, however, it appears that
the heart is particularly susceptible to cell wall-induced pathophysiology and that cardiac dysfunction is a strong contributor to acute
mortality in the context of circulating bacterial debris.
PAFr-MEDIATED EFFECTS OF CELL WALL
The Journal of Immunology
55. Marrache, A., F. Gobiel, S. Bernier, J. Stankova, M. Rola-Pleszczynski,
S. Choufani, G. Bkaily, A. Bourdeau, M. Sirois, A. Vazquez-Tello, et al. 2002.
Proinflammatory gene induction by platelet-activating factor mediated via its
cognate nuclear receptor. J. Immunol. 169: 6474 – 6481.
56. Marrache, A., F. Gobeil, T. Zhu, and S. Chemtob. 2005. Intracellular signaling of
lipid mediators via cognate nuclear G protein-coupled receptors. Endothelium 12:
63–72.
57. Kang, J., Y. Shi, B. Xiang, B. Qu, W. Su, M. Zhu, M. Zhang, G. Bao, F. Wang,
X. Zhang, et al. 2005. A nuclear function of ␤-arrestin 1 in GPCR signaling:
regulation of histone acetylation and gene transcription. Cell 123: 833– 847.
58. Venkatesha, R., J. Ahamed, C. Nuesch, A. Zaidi, and H. Ali. 2004. Plateletactivating factor-induced chemokine gene expression required NF-␬B activation
and Ca2⫹/calcineurin signaling pathways. J. Biol. Chem. 279: 44606 – 44612.
59. Mitchell, L., H. Smith, J. Braun, K. Herzog, J. Weber, and E. Tuomanen. 2004.
Dual phases of apoptosis in pneumococcal meningitis. J. Infect. Dis. 190:
2039 –2046.
60. Bermpohl, B., A. Halle, D. Freyer, E. Dagand, J. Braun, I. Bechman, N. Schroder,
and J. Weber. 2005. Bacterial programmed cell death of cerebral endothelial cells
involves dual death pathways. J. Clin. Invest. 115: 1607–1615.
61. Freyer, D., M. Weih, and J. Weber. 1996. Pneumococcal cell wall components
induce nitric oxide synthase and TNF-␣ in astroglial-enriched cultures. Glia
16: 1– 6.
62. Akira, S., and K. Takeda. 2004. Toll-like receptor signalling. Nat. Rev. Immunol.
4: 499 –511.
63. Girardin, S., and D. Philpott. 2004. Mini-review: the role of peptidoglycan recognition in innate immunity. Eur. J. Immunol. 34: 1777–1782.
64. Sansonetti, P. 2004. War and peace at mucosal surfaces. Nat. Rev. Immunol. 4:
953–964.
65. Ammann, P., T. Fehr, E. Minder, C. Gunter, and O. Bertel. 2001. Elevation of
troponin I in sepsis and septic shock. Intensive Care Med. 27: 965–969.
66. Charan, N., R. Mudumbi, P. Hawk, R. Vestal, and P. Carvalho. 1994. Streptococcus pneumoniae-induced pulmonary hypertension and systemic hypotension
in anesthetized sheep. J. Appl. Physiol. 77: 2071–2078.
67. Horton, J., D. Maass, J. White, and B. Sanders. 2003. Myocardial inflammatory
responses to sepsis complicated by previous burn injury. Surg. Infect. 4: 363–377.
68. Weiser, J., R. Austrian, P. Sreenivasan, and H. Masure. 1994. Phase variation in
pneumococcal opacity: relationship between colonial morphology and nasopharyngeal colonization. Infect. Immun. 62: 2582–2589.
69. Lee, D., A. Lanca, R. Cheng, T. Nguyen, X. Ji, F. Gobeil, Jr., S. Chemtob,
S. George, and B. O’Dowd. 2004. Agonist-independent nuclear localization of
the Apelin, angiotensin AT1, and bradykinin B2 receptors. J. Biol. Chem. 279:
7901–7908.
70. Girardin, S., L. Travassos, M. Herve, D. Blanot, I. Boneca, D. Philpott,
P. Sansonetti, and D. Mengin-Lecreulx. 2003. Peptidoglycan molecular requirements allowing detection by Nod1 and Nod2. J. Biol. Chem. 278: 41702– 41708.
71. Viala, J., C. Chaput, I. Boneca, A. Cardona, S. Girardin, A. Moran, R. Athman,
S. Memet, M. Huerre, A. Coyle, et al. 2004. Nod1 responds to peptidoglycan
delivered by the Helicobacter pylori cag pathogenicity island. Nat. Immunol. 5:
1166 –1174.
72. Gao, H., Y. Sun, Y. Wu, B. Luan, Y. Wang, B. Qu, and G. Pei. 2004. Identification of ␤-arrestin2 as a G protein-coupled receptor-stimulated regulator of
NF-␬B pathways. Mol. Cell 14: 303–317.
73. Tien, M.-T., S. Girardin, B. Regnault, L. Le Bourhis, M.-A. Dillies, J.-Y. Coppee,
R. Bourdet-Sicard, P. Sansonetti, and T. Perdron. 2006. Anti-inflammatory effect
of Lactobacillus casei on Shigella-infected human intestinal epithelial cells.
J. Immunol. 176: 1228 –1237.
74. Weichart, D., J. Gobom, S. Klopfleisch, R. Hasler, N. Gustavsson, S. Billmann,
H. Lehrach, D. Seegert, S. Schreiber, and P. Rosensteil. 2006. Analysis of NOD2meidated proteome response to muramyl dipeptide in HEK293 cells. J. Biol.
Chem. 281: 2380 –2389.
75. Shimizu, T., H. Mutoh, and S. Kato. 1996. Platelet-activating factor receptor.
Gene structure and tissue-specific regulation. Adv. Exp. Med. Biol. 416: 79 – 84.
Downloaded from http://www.jimmunol.org/ by guest on June 15, 2017
35. Cabellos, C., D. E. MacIntyre, M. Forrest, M. Burroughs, S. Prasad, and
E. Tuomanen. 1992. Differing roles of platelet-activating factor during inflammation of the lung and subarachnoid space. J. Clin. Invest. 90: 612– 618.
36. McCullers, J., and J. Rehg. 2005. Lethal synergism between influenza virus and
Streptococcus pneumoniae: characterization of a mouse model and the role of
platelet-activating factor receptor. J. Infect. Dis. 186: 341–350.
37. Rijneveld, A., S. Weijer, S. Florquin, P. Speelman, T. Shimizu, S. Ishii, and
T. van der Poll. 2004. Improved host defense against pneumococcal pneumonia
in platelet-activating factor receptor-deficient mice. J. Infect. Dis. 189: 711–716.
38. Ishii, S., T. Kuawki, T. Nagase, K. Maki, F. Tashiro, S. Sunaga, W. Cao,
K. Kume, Y. Fukuchi, K. Ikuta, et al. 1998. Impaired anaphylactic responses with
intact sensitivity to endotoxin in mice lacking a platelet activating factor receptor.
J. Exp. Med. 187: 1779 –1788.
39. Tettelin, H., K. Nelson, I. Paulsen, J. Eisen, T. Read, S. Peterson, J. Heidelberg,
R. DeBoy, D. Haft, R. Dodson, et al. 2001. Complete genome sequence of virulent isolates of Streptococcus pneumoniae. Science 293: 498 –506.
40. Tuomanen, E. I., A. Tomasz, B. Hengstler, and O. Zak. 1985. The relative role
of bacterial cell wall and capsule in the induction of inflammation in pneumococcal meningitis. J. Infect. Dis. 151: 535–540.
41. Pante, N., and M. Kann. 2002. Nuclear pore complex is able to transport macromolecules with diameters of about 39 nm. Mol. Biol. Cell 13: 425– 434.
42. Feldherr, C., D. Akin, and R. Cohen. 2001. Regulation of functional nuclear pore
size in fibroblasts. J. Cell Sci. 114: 4621– 4627.
43. Tomasz, A., A. Albino, and E. Zanati. 1970. Multiple antibiotic resistance in a
bacterium with suppressed autolytic system. Nature 227: 138 –140.
44. Girardin, S., I. Boneca, J. Viala, M. Chamaillard, A. Labigne, G. Thomas,
D. Philpott, and P. Sansonetti. 2003. Nod2 is a general sensor of peptidoglycan
through muramyl dipeptide (MDP) detection. J. Biol. Chem. 278: 8869 – 8872.
45. Gosink, K., E. Mann, C. Guglielmo, E. Tuomanen, and R. Masure. 2000. Role of
novel choline binding proteins in virulence of Streptococcus pneumoniae. Infect.
Immun. 68: 5690 –5695.
46. Gaber, M., H. Yuan, J. Killmar, M. Naimark, M. Kiani, and T. Merchant. 2004.
An intravital microscopy study of radiation-induced changes in permeability and
leukocyte-endothelial cell interactions in the microvessels of the rat pia mater and
cremaster muscle. Brain Res. Protocol 13: 1–10.
47. Hoffmann, O., N. Keilwerth, M. Bastholm Bille, U. Reuter, K. Angstwurm,
R. Schumann, U. Dirnagl, and J. Weber. 2002. Triptans reduce the inflammatory
response in bacterial meningitis. J. Cereb. Blood Flow Metab. 22: 988 –996.
48. Gao, F., E. Bao, T. Yue, E. Ohlstein, B. Lopez, T. Christopher, and X. Ma. 2002.
Nitric oxide mediates the antiapoptotic effect of insulin in myocardial ischemiareperfusion: the roles of PI3-kinase, Akt, and endothelial nitric oxide synthase
phosphorylation. Circulation 105: 1497–1502.
49. Blasig, I., H. Giese, M. Schroeter, A. Sporbert, D. Utepbergenov, I. Buchwalow,
K. Neubert, G. Schonfelder, D. Freyer, I. Schiimke, et al. 2001. *NO and oxyradical metabolism in new cell lines of rat brain capillary endothelial cells forming
the blood brain barrier. Microvasc. Res. 62: 114 –127.
50. Zhang, Z., H. Cheng, K. Onishi, N. Ohte, T. Wannenburg, and C. Cheng. 2005.
Enhanced inhibition of L-type Ca2⫹ current by ␤3-adrenergic stimulation in
failing rat heart. J. Pharm. Exp. Ther. 315: 1203–1211.
51. Harms, C., M. Lautenshclager, A. Bergk, J. Katchnov, D. Freyer, K. Kapinya,
U. Herwig, D. Megow, U. Dirnagl, and J. Weber. 2001. Differential mechanisms
of neruoprotection by 17 ␤-estradiol in apoptotic versus necrotic neurodegeneration. J. Neurosci. 21: 2600 –2609.
52. Biegel, D., D. Spender, and J. Pachter. 1995. Isolation and culture of human brain
microvessel endothelial cells for the study of blood-brain barrier properties in
vitro. Brain Res. 692: 183–187.
53. Dignam, J. 1983. Accurate transcription initiation by RNA polymerase II in a
soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11:
1475–1489.
54. Schumann, R., D. Pfeil, D. Freyer, W. Buerger, N. Lamping, C. Kirschning,
U. Goebel, and J. Weber. 1998. Lipopolysaccharide and pneumococcal cell wall
components activate the mitogen activated protein kinases (MAPK) ERK-1,
ERK-2, and p38 in astrocytes. Glia 22: 295–305.
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